United States Environmental Protection Agency
Office of Water
Office of Environmental Information
Washington, DC
EPA-841-B-07-009
National Rivers and Streams
Assessment
Field Operations
Manual
April 2009
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National Rivers and Streams Assessment Final Manual
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NOTICE
The intention of the National Rivers and Streams Assessment project is to provide a
comprehensive "State of the Flowing Waters" assessment for rivers and streams across the
United States. The complete documentation of overall project management, design, methods,
and standards is contained in four companion documents:
• National Rivers and Streams Assessment: Quality Assurance Project Plan (EPA-
841-B-07-007)
• National Rivers and Streams Assessment: Site Evaluation Guidelines (EPA-841-B-
07-008)
• National Rivers and Streams Assessment: Field Operations Manual (EPA-841-B-07-
009)
• National Rivers and Streams Assessment: Laboratory Methods Manual (EPA-841-B-
07-010)
This document (Field Operations Manual) contains a brief introduction and procedures to
follow at the base location and on-site, including methods for sampling water chemistry (grabs
and in situ measurements), periphyton, benthic macroinvertebrates, sediment enzymes, fish
composition, fish tissue (at non-wadeable sites), a fecal indicator, and physical habitat. These
methods are based on the guidelines developed and followed in the Western Environmental
Monitoring and Assessment Program (Baker, et al., 1997), the methods outlined in Concepts
and Approaches for the Bioassessment of Non-wadeable Streams and Rivers (Flotemersch, et
al., 2006), and methods employed by several key states that were involved in the planning
phase of this project. Methods described in this document are to be used specifically in work
relating to the National Rivers and Streams Assessment. All Project Cooperators must follow
these guidelines. Mention of trade names or commercial products in this document does not
constitute endorsement or recommendation for use. Details on specific methods for site
evaluation and sample processing can be found in the appropriate companion document.
The citation for this document is:
USEPA. 2007. National Rivers and Streams Assessment: Field Operations Manual.
EPA-841-B-07-009. U.S. Environmental Protection Agency, Washington, DC.
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TABLE OF CONTENTS
TABLE OF CONTENTS v
LIST OF TABLES xi
LIST OF FIGURES xv
ACRONYMS/ABBREVIATIONS xvii
CONTACT LIST xviii
1.0 BACKGROUND 1
1.1 Survey Design 1
1.1.1 Target Population and Sample Frame 2
1.1.2 Replacing Sites 2
1.2 Selection of NRSA Indicators 3
1.3 Description of NRSA Indicators 3
1.4 Supplemental Material to the Field Operations Manual 7
2.0 DAILY OPERATIONS SUMMARY 9
2.1 Sampling Scenario 9
2.1.1 Non-wadeable Sites 9
2.1.2 Wadeable Sites 9
2.2 Recording Data and Other Information 13
2.3 Safety and Health 15
2.3.1 General Considerations 15
2.3.2 Safety Equipment 17
2.3.3 Safety Guidelines for Field Operations 18
3.0 BASE SITE ACTIVITIES 21
3.1 Predeparture Activities 21
3.1.1 Daily Itineraries 22
3.1.2 Instrument Checks and Calibration 22
3.1.3 Equipment and Supply Preparation 22
3.2 Post Sampling Activities 23
3.2.1 Review Data Forms and Labels 23
3.2.2 Inspect and Prepare Samples 23
3.2.3 Equipment Cleanup and Check 24
3.2.4 Supply Inventory 24
3.2.5 Shipment of Samples and Forms 26
3.2.6 Status Reports and Communications 26
4.0 INITIAL SITE PROCEDURES 33
4.1 Site Verification Activities 33
4.1.1 Locating the X-Site 33
4.1.2 Determining the Sampling Status of a Stream 37
4.1.3 Sampling During or After Rain Events 40
4.1.4 Site Photographs 40
4.2 Laying out the sampling reach 40
4.3 Modifying Sample Protocols for High or Low Flows 47
4.3.1 Streams with Interrupted Flow 47
4.3.2 Partially Wadeable Sites 48
4.3.3 Braided Rivers and Streams 48
5.0 NON-WADEABLE RIVERS 49
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5.1 Water Quality 49
5.1.1 In Situ Measurements of Dissolved Oxygen, pH, Temperature, and
Conductivity 49
5.1.1.1 Summary of Method 49
5.1.1.2 Equipment and Supplies 49
5.1.1.3 Multi-Probe Sonde 49
5.1.1.4 Sampling Procedure 51
5.1.2 Water Chemistry Sample Collection and Preservation 52
5.1.2.1 Summary of Method 52
5.1.2.2 Equipment and Supplies 52
5.1.2.3 Sampling Procedure 54
5.1.3 Secchi Disk Transparency at Non-Wadeable Sites 54
5.1.3.1 Summary of Method 54
5.1.3.2 Equipment and Supplies 55
5.1.3.3 Sampling Procedure 55
5.1.4 Sediment Enzymes 56
5.1.4.1 Summary of Method 56
5.1.4.2 Equipment and Supplies 56
5.1.4.3 Sampling Procedure 56
5.2 Physical Habitat Characterization in Non-Wadeable Rivers and Streams 59
5.2.1 Equipment and Supplies 59
5.2.3 Summary of Workflow 62
5.2.4 Habitat Sampling Locations on the Study Reach 63
5.2.5 Work Flow and Reach Marking 64
5.2.6 Reconnaissance 65
5.2.7 Thalweg Profile 65
5.2.7.1 Thalweg Depth Profile 65
5.2.7.2 Pole Drag for Snags and Substrate Characteristics 65
5.2.7.3 Channel Habitat Classification 69
5.2.8 Channel Margin ("Littoral") and Riparian Measurements 70
5.2.8.1 Channel Margin Depth and Substrate 73
5.2.8.2 Large Woody Debris 73
5.2.8.3 Bank Angle and Channel Cross-Section Morphology 75
5.2.8.4 Canopy Cover (Densiometer) 78
5.2.8.5 Riparian Vegetation Structure 79
5.2.8.6 Fish Cover, Algae, Aquatic Macrophytes 80
5.2.8.7 Human Influences 81
5.2.8.8 Riparian "Legacy" Trees and Invasive Alien Species 82
5.2.9 Channel Constraint Assessment 85
5.2.10 Debris Torrents and Recent Major Floods 88
5.3 Periphyton 90
5.3.1 Summary of Method 90
5.3.2 Equipment and Supplies 90
5.3.3 Sampling Procedure 91
5.3.4 Sample Processing in the Field 92
5.4 Benthic Macroinvertebrates 93
5.4.1 Summary of Method 93
5.4.2 Equipment and Supplies 93
5.4.3 Sampling Procedure 94
5.4.4 Sample Processing in Field 94
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5.5 Fish 98
5.5.1 Summary of Method 98
5.5.2 Equipment and Supplies 98
5.5.3 Sampling Procedure 100
5.5.4 Processing Fish 102
5.5.5 Taxonomic Quality Assurance/Quality Control 103
5.6 Fish Tissue 107
5.6.1 Summary of Method 107
5.6.2 Equipment and Supplies 107
5.6.3 Sampling Procedure 110
5.7 Fecal I ndicator (Enterococci) 112
5.7.1 Summary of Method 112
5.7.2 Equipment and Supplies 112
5.7.3 Sampling Procedure 112
6.0 WADEABLE STREAMS 115
6.1 Water Quality 115
6.1.1 In Situ Measurements of Dissolved Oxygen, pH, Temperature, and
Conductivity 115
6.1.1.1 Summary of Method 115
6.1.1.2 Equipment and Supplies 115
6.1.1.3 Multi-Probe Sonde 117
6.1.1.4 Sampling Procedure 117
6.1.2 Water Chemistry Sample Collection and Preservation 118
6.1.2.1 Summary of Method 118
6.1.2.2 Equipment and Supplies 118
6.1.2.3 Sampling Procedure 118
6.1.3 Sediment Enzymes 119
6.1.3.1 Summary of Method 119
6.1.3.2 Equipment and Supplies 119
6.1.3.3 Sampling Procedure 123
6.2 Physical Habitat Characterization—Wadeable Streams 124
6.2.1 Components of the Habitat Characterization 124
6.2.2 Habitat Sampling Locations within the Reach 126
6.2.3 Logistics and Work Flow 126
6.2.4 Thalweg Profile and Large Woody Debris Measurements 128
6.2.4.1 Thalweg Profile 128
6.2.4.2 Large Woody Debris Tally 134
6.2.5 Channel and Riparian Measurements at Cross-Section Transects 134
6.2.5.1 Slope and Bearing 134
6.2.5.2 Substrate Size and Channel Dimensions 141
6.2.5.3 Bank Characteristics 143
6.2.5.5 Riparian Vegetation Structure 151
6.2.5.6 Instream Fish Cover, Algae, and Aquatic Macrophytes 154
6.2.5.7 Human Influence 154
6.2.5.8 Cross-section Transects on Side Channels 155
6.2.5.9 Riparian "Legacy" Trees and Invasive Alien Species 158
6.2.6 Channel Constraint, Debris Torrents, Recent Floods, and
Discharge 159
6.2.6.1 Channel Constraint 159
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6.2.6.2 Debris Torrents and Recent Major Floods 163
6.2.6.3 Stream Discharge 164
6.2.6.4 Velocity-Area Procedure 166
6.2.6.5 Timed Filling Procedure 169
6.2.6.6 Neutrally-Buoyant Object Procedure 170
6.2.7 Equipment and Supplies 172
6.3 Periphyton 173
6.3.1 Summary of Method 173
6.3.2 Equipment and Supplies 173
6.3.3 Sampling Procedure 173
6.3.4 Sample Processing in the Field 174
6.4 Benthic Macroinvertebrates 175
6.4.1 Summary of Method 175
6.4.2 Equipment and Supplies 175
6.4.3 Sampling Procedure 176
6.4.4 Sample Processing in Field 176
6.5 Fish 182
6.5.1 Summary of Method 182
6.5.2 Equipment and Supplies 182
6.5.3 Sampling Procedure 186
6.5.4 Processing Fish 189
6.5.5 Taxonomic Quality Assurance/Quality Control 190
6.6 Fecal Indicator (Enterococci) 196
6.6.1 Summary of Method 196
6.6.2 Equipment and Supplies 196
6.6.3 Sampling Procedure 196
7.0 FINAL SITE ACTIVITIES 198
7.1 General Site Assessment 199
7.1.1 Watershed Activities and Disturbances Observed 199
7.1.2 Site Characteristics 199
7.1.3 General Assessment 199
7.2 Processing the Fecal Indicator, Chlorophyll a, and Periphyton Samples 201
7.2.1 Equipment and Supplies (Fecal Indicator) 201
7.2.2 Procedures for Processing the Fecal Indicator Sample 201
7.2.3 Equipment and Supplies (Chlorophyll a from Water Sample) 203
7.2.4 Procedures for Processing the Chlorophyll a Water Sample 204
7.2.5 Equipment and Supplies (Periphyton Sample) 204
7.2.6 Procedures for Processing the Periphyton Samples 204
7.3 Data Forms and Sample Inspection 208
7.4 Launch Site Cleanup 208
8.0 FIELD QUALITY CONTROL 209
8.1 Repeat and Duplicate Sampling 209
8.1.1 Repeat Sampling 210
8.1.2 Duplicate Sampling 210
8.2 Field Evaluation and Assistance Visits 211
8.2.1 Specifications for QC Assurance 211
8.2.2 Reporting 212
9.0 LITERATURE CITED 213
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APPENDIXA LIST OF EQUIPMENT AND SUPPLIES A-1
APPENDIX B FIELD FORMS B-1
Beatable Forms Packet B-3
Wadeable Forms Packet B-33
APPENDIX C SHIPPING AND TRACKING GUIDELINES C-1
APPENDIX D COMMON & SCIENTIFIC NAMES OF FISHES OF THE UNITED STATES .... D-1
APPENDIX E PPCP and PFC SAMPLES AT SELECTED URBAN SITES E-1
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LIST OF TABLES
Table 1-1. Summary table of indicators for non-wadeable sites 6
Table 1-2. Summary table of indicators for wadeable sites 7
Table 2-1. Guidelines for recording field measurements and tracking information 14
Table 2-2. General health and safety considerations 17
Table 2-3. General safety guidelines for field operations 19
Table 3-1. Stock solutions, uses, and methods for preparation 23
Table 3-2. Postsampling equipment care 25
Table 4-1. Landscape and NHDPIus attributes for the watershed page (data were
summarized from NHDPIus and NLCD2001) 35
Table 4-2. Equipment and supplies list for site verification 37
Table 4-3. Site Verification Procedures 39
Table 4-4. Guidelines to determine the influence of rain events 40
Table 4-5a. Laying out the sampling reach at non-wadeable sites 41
Table 4-5b. Laying out the sampling reach at wadeable sites 43
Table 4-7. Sliding the sampling reach 46
Table 4-8. Reach layout modifications for interrupted streams 47
Table 4-9. Modifications for braided streams 48
Table 5.1-1. Equipment and supplies—DO, pH, temperature, and conductivity 49
Table 5.1-2. Sampling procedure—temperature, pH, conductivity and dissolved oxygen 51
Table 5.1-3. Equipment and supplies—water chemistry sample collection and
preservation 52
Table 5.1-4. Sampling procedure for non-wadeable sites—water chemistry sample
collection 54
Table 5.1 -5. Equipment and supplies—Secchi disc transparency 55
Table 5.1-6. Sampling procedure at non-wadeable sites—Secchi disk transparency 55
Table 5.1 -7. Equipment and supplies—sediment enzymes 56
Table 5.2-1. Checklist of equipment and supplies for physical habitat 59
Table 5.2-2. Components of river physical habitat protocol 61
Table 5.2-4. Thalweg profile procedure 66
Table 5.2-5 Channel unit categories 69
Table 5.2-6. Channel margin depth and substrate procedure 73
Table 5.2-7. Procedure for tallying large woody debris 74
Table 5.2-8. Procedure for bank angle and channel cross-section 75
Table 5.2-9. Procedure for canopy cover measurements 79
Table 5.2-10. Procedure for characterizing riparian vegetation structure 80
Table 5.2.11. Procedure for estimating fish cover 81
Table 5.2-12. Procedure for estimating human influence 82
Table 5.2-13. Procedure for identifying riparian legacy trees and alien invasive species 83
Table 5.2-14. Procedures for assessing channel constraint 85
Table 5.3-1. Equipment and supplies list for periphyton at non-wadeable sites 90
Table 5.3-2. Procedure for collecting composite index samples of periphyton at non-
wadeable sites 92
Table 5.4-1. Equipment and supplies list for benthic macroinvertebrate collection at non-
wadeable sites 93
Table 5.4-2. Procedure for benthic macroinvertebrate sampling at non-wadeable sites 96
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Table 5.4-3. Procedure for compositing samples for benthic macroinvertebrates at non-
wadeable sites 97
Table 5.5-1. Equipment and supplies — fish assessment at non-wadeable sites 98
Table 5.5-2. Procedure for electrofishing at non-wadeable sites 101
Table 5.5-3. Procedure for processing fish at non-wadeable sites 102
Table 5.5-4. Procedure for laboratory identification offish samples 104
Table 5.5-5. Procedure for vouchering of fish samples 106
Table 5.6-1. Equipment and supplies—fish tissue collection at non-wadeable sites 108
Table 5.6-2. Recommended target species for fish tissue collection (in order of
preference) at non-wadeable sites 110
Table 5.6-3. Sampling procedure for fish composite samples at non-wadeable sites 110
Table 5.7-1. Equipment and supplies list for fecal indicator sampling at non-wadeable
sites 112
Table 5.7-2. Procedure for fecal indicator (Enterococci) sample collection at non-
wadeable sites 112
Table 6.1-1. Equipment and supplies—DO, pH, temperature, and conductivity 115
Table 6.1-2. Sampling procedure—temperature, pH, conductivity and dissolved oxygen 118
Table 6.1-3. Equipment and supplies—water chemistry sample collection and
preservation 118
Table 6.1-4. Sampling procedure for wadeable sites—water chemistry sample collection... 119
Table 6.1-5. Equipment and supplies—sediment enzymes 120
Table 6.1 -6. Sampling procedure—sediment enzymes 123
Table 6.2-1. Components of physical habitat characterization 125
Table 6.2-2. Thalweg profile procedure 129
Table 6.2-3. Channel unit and pool forming element categories 133
Table 6.2-4. Procedure for tallying large woody debris 134
Table 6.2-5. Procedure for obtaining slope and bearing data 138
Table 6.2-6. Modified procedure for obtaining slope and bearing data 140
Table 6.2-7. Substrate measurement procedure 142
Table 6.2-8. Procedure for measuring bank characteristics 145
Table 6.2-9. Procedure for canopy cover measurements 150
Table 6.2-10. Procedure for characterizing riparian vegetation structure 153
Table 6.2-11. Procedure for estimating instream fish cover 154
Table 6.2-12. Procedure for estimating human influence 155
Table 6.2-13. Procedure for identifying riparian legacy trees 158
Table 6.2-14. Procedures for assessing channel constraint 161
Table 6.2-15. Velocity-Area procedure for determining stream discharge 167
Table 6.2-16. Timed filling procedure for determining stream discharge 169
Table 6.2-17. Neutrally buoyant object procedure for determining stream discharge 170
Table 6.2-18. Checklist of equipment and supplies for physical habitat 172
Table 6.3-1. Equipment and supplies list for periphyton at wadeable sites 173
Table 6.3-2. Procedure for collecting composite index samples of periphyton at
wadeable sites 173
Table 6.4-1. Equipment and supplies list for benthic macroinvertebrate collection at
wadeable sites 175
Table 6.4-2. Procedure for benthic macroinvertebrate sampling at wadeable sites 179
Table 6.4-3. Procedure for preparing composite samples for benthic macroinvertebrates
at wadeable sites 181
Table 6.5-1. Equipment and supplies — fish assessment at wadeable sites 182
Table 6.5-2. Procedure for electrofishing at wadeable sites <500 m 187
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Table 6.5-3. Procedure for electrofishing atwadeable sites >500 m 188
Table 6.5-4. Procedure for processing fish atwadeable sites 189
Table 6.5-5. Procedure for laboratory identification of fish samples 191
Table 6.5-6. Procedure for vouchering fish samples 192
Table 6.6-1. Equipment and supplies list for fecal indicator sampling atwadeable sites 196
Table 6.6-2. Procedure for fecal indicator (Enterococci) sample collection at wadeable
sites 196
Table 7.1. Equipment and supplies list for fecal indicator sample 201
Table 7.2. Processing procedure—fecal indicator sample 201
Table 7.3. Equipment and supplies list for chlorophyll a processing 203
Table 7.4. Processing procedure—chlorophyll a sample 204
Table 7.5. Equipment and supplies list for periphyton sample processing 204
Table 7.6. Procedure for ID/enumeration samples of periphyton 205
Table 7.7. Procedure for preparing chlorophyll samples of periphyton 205
Table 7.8. Procedure for preparing ash-free dry mass (AFDM) samples of periphyton 206
Table 7.9. Procedure for preparing acid alkaline phosphatase activity samples for
periphyton 208
Table 8.1. General information noted during field evaluation 211
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LIST OF FIGURES
Figure 2-1. Field sampling scenario for non-wadeable sites 11
Figure 2-2. Field sampling scenario for wadeable sites 12
Figure 2-3. Example sample labels for sample tracking and identification 13
Figure 3-1. Overview of base site activities 21
Figure 3-2. Tracking and Sample Status Form 29
Figure 3-3. Tracking (Batched and Retained) Form 30
Figure 3-5. Sample packaging and shipping procedures 32
Figure 4-1. Watershed page 34
Figure 4-2. Site page 36
Figure 4-3. Verification Form (page 1) 38
Figure 4-4. Verification Form (page 2) 44
Figure 4-5. Sampling reach features for a non-wadeable site 45
Figure 5.1-1. Field Measurement Form 50
Figure 5.1-2. Sample Collection Form, Side 1 53
Figure 5.1-3. Secchi disk diagram (EPA, 1991) 55
Figure 5.1-4. Sample Collection Form, Side 2 58
Figure 5.2-1. River reach sample layout 63
Figure 5.2-2. Littoral-Riparian Plots for characterizing riparian vegetation, human
influences, fish cover, littoral substrate, and littoral depths 64
Figure 5.2-3. Thalweg Profile Form 68
Figure 5.2-4. Channel/Riparian Transect Form, page 1 (front side) 71
Figure 5.2-5. Channel/Riparian Transect Form, page 2 (back side) 72
Figure 5.2-8. Field form for Riparian "Legacy" Trees and Invasive Alien Plants (Page 1) 84
Figure 5.2-9. Types of multiple channel patterns 86
Figure 5.4-1. Transect sample design for collecting benthic macroinvertebrates at non-
wadeable sites 94
Figure 5.4-2. Benthic macroinvertebrate collection at non-wadeable sites 95
Figure 5.5-1. Fish Collection Form, Side 1 99
Figure 5.5-2. Transect sampling design for fish sampling at non-wadeable sites 100
Figure 5.6-1. Fish Gear and Voucher/Tissue Sample Information 109
Figure 6.1-1. Field Measurement Form 116
Figure 6.1 -2. Sample Collection Form, Side 1 121
Figure6.1-3. Sample Collection Form, Side2 122
Figure 6.2-1. Reach layout for physical habitat measurements (plan view) 127
Figure 6.2-3. Large woody debris influence zones (modified from Robison and Beschta,
1990) 135
Figure 6.2-4. Channel slope and bearing measurements 137
Figure 6.2-5. Slope and Bearing Form 139
Figure 6.2-6. Substrate sampling cross-section 141
Figure 6.2-7. Channel/Riparian Cross-section Form 144
Figure 6.2-12. Riparian zone and instream fish cover plots for a stream cross-section
transect 152
Figure 6.2-13. Riparian and instream fish cover plots for a stream with minor and major
side channels 156
Figure 6.2-14. Channel/Riparian Cross-section Form for an additional major side channel
transect 157
Figure 6.2-15. Riparian "Legacy" Tree and Invasive Alien Plants Form (Page 1) 160
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Figure 6.2-16. Channel Constraint Form, showing data for channel constraint 162
Figure 6.2-17. Types of multiple channel patterns 163
Figure 6.2-18. Torrent Evidence Assessment Form 165
Figure 6.2-19. Layout of channel cross-section for obtaining discharge data by the
velocity-area procedure 166
Figure 6.2-21. Use of a portable weir in conjunction with a calibrated bucket to obtain an
estimate of stream discharge 171
Figure 6.4-1. Benthic macroinvertebrate collection atwadeable sites 177
Figure 6.4-2. Transect sample design for collecting benthic macroinvertebrates at
wadeable sites 178
Figure 6.5-1. Fish Collection Form for Small Wadeable Streams, Side 1 184
Figure 6.5-2. Fish Collection Form for Large Wadeable Streams (Subreach A-B) 185
Figure 6.5-4. Fish Identification Lab Sheet 195
Figure 7.1. Final site activities summary 198
Figure 7.2. Site Assessment Form 200
Figure 8.1. Summary of the repeat and duplicate sampling design 209
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ACRONYMS/ABBREVIATIONS
AFDM ash-free dry mass
ANC acid neutralizing capacity
APA acid/alkaline phosphatase activity
CPR cardiopulmonary resuscitation
DBH diameter at breast height
Dl deionized
DO dissolved oxygen
DOC dissolved organic carbon
EMAP Environmental Monitoring and Assessment Program
EPA Environmental Protection Agency
ETOH ethyl alcohol
GIS geographic information system
GPS global positioning device
HOPE high density polyethylene
IBI Index of Biotic Integrity
LWD large woody debris
NAD North American Datum
NAWQA National Water-Quality Assessment Program
NHD National Hydrography Dataset
NH4 ammonium
NIST National Institute of Standards
NO3 nitrate
NRSA National Rivers and Streams Assessment
O/E "observed" over "expected"
OSHA Occupational Safety and Health Administration
PFD personal floatation device
P-Hab physical habitat
PSI pounds per square inch
PVC polyvinyl chloride
QAPP Quality Assurance Project Plan
QA/QC quality assurance/quality control
SOPs Standard Operating Procedures
TN total nitrogen
TOC total organic carbon
TP total phosphorus
TSS total suspended solids
USGS United States Geological Survey
WSA Wadeable Streams Assessment
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CONTACT LIST
Information Management Coordinator
Marlys Cappaert
Computer Sciences Corporation
200 S.W. 35th Street
Corvallis, OR 97333
(541) 754-4467
(541)754-4799 fax
cappaert.marlys@epa.gov
Field Logistics Coordinator
Jennifer Pitt
Tetra Tech Center for Ecological Sciences
400 Red Brook Blvd., Suite 200
Owings Mills, MD21117
410-356-8993
410-356-9005 fax
iennifer.pitt@tetratech.com
USEPA HEADQUARTERS
Ellen Tarquinio
USEPA Office of Water
Office of Wetlands, Oceans and Watersheds
1200 Pennsylvania Avenue, NW(4503T)
Washington, D.C. 20460-0001
(202) 566-2267
tarquinio.ellen@epa.gov
Treda Smith
USEPA Office of Water
Office of Wetlands, Oceans and Watersheds
1200 Pennsylvania Avenue, NW(4503T)
Washington DC 20460
202-566-0916
Smith.treda@epamail.epa.gov
USEPA REGIONAL CONTACTS
USEPA Region 1
Tom Faber
USEPA Region 1 - New England Regional
Laboratory
11 Technology Drive
North Chelmsford, MA 01863-2431
(617)918-8672
faber.tom@epa.gov
USEPA Region 2
Darvene Adams
USEPA Facilities
Raritan Depot
2890 Woodbridge Avenue
Edison, NJ 08837-3679
(732) 321-6700
adams.darvene@epa.gov
USEPA Region 3
Louis Reynolds
USEPA Wheeling Operations Office
303 Methodist Building
11th and Chapline Streets
Wheeling, WV 26003
(304) 234-0244
reynolds.louis@epa.gov
USEPA Region 4
Larinda Tervelt
USEPA Region 4
61 Forsyth Street, S.W.
Atlanta, GA 30303-8960
(404) 562-9448
tervelt. larinda@epa.gov
USEPA Region 5
USEPA Region 6
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Sarah Lehmann
USEPA Region 5
77 West Jackson Boulevard
Chicago, IL 60604-3507
(312) 353-4328
lehmann.sarah@epa.gov
USEPA Region 7
Gary Welker
USEPA Region 7
901 North Fifth Street
Kansas City, KS66101
(913)551-7177
welker.gary@epa.gov
Mike Schaub
USEPA Region 6
1445 Ross Avenue
Suite 1200
Dallas, TX 75202-2733
(214)665-7314
schaub.mike@epa.gov
USEPA Region 8
Tina Laidlaw
USEPA Region 8 Montana Office
10 West 15th Street, Suite 3200
Helena, MT 59626
406-457-5016
laidlaw.tina@epa.gov
USEPA Region 9
Janet Hashimoto
USEPA Region 9
75 Hawthorne Street
San Francisco, CA 94105
(415)972-3452
hashimoto.ianet@epa.gov
USEPA Region 10
Gretchen Hayslip
USEPA Region 10
1200 Sixth Avenue
Seattle, WA 98101
(206) 553-1685
hayslip.gretchen@epa.gov
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Page 1
1.0 BACKGROUND
This manual describes field protocols and daily operations for crews to use in the
National Rivers and Streams Assessment (NRSA). The NRSA is a probability-based survey of
our Nation's rivers and streams and is designed to:
• Assess the condition of the Nation's rivers and streams
• Establish a baseline to compare future rivers and streams surveys for trends
assessments
• Evaluate changes in condition from the 2004 Wadeable Streams Assessment
• Help build State and Tribal capacity for monitoring and assessment and promote
collaboration across jurisdictional boundaries
This is one of a series of water assessments being conducted by states, tribes, the U.S.
Environmental Protection Agency (EPA), and other partners. In addition to rivers and streams,
the water assessments will also focus on coastal waters, lakes, and wetlands in a revolving
sequence. The purpose of these assessments is to generate statistically-valid reports on the
condition of our Nation's water resources and identify key stressors to these systems.
The goal of the NRSA is to address two key questions about the quality of the Nation's
rivers and streams:
• What percent of the Nation's rivers and streams are in good, fair, and poor condition
for key indicators of water quality, ecological health, and recreation?
• What is the relative importance of key stressors such as nutrients and pathogens?
The NRSA is designed to be completed during the index period of late May through the
end of September. Field crews will collect a variety of measurements and samples from
predetermined sampling locations (located with an assigned set of coordinates), and from
randomized stations along the sampling reach.
1.1 Survey Design
EPA selected sampling locations using a probability based survey design. Sample
surveys have been used in a variety of fields (e.g., election polls, monthly labor estimates, forest
inventory analysis) to determine the status of populations or resources of interest using a
representative sample of a relatively few members or sites. Using this survey design allows data
from the subset of sampled sites to be applied to the larger target population, and assessments
with known confidence bounds to be made.
The objectives, or design requirements, for the National Rivers and Streams
Assessment are to produce:
• estimates of the 2008-2009 status of all flowing waters nationally and regionally (9
aggregated Omernik ecoregions),
• estimates of the 2008-2009 status of wadeable streams and non-wadeable rivers
nationally and regionally (9 aggregated Omernik ecoregions),
• estimates of the 2008-2009 status of urban flowing waters nationally, and
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• estimates of the change in status in wadeable streams between 2008-2009 and
2004, nationally and regionally (nine aggregated Omernik ecoregions).
With input from the states and other partners, EPA used an unequal probability design to
select 900 wadeable streams and 900 non-wadeable rivers. For purposes of this study, a
wadeable stream segment is defined being >50% wadeable; if it is <50% wadeable, it is defined
as non-wadeable. To evaluate change in wadeable streams from the 2004 WSA, 450 of the 900
wadeable sites were selected using an unequal probability design from the WSA original sites.
The result was the selection of 1800 river and stream sites, with approximately 10%, or 200, of
these sites scheduled for revisits. The NRSA design is explicitly stratified by state. An
"oversample" of additional sites also is available so that any state wishing to conduct a state
scale assessment could be accommodated.
1.1.1 Target Population and Sample Frame
The target population consists of all streams and rivers within the 48 contiguous states
that have flowing water during the study index period excluding portions of tidal rivers up to
head of salt. The study index period extends from late May to the end of September and is
characterized by base flow conditions. The target population includes the Great Rivers. Run-of-
the-river ponds and pools with a residency time of less than 7 days, are included while
reservoirs are excluded. Tidal freshwater rivers and streams are included above the head of
salt. For the purposes of this study the head of salt is < .05ppt. Please refer to the Site
Evaluation Guidelines (EPA-841-B-07-008) and the NRSA Web site
(http://www.epa.gov/owow/riverssurvey/index.html) for more detailed information on the target
population.
The sample frame was derived from the National Hydrography Dataset, NHD-Plus, from
1:100,000 scale maps. Attributes that are used in the NRSA design include:
• State • WSA aggregated ecoregions
• EPA Region • Strahler order (1st through 8th+)
• NAWQA Mega Region • Strahler order categories
• Omernik Ecoregion Level 3 • Urban (site is within "urban" boundary)
1.1.2 Replacing Sites
Sites are organized to be replaced within each state. If a stream or river site is evaluated
and it is determined that it cannot be sampled, then it is to be replaced by another site within the
state. Sites that are coded as 1st through 4th order are to be replaced by oversample sites that
are coded 1st through 4th order, ignoring order within this range. For example, a 2nd order
stream would be replaced by the next 1st, 2nd, 3rd or 4th order stream on the state list. Sites that
are coded as 5th through 10th order are to be replaced by oversample sites that are coded 5th
through 10th order, again ignoring order within the range. For example, a 5th order river would be
replaced by the next 5th, 6th, 7th, 8th, 9th, or 10th order river on the state list. In each case the next
site that is within the Strahler order range is used for the replacement. Please refer to the
Site Evaluation Guidelines (EPA-841-B-07-008) for more detailed information.
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1.2 Selection of NRSA Indicators
As part of the indicator selection process, EPA worked with state and tribal partners and
technical expert consultants through technical conferences and indicator workgroup
teleconferences. The Agency formed a National Rivers and Streams Assessment Steering
Committee with state and regional representatives to develop and refine methodologies. This
section summarizes the Steering Committee recommendations to EPA for selecting NRSA
indicators.
The EPA and partners developed screening and evaluation criteria and identified
potential indicators based on recommendations received at the Large Rivers Assessment
Planning Meeting in San Antonio, Texas (January 10-12, 2007), and the National Rivers and
Streams Planning Session held in Washington, D.C, (April 12, 2007). Key screening and
evaluation criteria included indicator applicability on a national scale, the ability of an indicator to
reflect various aspects of ecological condition, repeatability, and cost-effectiveness.
Participants in indicator discussions included partners and consultants with a technical
background in water monitoring program design and execution, as well as those with knowledge
of state and regional water monitoring programs. Workgroup participants provided feedback on
indicators, field protocols, and analytical procedures for the NRSA. EPA, states, tribes, and
others discussed approaches and options on the chemical, physical, and biological parameters
to be measured. Participants explored the technical and budgetary feasibility of sampling and
analysis methods, the use of specialized technologies (e.g., remote sensing), practical
considerations for completing the assessment (e.g., use of volunteers, availability of labs,
timeframes, funding), and emerging pollutants and contaminant issues.
The remainder of this section briefly describes the indicators that will be used for the
NRSA to assess water quality, ecological integrity, recreational value, and site characteristics
(also see Table 1-1 and Table 1.2).
1.3 Description of NRSA Indicators
In Situ Water Quality Measurements
Measurements for temperature, pH, dissolved oxygen (DO), and conductivity will be
taken with a calibrated water quality probe meter or multi-probe sonde at the X-site (center)
transect in each river or stream. This information will be used to detect extremes in condition
that might indicate impairment.
Secchi Disk Transparency
A Secchi disk is a commonly used black and white patterned disk used to measure the
clarity of water in visibility distance.
Water Chemistry and Associated Measurements
Water chemistry measurements will be used to determine the acidic conditions and
nutrient enrichment, as well as classification of water chemistry type.
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Sediment Enzymes
Benthic organisms are in intimate contact with river sediments, and they are influenced
by the physical and chemical properties of the sediment. Sediment enzyme activity serves as a
functional indicator of key ecosystem processes. Analytical tests include DIN, DIG, TP and TN.
Chlorophyll a
Chlorophyll a is the pigment that makes plants and algae green. Its measurement is
used to determine algal biomass in the water.
Periphyton Assemblage
Periphyton are diatoms and soft-bodied algae that are attached or otherwise associated
with channel substrates. They can contribute to the physical stability of inorganic substrate
particles, and provide habitat and structure. Periphyton are useful indicators of environmental
condition because they respond rapidly and are sensitive to a number of anthropogenic
disturbances, including habitat destruction, contamination by nutrients, metals, herbicides,
hydrocarbons, and acidification.
Benthic Macroinvertebrate Assemblage
Benthic macroinvertebrates are bottom-dwelling animals without backbones
("invertebrates") that are large enough to be seen with the naked eye ("macro"). Examples of
macroinvertebrates include: crayfish, snails, clams, aquatic worms, leeches, and the larval and
nymph stages of many insects, including dragonflies, mosquitoes, and mayflies. Populations in
the benthic assemblage respond to a wide array of stressors in different ways so that it is often
possible to determine the type of stress that has affected a macroinvertebrate assemblage
(Klemm et al., 1990). Because many macroinvertebrates have relatively long life cycles of a
year or more and are relatively immobile, the structure of the macroinvertebrate assemblage is
a response to exposure of present or past conditions.
Fish Assemblage
Monitoring of the fish assemblage is an integral component of many water quality
management programs. The assessment will measure specific attributes of the overall structure
of the ichthyofaunal community to evaluate biological integrity and water quality.
Physical Habitat Assessment
The physical habitat assessment of the sampling reach and the riparian zone (the region
lying along a bank) will serve three purposes. First, habitat information is essential to the
interpretation of what ecological condition is expected to be like in the absence of many types of
anthropogenic impacts. Second, the habitat evaluation is a reproducible, quantified estimate of
habitat condition, serving as a benchmark against which to compare future habitat changes that
might result from anthropogenic activities. Third, the specific selections of habitat information
collected aid in the diagnosis of probable causes of ecological degradation in rivers and
streams. For example, some of the data collected will be used to calculate relative bed stability
(RBS). RBS is an estimate of stream stability that is calculated by comparing the mean
sediment size present to the sediment size predicted by channel and slope.
In addition to information collected in the field by the physical habitat assessment, the
physical habitat description of each site includes many map-derived variables such as stream
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order and drainage area. Furthermore, an array of information, including watershed topography
and land use, supplements the physical habitat information. Together with water chemistry, the
habitat measurements and observations describe the variety of physical and chemical
conditions that are necessary to support biological diversity and foster long-term ecosystem
stability.
Fecal Indicator (Enterococci)
Enterococci are bacteria that are endemic to the guts of warm blooded creatures. These
bacteria, by themselves, are not considered harmful to humans but often occur in the presence
of potential human pathogens (the definition of an indicator organism). Epidemiological studies
of marine and fresh water bathing beaches have established a direct relationship between the
density of enterococci in water and the occurrence of swimming-associated gastroenteritis.
Fish Tissue
The fish tissue contaminants indicator, which measures bioaccumulation of persistent
toxics, is used to estimate national risks offish consumption to humans. Various studies have
been done on fish tissue contaminants focusing on different parts of the fish (e.g., whole fish,
fillets, livers). The NRSA will focus on fillets because of its emphasis on human health.
Other Indicators / Site Characteristics
Pharmaceuticals and Personal Care Products (PPCP) will be sampled from fish tissue
and water column at 154 pre-selected sites. These sites are defined as urban, beatable sites
and will have an additional water grab taken to look at these emerging contaminants.
Observations and impressions about the site and its surrounding catchment by field teams will
be useful for ecological value assessment, development of associations and stressor indicators,
and data verification and validation.
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Table 1 -1. Summary table of indicators for non-wadeable sites
Indicator
In Situ measurements (pH, DO,
temperature, conductivity)
Water chemistry (TP, TN [NH4, NO3),
basic anions and cations, alkalinity
[ANC], DOC, TOC, TSS, conductivity
Secchi Disk transparency
Chlorophyll a
Sediment enzymes
Periphyton
Benthic macroinvertebrate assemblage
(Littoral)
Fish Assemblage
Physical habitat assessment
Fecal indicator (enterococci)
Fish Tissue
Drainage area
Characteristics of watershed
PPCP (Only at pre-defined urban sites)
Specs/Location in Sampling Reach
Measurements taken at X site at midchannel; readings are
taken at 0.5 m depth
Collected from a depth of 0.5 m at the cross site at the center
of the stream
Measured at X site at midchannel
Collected as part of water chemistry and periphyton samples
Collected from 1 1 locations systematically placed at each site
and combined into a single composite sample
Collected from 1 1 locations systematically placed at each site
and combined into a single composite sample
Collected from 1 1 locations systematically placed at each site
and combined into a single composite sample
Sampled throughout the sampling reach at specified
locations
Measurements collected throughout the sampling reach at
specified locations
Collected at the last transect one meter off the bank
Target species collected throughout the sampling reach as
part offish assemblage sampling
Done at desktop, and used in target population selection
Done at desktop using CIS and verified by state agencies
Collected only at specified sites at the X site
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Table 1 -2. Summary table of indicators for wadeable sites
Indicator
In Situ measurements (pH, DO,
temperature, conductivity)
Water chemistry (TP, TN [NH4, NO3),
basic anions and cations, alkalinity
[ANC], DOC, TOC, TSS, conductivity
Chlorophyll a
Sediment enzymes
Periphyton
Benthic macroinvertebrate assemblage
(Littoral)
Fish Assemblage
Physical habitat assessment
Fecal indicator (enterococci)
Drainage area
Characteristics of watershed
Specs/Location in Sampling Reach
One set of measurements taken at the X site in the center of the
stream; readings are taken at 0.5 m depth
Collected from a depth of 0.5 m at the X site at the center of the
stream
Collected as part of water chemistry and periphyton samples
Collected from 1 1 locations systematically placed at each site and
combined into a single composite sample
Collected from 1 1 locations systematically placed at each site and
combined into a single composite sample
Collected from 1 1 locations systematically placed at each site and
combined into a single composite sample
Sampled throughout the sampling reach at specified locations
Measurements collected throughout the sampling reach at
specified locations
Collected at the last transect one meter off the bank
Done at desktop, and used in target population selection
Done at desktop using CIS and verified by state agencies
1.4 Supplemental Material to the Field Operations Manual
The Field Operations Manual describes field protocols and daily operations for crews to
use in the NRSA. Following these detailed protocols will ensure consistency across regions and
reproducibility for future assessments. Before beginning sampling at a site, crews should
prepare a packet for each site containing pertinent information to successfully conduct
sampling. This includes a road map and set of directions to the site, topographic maps, land
owner access forms, sampling permits (if needed), site evaluation forms and other information
necessary to ensure an efficient and safe sampling day.
Field crews will also receive a quick-reference handbook that contains tables and figures
summarizing field activities and protocols from the Field Operations Manual. This waterproof
handbook will be the primary field reference used by field teams after completing the required
field training session. The field teams are also required to keep the field operations manual
available in the field for reference and for possible protocol clarification.
Large-scale and/or long-term monitoring programs such as those envisioned for national
surveys and assessments require a rigorous QA program that can be implemented consistently
by all participants throughout the duration of the monitoring period. Quality assurance is a
required element of all EPA-sponsored studies that involve the collection of environmental data
(USEPA 2000a, 2000b). Field teams will be provided a copy of the integrated Quality Assurance
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and Project Plan (QAPP). The QAPP contains more detailed information regarding QA/QC
activities and procedures associated with general field operations, sample collection,
measurement data collection for specific indicators, and data reporting activities. For more
information on the Quality Assurance procedures, refer to the National Rivers and Streams
Assessment: Quality Assurance Project Plan (EPA 841-B-07-007).
Related NRSA documents include the following: National Rivers and Streams
Assessment: Quality Assurance Project Plan (EPA 841-B-07-007), National Rivers and Streams
Assessment: Site Evaluation Guidelines (EPA 841-B-07-008), and National Rivers and Streams
Assessment: Laboratory Methods Manual (EPA 841-B-07-010 or 841-B-07-011). These
documents are available at: http://www.epa.gov/owow/riverssurvey/index.html.
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2.0 DAILY OPERATIONS SUMMARY
This Field Operations Manual will be used for sampling at both wadeable and non-
wadeable sites. The same indicators will be collected (with the exception of Secchi transparency
and fish tissue, which are only collected at non-wadeable sites), but the sampling will be
conducted with different protocols and equipment. This section presents a general overview of
the activities that a field team is to conduct during a typical 1-day sampling visit to a site,
whether wadeable or non-wadeable. General guidelines for recording data using standardized
field data forms and sample labels are also presented. Finally, safety and health considerations
and guidelines related to field operations are described.
2.1 Sampling Scenario
The Field methods for the NRSA are designed to be completed in one field day for most
sites. Depending on the time needed for both the sampling and travel for the day, an additional
day may be needed to complete sampling or for pre-departure and post-sampling activities
(e.g., cleaning equipment, repairing gear, shipping samples, and traveling to the next site).
Remote sites with lengthy or difficult approaches may require more time, and field crews will
need to plan accordingly.
Each field team should define roles and responsibilities for each team member to
organize field activities efficiently. Minor modifications to the sampling scenario may be made by
teams; however the sequence of sampling events presented in the Figures 2-1 and 2-2
cannot be changed and is based on the need to protect some types of samples from
potential contamination and to minimize holding times once samples are collected.
2.1.1 Non-wadeable Sites
A field crew for a non-wadeable field team typically will consist of four or five people in 2
boats. A minimum of two people are always required in a boat together to execute the sampling
activities and to ensure safety. Typically, in non-wadeable sites, two crew members will work in
the "habitat" boat, and two or three will work in the "fish" boat. One crew member on each boat
is primarily responsible for boat operation and navigation. Any additional team members may
either help collect samples, or may remain on the bank to provide logistical support. A daily field
sampling scenario showing how the work load may be split between team members is
presented in Figure 2-1. The following sections further define the sampling sequence and the
protocols for sampling activities.
2.1.2 Wadeable Sites
A field crew for wadeable sites will typically consist of four people. Any additional team
members may either help collect samples, or may remain on the bank to provide logistical
support. A daily field sampling scenario showing how the work load may be split between team
members is presented in Figure 2-2. The following sections further define the sampling
sequence and the protocols for sampling activities.
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The field team arrives at the site in the early morning to complete the sampling in a
single day. The sampling sequence is to:
• verify site and locate x-site (whole crew),
Divide into 2 groups and:
• conduct in situ measurements of dissolved oxygen, pH, temperature, and
conductivity
• take Secchi disk transparency depth measurements at non-wadeable sites,
• collect water chemistry and chlorophyll a,
• conduct physical habitat characterization,
• collect periphyton samples,
• collect benthic samples,
• collect sediment enzyme samples,
• collect fish samples,
• collect fish tissue samples at non-wadeable sites,
• collect fecal indicator sample,
• filter fecal indicator, chlorophyll a, and periphyton samples,
• preserve and prepare all samples for shipment,
• review field forms,
• report sampling event,
• ship time-sensitive samples.
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Whole Crew
Locate X-site
Verify site as target
Determine launch site & set up staging area
UIUU|J H HOUVIUeb. X
Prepare forms, equipment & supplies
Calibrate multi-probe meter
Load equipment and supplies onto boat
Measure Secchi depth &
in situ temperature, pH,
DO, & conductivity
Collect water chemistry
samples
LOCATE & TRAVEL TO SAMPLING STATIONS
Conduct habitat
characterizations
Collect benthic macroinvertebrate,
periphyton, & sediment enzyme samples
Conduct fish assessment
Collect fecal indicator
sample at Transect K
RETURN TO STAGING AREA
Collect fish tissue samples
Preserve benthic macroinvertebrate,
periphyton, & sediment enzyme samples
& prepare for transport
Filter fecal indicator, chlorophyll-a, & AFDM
samples; prepare for transport
Preserve & prepare fish
tissue & voucher
samples for transport
Inspect and clean boat, motor, & trailer to prevent
transfer of nuisance species and contaminants
Review data forms for completeness
Clean and organize equipment for loading
Report back to Field Logistics Coordinator and
Information Management Coordinator
SHIP SAMPLES
Figure 2-1. Field sampling scenario for non-wadeable sites.
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Whole Crew
Locate X-site
Verify site as target
Set up staging area
Group A Activities:
Prepare forms, equipment & supplies
Group B Activities:
Calibrate multi-probe meter
Lay out sampling reach (from X-site to Transect A)
BEGIN SAMPLING ACTIVITIES AT TRANSECT A
Conduct habitat
characterizations
Collect benthic macroinvertebrate,
periphyton, & sediment enzyme samples
Collect fecal indicator
sample at Transect K
RETURN TO STAGING AREA
Preserve benthic macroinvertebrate, periphyton, &
sediment enzyme samples & prepare for transport
Filter fecal indicator, chlorophyll-a, & AFDM
samples; prepare for transport
Lay out sampling reach (from X-site to Transect K)
RETURN TO TRANSECT F (X-site)
Measure in situ temperature,
pH, DO, & conductivity
Collect water chemistry
samples
TRAVEL TO TRANSECT A
Conduct fish assessment
RETURN TO STAGING AREA
Preserve & prepare fish voucher
samples for transport
Review data forms for completeness
I
Clean and organize equipment for loading
I
Report back to Field Logistics Coordinator and
Information Management Coordinator
SHIP SAMPLES
Figure 2-2. Field sampling scenario for wadeable sites.
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2.2 Recording Data and Other Information
All samples need to be identified and tracked, and associated information for each
sample must be recorded. To assist with sample identification and tracking, labels are
preprinted with sample ID numbers (Figure 2-3).
WATER CHEMISTRY
FWOS
/ 120
999001
WATER CHEM-PPCP
FWOS
; ;20
999003
PERIPHYTON ID
FWOS
; 120
Sample volume:
_
999005
PERIPHYTON BIO
FW08 _____
_ / _ 120
Sample volume:
Vol Filtered /-COiL
999006
BENTHOS - LOW GRADIENT
FWOS
/ 120
Jar 1 of
999008
SAMPLE TYPE
FWOS
CHLOROPHYLL
FWOS
; 120
999002
PHYTOPLANKTON
FWOS
; 120
Vol Filtered mL
999004
PERIPHYTON CHL
Cample volume:
_
999005
PERIPHYTON APA
FW08 _____
/ _ 120 __
Sample volume: _ ml
999005
BENTHOS - REACH WIDE
FWOS _____
_ ; _ 120 __
Jar 1 of _
999007
SAMPLE TYPE _
FWOS
Sampie volume:
SAMPLE ID:
SAMPLE TYPE.
FWOS
FISH TISSUE
FW08
, 20 TOT,
FISH TISSUE
FWD8
Sample volume:
SAMPLE ID:
Sampie volume:
SAMPLE ID:
Figure 2-3. Example sample labels for sample tracking and identification.
It is imperative that field and sample information be recorded accurately, consistently,
and legibly. The cost of a sampling visit coupled with the short index period severely limits the
ability to resample a site if the initial information recorded was inaccurate or illegible. Guidelines
for recording field measurements are presented in Table 2-1.
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Table 2-1. Guidelines for recording field measurements and tracking information
Activity
Guidelines
Field Measurements
Data Recording
Record measurement values and observations on data forms preprinted on water-
resistant paper.
Use No. 2 pencil only (fine-point indelible markers can be used if necessary) to
record information on forms.
Record data and information using correct format as provided on data forms.
Be sure to accurately record site IDs and sample numbers. For revisit samples use
(site ID)-R to indicate the samples are from revisit sites. For duplicate samples,
use (site ID)-D to indicate the samples are duplicates.
Print legibly (and as large as possible). Clearly distinguish letters from numbers
(e.g., 0 versus O, 2 versus Z, 7 versus T or F, etc.), but do not use slashes.
In cases where information is recorded repeatedly on a series of lines (e.g., physical
habitat characteristics), do not use "ditto marks" (") or a straight vertical line.
Record the information that is repeated on the first and last lines, and then
connect these using a wavy vertical line.
When recording comments, print or write legibly. Make notations in comments field
only; avoid marginal notes. Be concise, but avoid using abbreviations or
"shorthand" notations. If you run out of space, attach a sheet of paper with the
additional information, rather than trying to squeeze everything into the space
provided on the form.
Data Qualifiers
(Flags)
Use only defined flag codes and record on data form in appropriate field.
K = Measurement not attempted or not recorded.
Q = Failed quality control check; remeasurement not possible.
U = Suspect measurement; remeasurement not possible.
Fn = Miscellaneous flags (n = 1, 2, etc.) assigned by a field team during a
particular sampling visit (also used for qualifying samples).
Explain reason for using each flag in comments section on data form.
Sample Labels
Use adhesive labels with preprinted ID numbers and follow the standard recording
format for each type of sample.
Use a pencil to record information on label. Cover the completed label with clear
tape.
Record sample ID number from label and associated collection information on
sample collection form preprinted on water-resistant paper.
Sample Collection and Tracking
Sample
Qualifiers
(Flags)
Use only defined flag codes and record on sample collection form in appropriate
field.
K = Sample not collected or lost before shipment; resampling not possible.
U = Suspect sample (e.g., possible contamination, does not meet minimum
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Activity
Guidelines
acceptability requirements, or collected by non-standard procedure).
Fn = Miscellaneous flags (n=1, 2, etc.) assigned by a field team during a
particular sampling visit (also used for field measurements).
Explain reason for using flags in "Comments" on sample collection form.
Review of
Labels and
Data Collection
Forms
Compare information recorded on labels and sample collection form for accuracy
before leaving site.
Review labels and data collection forms for accuracy, completeness, and legibility
before leaving site.
The Field Team Leader must review all labels and data collection forms for
consistency, correctness, and legibility before transfer to the Information
Management Center.
2.3 Safety and Health
Collection and analysis of samples can involve significant risks to personal safety and
health. This section describes recommended training, communications, and safety
considerations, safety equipment and facilities, and safety guidelines for field operations.
2.3.1 General Considerations
Important considerations related to field safety are presented in Table 2-2. It is the
responsibility of the state or contractor project leader to ensure that the necessary safety
courses are taken by all field personnel and that all safety policies and procedures are followed.
Please follow your own agency's health and safety protocols, or refer to the Health and Safety
Guidance for Field Sampling: National Rivers and Streams Assessment (available from the EPA
Regional Coordinator) and Logistics of Ecological Sampling on Large Rivers (Flotemersch, et al.
(editors) 2000). Additional sources of information regarding safety-related training include the
American Red Cross (1979), the National Institute for Occupational Safety and Health (1981),
U.S. Coast Guard (1987) and Ohio EPA (1990).
Field crew members should become familiar with the hazards involved with sampling
equipment and establish appropriate safety practices prior to using them. Make sure all
equipment is in safe working condition. Personnel must consider and prepare for hazards
associated with the operation of motor vehicles, boats, winches, tools, and other incidental
equipment. Boat operators should meet any state requirements for boat operation and be
familiar with U.S. Coast Guard rules and regulations for safe boating contained in a pamphlet,
"Federal Requirements for Recreational Boats," available from a local U.S. Coast Guard
Director or Auxiliary or State Boating Official (U.S. Coast Guard, 1987). Life jackets must be
worn by crew members at all times on the water. All boats with motors must have fire
extinguishers, boat horns, life jackets or flotation cushions, and flares or communication
devices. Boats should stay in visual contact with each other, and should use 2-way radios to
communicate.
Primary responsibility for safety while electrofishing rests with the crew chief.
Electrofishing units may deliver a fatal electrical shock, and should only be used by qualified,
experienced operators. Field crew members using electrofishing equipment must be insulated
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from the water, boat, and electrodes via rubber boots and linesman gloves. Use chest waders
with nonslip soles and linesman gloves. DO NOT wear breathable waders while electrofishing. If
waders become wet inside, stop fishing until they are thoroughly dry or use a dry pair. Avoid
contact with the anode and cathode at all times due to the potential shock hazard. If you
perspire heavily, wear polypropylene or some other wicking and insulating clothing instead of
cotton. If it is necessary for a team member to reach into the water to pick up a fish or
something that has been dropped, do so only after the electrical current is off and the anode is
removed from the water. Do not resume electrofishing until all individuals are clear of the
electroshock hazard. The backpack electrofishing equipment is equipped with a 45° tilt switch
that interrupts the current. Do not make any modifications to the electrofishing unit that would
hinder this safety switch. Avoid electrofishing near unprotected people, pets, or livestock.
Discontinue activity during thunderstorms or rain. Team members should keep each other in
constant view or communication while electrofishing. For each site, know the location of the
nearest emergency care facility. Although the team leader has authority, each team member
has the responsibility to question and modify an operation or decline participation if it is unsafe.
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Table 2-2. General health and safety considerations.
Recommended Training
• First aid and cardiopulmonary resuscitation (CPR)
• Vehicle safety (e.g., operation of 4-wheel drive vehicles)
• Boating and water safety; Whitewater safety if applicable
• Field safety (weather, personal safety, orienteering, site reconnaissance of prior to sampling
• Equipment design, operation, and maintenance
• Handling of chemicals and other hazardous materials
Communications
• Check-in schedule
• Sampling itinerary (vehicle used & description, time of departure & return, travel route)
• Contacts for police, ambulance, hospitals, fire departments, search and rescue personnel
• Emergency services available near each sampling site and base location
• Cell (or satellite) phone and VHF radio if possible
Personal Safety
• Field clothing and other protective gear including lifejackets for all team members
• Medical and personal information (allergies, personal health conditions)
• Personal contacts (family, telephone numbers, etc.)
• Physical exams and immunizations
A communications plan to address safety and emergency situations is essential. All field
personnel need to be fully aware of all lines of communication. Field personnel should have a
daily check-in procedure for safety. An emergency communications plan should include
contacts for police, ambulance, fire departments, hospitals, and search and rescue personnel.
Proper field clothing should be worn to prevent hypothermia, heat exhaustion, sunstroke,
drowning, or other dangers. Field personnel must be able to swim, and personal flotation
devices must be used. Chest waders made of rubberized or neoprene material must always be
worn with a belt to prevent them from filling with water in case of a fall. A personal flotation
device (PDF) and suitable footwear must be worn at all times while on board a boat.
Many hazards lie out of sight in the bottoms of rivers and streams. Broken glass or sharp
pieces of metal embedded in the substrate can cause serious injury if care is not exercised
when walking or working with the hands in such environments. Infectious agents and toxic
substances that can be absorbed through the skin or inhaled may also be present in the water
or sediment. Personnel who may be exposed to water known or suspected to contain human or
animal wastes that carry causative agents or pathogens must be immunized against tetanus,
hepatitis, typhoid fever, and polio. Biological wastes can also be a threat in the form of viruses,
bacteria, rickettsia, fungi, or parasites.
2.3.2 Safety Equipment
Appropriate safety apparel such as waders, linesman gloves, safety glasses, etc. must
be available and used when necessary. First aid kits, fire extinguishers, and blankets must be
readily available in the field. Cellular or satellite telephones and/or portable radios should be
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provided to field teams working in remote areas in case of an emergency. Supplies (e.g., clean
water, anti-bacterial soap, ethyl alcohol) must be available for cleaning exposed body parts that
may have been contaminated by pollutants in the water.
2.3.3 Safety Guidelines for Field Operations
General safety guidelines for field operations are presented in Table 2-3. Personnel
participating in field activities should be in sound physical condition and have a physical
examination annually or in accordance with organizational requirements. All surface waters and
sediments should be considered potential health hazards due to potential toxic substances or
pathogens. Persons must become familiar with the health hazards associated with using
chemical fixing and/or preserving agents. Chemical wastes can be hazardous due to
flammability, explosiveness, toxicity, causticity, or chemical reactivity. All chemical wastes must
be discarded according to standardized health and hazards procedures (e.g., National Institute
for Occupational Safety and Health [1981]; U.S. EPA [1986]).
During the course of field research activities, field teams may observe violations of
environmental regulations, may discover improperly disposed hazardous materials, or may
observe or be involved with an accidental spill or release of hazardous materials. In such cases
it is important that the proper actions be taken and that field personnel do not expose
themselves to something harmful. The following guidelines should be applied:
First and foremost, protect the health and safety of all personnel. Take necessary steps
to avoid injury or exposure to hazardous materials. If you have been trained to take action such
as cleaning up a minor fuel spill during fueling of a boat, do it. However, you should always err
on the side of personal safety.
Field personnel should never disturb or retrieve improperly disposed hazardous
materials from the field to bring back to a facility for "disposal". To do so may worsen the impact,
incur personal liability for the team members and/or their respective organizations, cause
personal injury, or cause unbudgeted expenditure of time and money for proper treatment and
disposal of the material. Notify the appropriate authorities so they may properly respond to the
incident.
For most environmental incidents, the following emergency telephone numbers should be
provided to all field teams: State or Tribal department of environmental quality or protection,
U.S. Coast Guard, and the U.S. EPA regional office. In the event of a major environmental
incident, the National Response Center may need to be notified at 1-800-424-8802.
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Table 2-3. General safety guidelines for field operations
• Two crew members must be present during all sample collection activities, and no one should be left
alone while in the field. Boats should proceed together down the river.
• Use caution when sampling in swift or deep water. Wear a suitable PFD and consider using a safety
tether held by an assistant.
• Use extreme care walking on riprap. Rocks can shift unexpectedly and serious falls are possible.
• Field crew members using electrofishing equipment must be insulated from the water, boat, and
electrodes via non-breathable waders and linesman gloves. Use chest waders with nonslip soles.
• Electrofishing units may deliver a fatal electrical shock, and should only be used by qualified,
experienced operators.
• Do not attempt to collect samples from vertical or near vertical banks.
• Professional-quality breathable waders with a belt are recommended for littoral sampling only, and at a
safe distance from the electrofishing sampling. Neoprene boots are an alternative, but should have
sturdy, puncture-resistant soles.
• Use caution using the Ponar-type samplers. The jaws are sharp and may close unexpectedly.
• Exposure to water and sediments should be minimized as much as possible. Use gloves if necessary,
and clean exposed body parts as soon as possible after contact.
• All electrical equipment must bear the approval seal of Underwriters Laboratories and must be
properly grounded to protect against electric shock.
• Use heavy gloves when hands are used to agitate the substrate during collection of benthic
macroinvertebrate samples.
• Use appropriate protective equipment (e.g., gloves, safety glasses) when handling and using
hazardous chemicals.
• Crews working in areas with poisonous snakes must check with the local Drug and Poison Control
Center for recommendations on what should be done in case of a bite from a poisonous snake.
• Any person allergic to bee stings, other insect bites, or plants (i.e., poison ivy, oak, sumac, etc.) must
take proper precautions and have any needed medications handy.
• Field personnel should also protect themselves against deer or wood ticks because of the potential
risk of acquiring pathogens that cause Rocky Mountain spotted fever and Lyme disease.
• Field personnel should be familiar with the symptoms of hypothermia and know what to do in case
symptoms occur. Hypothermia can kill a person at temperatures much above freezing (up to 10°C or
50°F) if he or she is exposed to wind or becomes wet.
• Field personnel should be familiar with the symptoms of heat/sun stroke and be prepared to move a
suffering individual into cooler surroundings and hydrate immediately.
• Handle and dispose of chemical wastes properly. Do not dispose any chemicals in the field.
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3.0 BASE SITE ACTIVITIES
Field teams conduct a number of activities at their base site (i.e., office or laboratory,
camping site, or motel). These include tasks that must be completed both before departure to
the site and after return from the field (Figure 3-1). Close attention to these activities is required
to ensure that the field teams know (1) where they are going, (2) that access is permissible and
possible, (3) that equipment and supplies are available and in good working order to complete
the sampling effort, and (4) that samples are packed and shipped appropriately.
PREDEPARTURE ACTIVITIES
Team Leader
• Prepare daily itinerary
Crew Members
• Instrument checks & calibration
• Equipment & supplies preparation
Whole Crew
Site Verification
SAMPLE SITE
POSTSAMPLING ACTIVITIES
Team Leader
• Review forms & labels
• File status report by email to the
tracking team email addresses
Crew Members
• Filter, preserve, & inspect samples
• Clean boats with 1-10% bleach solution & perform
safety checks (boat, trailer, equipment)
• Clean (and repair, if needed) sampling gear
• Charge or replace batteries
• Refuel vehicle and boat
• Obtain ice and other consumable supplies as needed
• Package and ship samples & data forms
Figure 3-1. Overview of base site activities.
3.1 Predeparture Activities
Predeparture activities include the development of a daily itinerary, instrument checks
and calibration, and equipment and supply preparation. Procedures for these activities are
described in the following sections.
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3.1.1 Daily Itineraries
The Field Team Leaders are responsible for developing daily itineraries. This entails
compiling maps, contact information, copies of permission letters, and access instructions (a
"site packet"). Additional activities include confirming the best access routes, calling the
landowners or local contacts, confirming lodging plans, and coordinating rendezvous locations
with individuals who must meet with field teams prior to accessing a site. Changes in the
itinerary during the week, such as canceling a sampling day, must be relayed by the crew leader
to the Field Logistics Coordinator as soon as possible.
3.1.2 Instrument Checks and Calibration
Each field team must test and calibrate instruments prior to sampling. Calibration can be
conducted prior to departure for the site or at the site, with the exception of dissolved oxygen
(DO) calibration. Because of the potential influence of altitude, DO calibration is to be performed
only at the site. Field instruments include a global positioning system (GPS) receiver, a
multiprobe unit for measuring DO, pH, temperature, and conductivity, and electrofishing
equipment. Field teams should have access to backup instruments if any instruments fail the
manufacturer performance tests or calibrations. Prior to departure, field teams must:
• Turn on the GPS receiver and check the batteries. Replace batteries immediately if a
battery warning is displayed.
• Test and calibrate the multi-probe meter. Each field team should have a copy of the
manufacturer's calibration and maintenance procedures. All meters should be
calibrated according to manufacturer specifications provided along with the meter.
Once a week, crews should check their multiprobe against the provided Quality
Check Solution. This QCS is provided to all crews in their base kits and is used to
check pH and conductivity measurements.
• Turn on the electrofishing unit and check the batteries. Be sure to have fully charged
backup batteries. If using a gas powered electrofishing unit, check the oil and gas
supply.
3.1.3 Equipment and Supply Preparation
Field teams must check the inventory of supplies and equipment prior to departure using
the equipment and supplies checklists provided in Appendix A; use of the lists is mandatory.
Specific equipment will be used for wadeable vs. non-wadeable sites; be sure to bring both sets
of equipment if you are unsure what type of site you will be visiting that day. Pack meters,
probes, and sampling gear in such a way as to minimize physical shock and vibration during
transport. Pack stock solutions as described in Table 3-1. Follow the regulations of the
Occupational Safety and Health Administration (OSHA).
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Table 3-1. Stock solutions, uses, and methods for preparation.
Solution
Bleach (1-10%)
Calibration
QCS
Lugol's
95% Ethanol
Formalin
Use
Clean nets, gear, and inside of boat
QCS for pH and conductivity calibration
Preserve periphyton ID samples
Preserve benthic samples
Preserve fish voucher samples
Preparation
Add 10 -100 ml bleach to 1
L distilled water.
None (included in site kits)
None (included in site kits)
None
None
Site kits of consumable supplies for each sampling site will be delivered based on the
schedule each crew provides prior to the sampling season. Field crew leaders MUST provide
a schedule in order to receive the site kits. If your schedule changes, report the change as
soon as possible to the Field Logistics Coordinator (Jennifer Pitt: jennifer.pitt@tetratech.com:
copyTara.Kolodiei@tetratech.com: 410-356-8993). The site kit will include data forms, labels,
sample jars, bottles, filters, and other supplies (see complete list in Appendix A). The teams
must inventory these site kits before departure. The teams should also label and package the
sample containers into site kits prior to departure. Container labels should not be covered with
clear tape until all information is completed during sampling at the river/stream. Store extra site
kits of sampling supplies in the vehicles. Inventory these extra site kits prior to each site visit.
3.2 Post Sampling Activities
Upon return to the launching location after sampling, the team must review all completed
data forms and labels for accuracy, completeness, and legibility and make a final inspection of
samples. If information is missing from the forms or labels, the Field Team Leader is to provide
the missing information. The Field Team Leader is to initial all data forms after review. If
obtainable samples are missing, the site should be rescheduled for complete sampling. Other
post sampling activities include: inspection and cleaning of sampling equipment, supply
inventory, sample and data form shipment, and communications.
3.2.1
Review Data Forms and Labels
The field crew leader is ultimately responsible for reviewing all data forms and labels for
accuracy, completeness, and legibility. Ensure that written comments use no "shorthand" or
abbreviations. The data forms must be thoroughly reviewed. Upon completing the review, the
field crew leader must initial the field forms to indicate that they are ready to be sent to the
Information Management Center. Each sample label must also be checked for accuracy,
completeness, and legibility. The field crew leader must cross-check the sample numbers on the
labels with those recorded on the data forms.
3.2.2 Inspect and Prepare Samples
All samples need to be inspected and appropriately preserved and packaged for
transport. Check that all samples are labeled, and all labels are completely filled in. Check that
each label is covered with clear plastic tape. Check the integrity of each sample container, and
be sure there are no leaks. Make sure that all sample containers are properly sealed. Make sure
that all sample containers are properly preserved for storage or immediate shipment.
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3.2.3 Equipment Cleanup and Check
All equipment and gear must be cleaned and disinfected between sites to reduce the risk
of transferring nuisance species and pathogens. Species of primary concern in the U.S. include
Eurasian watermilfoil (Myriophyllum spicatum), zebra mussels (Dreissena polymorpha), New
Zealand mud snails (Potamopyrgus antipodarum), Myxobolus cerebralis (sporozoan parasite
that causes salmonid whirling disease), and Batrachochytrium dendrobatidis (a chytrid fungus
that threatens amphibian populations). Field crews must be aware of regional species of
concern, and take appropriate precautions to avoid transfer of these species. There are several
online resources regarding invasive species, including information on cleaning and disinfecting
gear, such as the Whirling Disease Foundation (www.whirling-disease.org), the USDA Forest
Service (Preventing Accidental Introductions of Freshwater Invasive Species, available from
http://www.fs.fed.us/invasivespecies/documents/Aquatic is prevention.pdf), and the California
Dept. of Fish and Game (Hosea and Finlayson 2005). General information about freshwater
invasive species is available from the U.S. Geological Survey Nonindigenous Aquatic Species
website (http://nas.er.usgs.gov), the Protect Your Waters website that is co-sponsored by the
U.S. Fish and Wildlife Service (http://www.protectyourwaters.net/hitchhikers), and the Sea Grant
Program (http://www.sgnis.org).
Handle and dispose of disinfectant solutions properly, and take care to avoid damage to
lawns or other property. Table 3-2 describes equipment care. Inspect all equipment, including
nets, boat, and trailer, and clean off any plant and animal material. Prior to leaving a site, drain all
bilge water and live wells in the boat. Inspect, clean, and handpick plant and animal remains from
vehicle, boat, motor, and trailer. Before moving to the next site, if a commercial car wash facility is
available, wash vehicle, boat, and trailer and thoroughly clean (hot water pressurized rinse-no
soap). Rinse equipment and boat with 1% -10% bleach solution to prevent the spread of exotics.
Note that many organizations now recommend against using felt-soled wading boots in affected
areas due to the difficulty in removing myxospores and mudsnails.
3.2.4 Supply Inventory
A site kit containing field forms, labels, and consumable supplies (see App. A) will be
provided to the field crews for each sampling site. Site kits will be shipped out based on the
schedule that each field crew provides prior to the start of the sampling season. Field crew
leaders MUST provide a schedule in order to receive the site kits. Crews should include
in this schedule the primary fish taxonomist at each site. If your schedule changes, please
report the change as soon as possible to the Field Logistics Coordinator (Jennifer Pitt:
iennifer.pitt@tetratech.com: copy Tara.Kolodiei(S)tetratech.com: 410-356-8993). Prior to
sampling, inspect each site kit to ensure all supplies are included. Store an extra, complete
backup site kit in the vehicle. Check the inventory of supplies and equipment at the end of the
day using the checklists provided in Appendix A. Make sure specific supplies are not running
low due to sampling errors, accidental loss, or increased demand at certain sites (e.g., some
sites may require extra benthic macroinvertebrate bottles). Make sure you have enough site kits
for sites that will require duplicate samples.
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Table 3-2. Postsampling equipment care
1. Clean for biological contaminants.
• Prior to departing site, drain all water from live wells and buckets used to hold and process fish, and
drain all bilge water from the boat.
• Inspect motor, boat, trailer, sampling gear, waders, boots, etc. for evidence of mud, snails, plant
fragments, algae, animal remains, or debris, and remove using brushes or other tools.
• At the base location, inspect and rinse periphyton sampling equipment, dip nets, kick nets, waders,
and boots with water and dry. Use one of the procedures below to disinfect gear if necessary.
Additional precautions to prevent transfer of Whirling Disease spores, New Zealand mudsnails,
and amphibian chytrid fungus.
Before visiting the site, consult the site dossier and determine if it is in an area where whirling
disease, New Zealand mud snails, or chytrid fungus are known to exist. Contact the local State
fishery biologist to confirm the existence or absence of these organisms.
• If the stream is listed as "positive" for any of the organisms, or no information is available, avoid
using felt-soled wading boots, and, after sampling, disinfect all fish and benthos sampling gear
and other equipment that came into contact with water or sediments (i.e., waders, boots, etc.) by
one of the following procedures:
Option A:
1. Soak gear in a 10% household bleach solution for at least 10 minutes, or wipe or spray on
a 50% household bleach solution and let stand for 5 minutes
2. Rinse with clean water (do not use stream water), and remove remaining debris
3. Place gear in a freezer overnight or soak in a 50% solution of Formula 409® antibacterial
cleaner for at least 10 minutes or soak gear in 120°F (49°C) water for at least 1 minute.
4. Dry gear in direct sunlight (at least 84 °F) for at least 4 hours.
Option B:
1. Soak gear in a solution of Sparquat® (4-6 oz. per gallon of water) for at least 10 minutes
(Sparquat is especially effective at inactivating whirling disease spores).
2. Place gear in a freezer overnight or soak in 120°F (49°C) water for at least 1 min.
3. Dry gear in direct sunlight (at least 84 °F) for at least 4 hours.
2. Clean and dry other equipment prior to storage.
• Rinse coolers with water to clean off any dirt or debris on the outside and inside.
• Rinse periphyton sampling equipment with tap water at the base location.
• Make sure conductivity meter probes are rinsed with deionized water and stored moist.
• Rinse carboy and all beakers used to collect water chemistry samples three times with deionized
water. Place beakers in a 1-gallon scalable plastic bag with a cube container for use at the next
stream.
• Check nets for holes and repair or locate replacements.
3. Inventory equipment and supply needs and relay orders to the Field Logistics Coordinator.
4. Remove GPS, multi-probe meter, and electrofishing unit from carrying cases and set up for
predeparture checks and calibration. Examine the oxygen membranes for cracks, wrinkles, or
bubbles. Replace if necessary, allowing sufficient time for equilibration.
5. Recharge/replace batteries as necessary.
6. Replenish fuel and oil; if a commercial car wash facility is available, thoroughly clean vehicle and boat
(hot water pressurized rinse—no soap).
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3.2.5 Shipment of Samples and Forms
The field team must ship or deliver time-sensitive samples (i.e., water chemistry,
chlorophyll a) to the appropriate analytical laboratories as soon as possible after collection.
Other samples (see App. C) may be shipped or delivered in batches provided they can be
adequately preserved. Batched samples should be shipped every two weeks. Field teams are to
fill out one sample tracking form for each sample shipment. On each sample tracking form, the
following information must be recorded:
• Airbill or package tracking number
• Date sample(s) were sent
• Site ID where each sample was collected
• Sample type code:
CHEM - Chemistry ENTE - Enterococci
CHLA - Chlorophyll a BERW - Benthos (reach-wide sample)
SEDE- Sediment enzymes BELG - Benthos (low gradient)
PERI - Periphyton FTIS - Fish Tissue
PAPA - Periphyton APA VOUC-- Fish voucher sample
Date when the sample(s) was collected (1st day if sampling took >1 day)
Site visit number (e.g., 1 for first visit, 2 for re-visit)
Sample ID number encoded on label
Number of containers for each sample
For Fish Tissue samples (FTIS), record species and length of each fish specimen
under Comments
Any additional comments
Packaging and shipping guidelines for each type of sample are summarized in
Figure 3-3. Detailed sample shipping instructions are presented in Appendix C.
After checking the Field Forms for completeness and accuracy, the Field Crew Leader
will make copies of all Field Forms and retain the copies. The original forms will be mailed to
Marlys Cappaert in the FedEx envelope provided in the site kit. A pre-addressed airbill will
be provided. The original forms must be sent because they are printed specifically to be used in
a scanner for automated data entry. Field forms may be retained and mailed in batches
throughout the field season (about every 2 weeks) when it is convenient to make the copies.
3.2.6 Status Reports and Communications
After each sampling event, the field crew leader must file a status report via email. This
status report email must be sent before the water chemistry/chlorophyll sample is shipped, and
no later than the following morning after each sampling event. An electronic tracking and
sample status report form will be emailed to the field crew leaders after their training session.
Complete the tracking and sample status report form for each site, even sites that are
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visited but not sampleable, and email the form to SampleTracking@epa.gov. If you are not
able to fill out the electronic form, the Tracking and Sample Status form provided in the field kits
can be faxed on a non VOIP fax machine or called into the number provided on the bottom of
the TSS form.
The separate, scanable Tracking and Sample Status form (Fig 3.2) provided in the set of
field forms must be filled out first; the information from this form will be used to fill out the status
report form. The scanable Tracking and Sample Status form will then be shipped in the
container with the samples. A tracking form must accompany every sample.
You must follow a standardized naming convention when naming the electronic status
report files. The naming convention for fresh samples is "labid_siteid_datecollected.doc:"
ex. WRS_FW08OR123_05_05_2008.doc
For batch/retained samples, the naming convention is "BR_siteid_datecollected.doc:"
ex. BR_FW08OR123_05_05_2008.doc (in this case, the site id and date collected will
refer to the first sample on the page)
It is very important to complete the status report after every sampling event. This will
enable the Field Logistics Coordinator to track sampling progress. More importantly, it will
enable the Information Management Center to track which samples were collected at each site,
and to immediately track the shipment of the time-sensitive water chemistry and chlorophyll
samples that will be shipped after each sampling event. If the form cannot be emailed by the
following morning after sampling, fax the scanable Tracking and Sample Status form (Fig 3.2) or
phone in ALL of the information (read the ENTIRE form to the voice mail machine) to the
Information Management Coordinator:
Information Management Coordinator: Marlys Cappaert
Sample Tracking (phone): 541-754-4663; Sample Tracking (fax): 541-754-4637
A second form will be provided to track batched and retained samples while they are
being held and when they are in transit to the appropriate laboratory. This form must be filled
out and emailed right away when samples are brought into your lab or holding facility, and then
again when the samples are shipped. The scanable Tracking (Batched and Retained) Form (Fig
3.3) will be filled out and shipped in the container with the samples.
The field crews should call or email the Field Logistics Coordinator (Jennifer Pitt; 410-
356-8993; Jennifer.Pitt@tetratech.com) to report any problems encountered. The Field Logistics
Coordinator monitors all aspects of field sampling activities. The Field Logistics Coordinator and
Information Management Coordinator will contact the EPA Headquarters Coordinator regularly
to provide regional updates throughout the sampling period. The EPA Headquarters Coordinator
will maintain a database of all sampling activities and reconnaissance information. For questions
or problems related to fish tissue or PPCP water sampling, contact Leanne Stahl or Elaine
Snyder. See Appendix E for contact information.
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The EPA Regional Coordinator serves as the central point of contact for information
exchange among field teams, the management and QA staffs, the information management
team, and the public. A list of EPA Regional Coordinators and their contact information can be
found at the beginning of this manual on page xv.
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TRACKING AND SAMPLE STATUS - WRS
SITE ID: FW08XJX"^£?O
SENT BY: J. fHfUUlf
State of Site Location. yf V i
Visit*:* 1 O2
0»ta toHi-cteb Q J 1 Q 1 1 2 0 @ f
SENDER PHONE: ^^ tfff_^fo
rE^ XX- 1 DATE SENT ^7/^^/20^^
SHIPPED • FedEx OUPS O Hand Delivery
BY: ,-, <-..., AIRBILL/TRACKING
U Other; NUMBER:
///4«u3yvy
Site Status Report
SAMPLEABLE J NOT SAMPLEABLE
O Wadeabie JO Dry - Visited
• Beatable O Dry - Not Visited
O Partial Wadeable O Wetland
O Partial Boatable O Map Error
O Wadeable interrupted O Impounded
O Boatablo Interrupted O Other
O Altered
TemporarMy
Not Sampleable —
O Not Boatable
O Not Wadeable '
O Other °
o
NO ACCESS
O Access Denied
O
O Inaccessible ,_
O Temp Inaccessible p,
SAMPLE STATUS
O No Samples * All Sample Types
Collected COHPC|BIJ
only some samp^ts w^ns coMected, indicate those betew:
«ater Chem (CHEM; Q Enlerococci (ENTE)
Water Chi (WCHL) O Sediment (SEDE)
-'erphyton Chi (PCHL) O Fish Tissue (FT1S)
=>eriphyton Bio (PBIOj O Ben! Reachwide (8ERW)
Deriphyton 10 (PERI) O Bent Low Gradient (BElG)
3erphyfon APA (PAPA)O FJhy1oplankton (PHYT)
Status Comments
S«,pl.O Sw,pl.fyp,
9 f f.O.O. 1 , CH E t.; |_
-S—^-Jf" ^ O.JL , C H L A
?,?,?£* O £~ 2^ P C H L
*f 1 t O. O f 3 P B t r,
.,,=-,-,-.-*.„* _J£.j....™jL—" — 3K-t — ILf - IMf j Bil..,._j s. t j
• • , ._ 1
. .
Sample Types Condition Codes
CHE&f . Water chemtelry Filled in b^ reciPle
WCHl -Wftlftr Column C = Cracked Jar
Chlofopliyll f s FfOfstt
PCHt - Peripdylan t = Leaktn-g
Chlorophyll ML ffi Mfsstng iab&l
PSIO - Pnrtphyi.on NP ss Not preserved
Biomasa W - Warm
OK x Sample OK
T = Thawed
j
Ccnnwtt.
Chain of Custody
nt Filled in by recipii
Date Received:
Received toy:
Contact Information
-nt Tracking Help:
Marlys Cappaert
PH: 541-754-4467
Lab:
Attn: Phil Monaco, Dynarnac
c/o U.S. EPA
1350 Goodnight A ve
Corvallis, OR 97333
PH: 541-754-4787
monaco.phll@epamail, epa.gov
FAX THIS FORM TO 541 -754-4637 Ur»ti
mm OR READ TRACKING INFO TO VOfCE MESSAGE CENTER: * ^m mm
mm 541-754-4663 Ufc»J •
Figure 3-2. Tracking and Sample Status Form
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page 30
TRACKING {BATCHED and RETAINED SAMPLES) National Rivers and Streams Assessment
D0tf
SENDER
PHONE
STATE OF
MUtlTAIION X
BATCHED SAMPLES • UNPRESERVED
, i i . , „ . , i, , , , , , _ ,„ , t ,
„ SHIPPED
FedEx "i UPS ' Hand Delivery Other
AIRB1LUT RACKING
NUMBER:
/ / 2 0
Dal.SampU
Coll«(.d
tea... }j
*•*».
RETAINED SAMPLES - PRESERVED
/ 1 f \ 2 Q 0
COI 1 FfTFD
C*rtr*i.
OPJVTPFD OFF AJ HOLLilNb I AL.lu.ITi iADUF-tSa
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xx
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* '
0
» 1
io 2
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Lab
Chain of Custody
Sample Types
Condition Codes
MED - DULUTH
NERL
FISH TISSUE LAB
j PFRIPHYTON LAB
/) BENTHIC LAB
FISH MUSEUM
OTHER
Filled in by recipient
Date Received.
/ I ...
Received by:
Tracking Help:
Marlys Cappffiffsl
p) 541-7MJ»4«7
PRESERVED -RETAINED
B£RW - eeothos Reach Wide
BELG - e*nB»« Low Oradi.nl
VERT - Ftsh Vouehefs
PERI - Psrfphylen Dot
UNPBESERVED - BATCHED
PAP* - P>npnyUkn APA i 4)
ENTE - Ens^roccxx
Filled In by recipient
C " Ct»Ck«<( )«r
F = frozen
L = Leaking
ML '" Minsing label
Np . Ng, p,.s.rvK)
W.W.rm
OK = Sample OK
T = Thawed
FAX THIS FORM TO 541-754-4637
OR READ TRACKING INFO TO VOICE MESSAGE CENTER: 541-754-4663
Figure 3-3. Tracking (Batched and Retained) Form
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page 31
PPCPs
2 5WmL
glass jars
WATER
CHEM
4L
cutaitainer
AFDM
(50-mL
tube)
CHLOR-a
(2 filters,
each in
50-mL tube)
SEDIMENT
(500 ml
jar)
APA
(50-mL
tube)
ENTEROCOCCI
(filters in vials)
o
LU
Preserve
on wet ice
LU
III
Freeze
immediately
on Dry Ice
PPCP FISH
TISSUE
(in foil &
double
bagged)
Preserve
on wet ice
Refrigerate
until
shipping
Keep frozen
until
shipping
SHIP ON WET ICE
ASAP AFTER
COLLECTION
OVERNIGHT
COURIER
REQUIRED
Ship M-Th
No Sat
delivery
Ship in batches on wet ice
(1-2 weeks)
OVERNIGHT
COURIER
REQUIRED
Saturday
delivery OK
FISH TISSUE
(in foil &
double
bagged)
, _•
i
Hold on wet
ice; freeze
within 6 hours
on dry ice
Keep frozen until
shipping (1-2 weeks)
Ship in batches on dry ice
OVERNIGHT COURIER
REQUIRED
Ship M-Th
overnight
No Sat delivery
OVERNIGHT COURIER
REQUIRED
Package and ship using
dry ice protocols
Ship M-Th
No Sat delivery
GLEC lab
(remaining
boatable
sites)
EPA
Cincinnati,
OH lab (PPCP
sites only)
*PPCP samples are only collected at a subset of pre-selected sites
Figure 3-4. Sample packaging and shipping procedures for unpreserved samples
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National Rivers and Streams Assessment
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Date: April 2009
Page 32
Q
LU
PERIPHYTON
(ID sample in
50-mLtube)
Ship in batches (1-2 weeks)
LU
CO
LLI
OVERNIGHT OR
GROUND COURIER
Formalin and Ethanol Must be
Packaged & Shipped
as Dangerous Goods
Ship M-Th if overnight
No Sat. delivery
Ml State
University & Philadelphia
Academy of Natural
Sciences
Figure 3-5. Sample packaging and shipping procedures.
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page 33
4.0 INITIAL SITE PROCEDURES
When you arrive at a site, you must first confirm you are at the correct site, and then
determine if the site meets the criteria for sampling and data collection activities (See Site
Evaluation Guidelines EPA-841-B-07-008). Inspect the selected reach for appropriate access,
safety, and general conditions. Decide whether the site is at base flow condition and not unduly
influenced by rain events which could affect the representativeness of field data and samples. If
you determine that the site can be sampled, lay out a defined reach within which all sampling
and measurement activities are conducted.
4.1 Site Verification Activities
4.1.1 Locati ng the X-Site
River and stream sampling points were chosen using the National Hydrography Dataset
(NHD), in particular NHD-Plus, following a systematic randomized selection process (Stevens
and Olsen, 2004). Each point is referred to as the "X-site." The "X-site" is the mid-point of the
sampling reach, and it will determine the location and extent for the rest of the sampling reach.
The latitude/longitude of the "X-site" is listed on the site spreadsheet that was distributed by the
EPA Regional Coordinators.
Conditions encountered at rivers and streams across the country will vary tremendously.
To orient the crews and help them anticipate sampling and access challenges, EPA MED
prepared site dossiers for all of the sampling sites. Each dossier contains maps with the X-site
plotted, and they show general conditions at each site at two scales. The "watershed" scale
page shows the position of the site in the landscape and stream network. The "site" scale page
shows the area around the site where samples will be taken.
Watershed Page Overview
The watershed page (Figure 4-1) shows land cover (National Land Cover Data 2001),
cities, major roads, stream networks, and county, state, watershed and catchment boundaries of
the site's watershed. The map scale and level of detail for this page varies according to
watershed size. Catchments (nominally, a site's local watershed) are spatially nested within the
stream's watershed. Catchment boundaries and hydrologic connectivity were defined in the
National Hydrography Dataset Plus (http://www.horizon-systems.com/nhdplus/: NHDPIus) using
a Digital Elevation Model (DEM). Watersheds are aggregates of all the catchments upstream
from a site. In small watersheds, the catchment may be the entire watershed. In large
watersheds, the catchment may not be visible. Pour-points are the downstream end of the
watershed. Catchment and watershed attributes (Table 4-1) include areas downstream of the
site to the pour-point.
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National Rivers and Streams Assessment
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7r2?27"K 7T>\y-rms! 7ri7;rw
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Figure 4-1. Watershed page
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National Rivers and Streams Assessment
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Table 4-1. Landscape and NHDPIus attributes for the watershed page (data were summarized from
NHDPIusandNLCD2001)
Measure
Area
NLCD2001 land cover classes
Mean annual precipitation
Mean annual temperature
Stream order
Flow
Velocity
Elevation
Slope
Area-weighted mean annual precipitation
Area-weighted mean annual temperature
Scale
Catchment
Watershed
Catchment
Watershed
Catchment
Catchment
Stream (flowline)
Stream (flowline)
Stream (flowline)
Stream (flowline)
Stream (flowline)
Stream (flowline)
Stream (flowline)
Units
km2
% area
mm
C°x10
Strahler units
cfs
fps
meters
cm/cm
mm
C°x10
Site Page Overview
The site page (Figure 4-2) shows the area immediately surrounding the sampling site.
The sampling site, roads, and stream lines are labeled on an aerial photograph. Aerial imagery
is provided by ArcGIS Online and features i-cubed Nationwide Select imagery. This dataset
consists of imagery from various sources and time periods. For more information on the imagery
in these maps, please see http://arcgisonline.esri.com (Layer name: ESRI_lmagery_World_2D).
Road data is provided by the U.S. EPA and features 2007 Tele Atlas North America data. The
catchment boundary and pour-point are noted. The map scale is fixed at 1:8,000. In some wide
rivers, the scale ratio was reduced in order to show shorelines. Sampling stations within the site
are distributed according to mean channel width (refer to National Rivers and Streams
Assessment Field Operations Manual; EPA 841-B-07-009, 2008). Tabular information includes
Site ID, river name, stream order, state, county, latitude and longitude coordinates of the site.
An inset map locates the site in the state.
Table 4-2 is the checklist for equipment and supplies required to conduct site verification
protocols described in this section. It is a subset of the checklist in Appendix A that is used at a
base site to assure that all equipment and supplies are taken to and available at the site. While
traveling from a base location to a site, record a detailed description of the route taken on page
1 of the Verification Form (Figure 4-3). This information will help others find the site again in the
future. Upon reaching the X-site, confirm its location and verify that you are at the correct
stream. Use all available means to accomplish this, including map coordinates, locational data
from the GPS, and any other evidence such as signs or conversations with local residents, and
record the information on page 1 of the Verification Form (Figure 4-3). Complete a verification
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National Rivers and Streams Assessment
Field Operations Manual
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Date: April 2009
Page 36
form for each site visited (regardless of whether you end up sampling it), following the
procedures described in Table 4-3.
River Na me
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Figure 4-2. Site page.
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National Rivers and Streams Assessment
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Date: April 2009
Page 37
Table 4-2. Equipment and supplies list for site verification.
For locating and
verifying site
Sampling permit and landowner access(if required)
Field Operations Manual and/or laminated quick reference guide
Site dossier, including access information, site spreadsheet with map
coordinates, street and/or topographic maps with "X-site" marked
NRSA Fact Sheets
GPS unit (preferably one capable of recording waypoints) with manual,
reference card, extra battery pack
Surveyor's flagging tape (to mark transects if not using GPS waypoints)
Laser rangefinder
50 m or 100 m measuring tape with reel (if not using rangefinder)
For recording
measurements
Clipboard
#2 pencils
Site Verification Form
Fine-tipped indelible markers to write on flagging
4.1.2 Determining the Sampling Status of a Stream
After you confirm the location of the X-site, evaluate the stream reach surrounding the X-
site and classify the stream into one of three major sampling status categories: sampleable,
non-sampleable, or no access (Table 4-3). The primary distinction between "Sampleable" and
"Non-Sampleable" streams is based on the presence of a defined stream channel, water
content during base flow, and adequate access to the site.
Even if there is no water at the X-site coordinates, you may still sample the site as an
"interrupted flow" stream (Section 4.3.1). If the channel is dry at the X-site, determine if there is
water present anywhere within the sampling reach. There must be greater than 50% water
throughout the channel reach. If there are isolated pools of water within the reach that equal
greater than 50% of the reach length, proceed to sample using the modified procedures outlined
in Section 4.3.1. If less than 50% of the reach has water, classify the site as "Dry-visited" on the
verification form. NOTE: Do not "slide" the reach (Section 4.2) for the sole purpose of obtaining
more water to sample (e.g., the downstream portion of the reach has water, but the upstream
portion does not).
Record the sampling status and pertinent site verification information on the Verification
Form (Figure 4-3). If the site is non-sampleable or inaccessible, no further sampling activities
are conducted. Replace the site with the first oversample site on the state list within the
appropriate Strahler order category (Section 1.1.2). Notify the EPA Regional Coordinator and
Field Logistics Coordinator (Section 3.2.6) that the site was replaced.
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National Rivers and Streams Assessment
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STREAM VERIFICATION FORM - WADEABLE/BOATABLE (Front)
SITE NAME: f/^fff KlVeTK. DATE Oil O j / 2 0 O g» VISIT: • 1 O2 O3
SITE ID: FWQ8XX 0£5O Slate of Site
Don"* forget to record **
Reach Lengtti on back. TEAM, /*?**' I
STREAM/R1VER VERIFICATION INFORMATION
Stream/River Verified by jfill m all that appty): 0 GPS O Local Contact • Signs 4 Roads 0 Taps, Msip
O Other (Describe Here): O Not Verified (Explain In Cwnments)
Coordinates Latitude North
"•E^^T if f # 7 / »
MAP OR '
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an-d Sfficomlsi *_/ c /I / / r
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OR
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O Farias! - Sampl&d by boa! f>SO% of reach sampled). Explain beEow.
O Watd^ablt Mtefrupied - Not continuous waiw ^long reach
O Boatabie fnterrupfed « NoS cortiinoou'S water along reach
O AfNwd - Stre'Sm/Rsver Channel Present but differs from Map
>, ! NO M NO, check one below
NON -SAMPLE ABLE-PERMANENT
• "i Dry • Vmuid
O Dry - Not vm£«ri
0 W«rt!and (No Definable Channel)
Q Map Ertor - No evidence channehi«3l6fbody ever present
O Impounded (UndemMth L^ikes or Pofsd)
O Other ^explain in comments)
NON-SAMPLEABLE-TEMPORARY
O Not beatable - N«d a different crew • K©*eh*du!tr fef this year
O Not wadeabUe - N€>sd a diifferenl crew - Reschedule for this year
O Other (Explnln In comments)
NO ACCESS
O Access Pernns'siiDn DeBie-d
O Permanently inaccessible ^UnabteAJnsate? to Keach Srte)
U Temporarily Inaccessibte-Fire, etc. - Reschedule For next year
GENERAL COMMENTS; |
DIRECTIONS TO STREAM/RIVER SITE: | f~^om
B£*T0*>. G-o sovrtf- a*j fiivr*.
Af-tess Hens C €**r 8**>K"\. PUBLIC L*u*>e*f r/Tw jtr fi^&t- ctffit
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Record Infofn^atiori y^ed to define length of reach, and sketch general f
03^06/2008 NRSA Slnsam Verification
iL^es of reach on
Figure 4-3. Verification Form (page 1).
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National Rivers and Streams Assessment Final Manual
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Page 39
Table 4-3. Site Verification Procedures
1. Find the stream/river location in the field corresponding to the X-site coordinates and the "X" marked
on the maps prepared for each site in the site dossier. Record the routes taken and other directions on
the Verification Form so that others can visit the same location in the future. If the site is non-
wadeable, locate public or private launch sites.
2. Use a GPS receiver to confirm the latitude and longitude at the X-site with the coordinates provided for
the site (datum = NAD 27). Record these on the Verification Form.
3. Use all available means to insure you are at the correct stream/river as marked on the map, including
1:24,000 USGS maps, topographic landmarks, road maps, signs, local contacts, etc.
4. Scan the channel upstream and downstream from the X-site, decide if the site is sampleable, and
mark the appropriate circle on the verification form. If the channel is dry at the X-site, determine if
water is present within 75 m upstream and downstream of the X-site. Assign one of the following
sampling status categories to the stream. Record the category on the Verification Form.
SAMPLEABLE CATEGORIES
• Wadeable - Continuous water, >50% wadeable
• Boatable
• Partial - Sampled by wading (>50% of reach sampled)
• Partial - Sampled by boat (>50% of reach sampled)
• Wadeable Interrupted: not continuous water along reach
• Boatable Interrupted: not continuous water along reach
• Altered Channel: Stream/river channel present but differs from map.
NON-SAMPLEABLE CATEGORIES
Permanent
• Dry Channel: Less than 50% water within the reach. Record as "Dry-Visited." If site was
determined to be dry (or otherwise non-perennial) from another source and/or field verified before
the actual sampling visit, record as "Dry-Not visited".
• Wetland: Standing water present, but no definable stream channel. If wetland is surrounding a
stream channel, define the site as Target but restrict sampling to the stream channel.
• Map Error: No evidence that a water body or stream channel was ever present at the X-site.
• Impounded stream: Stream is submerged under a lake or pond due to man-made or natural (e.g.,
beaver dam) impoundments. If the impounded stream is still wadeable, record it as "Altered" and
sample.
• Other: Examples would include underground pipelines, or a non-target canal. A sampling site
must meet both of the following criteria to be classified as a non-target canal:
The channel is constructed where no natural channel has ever existed.
The sole purpose/usage of the reach is to transfer water. There are no other uses of the
waterbody by humans (e.g., fishing, swimming, boating).
Temporary
• Not Boatable - need a different crew
• Not Wadeable - need a different crew
• Other: The site could not be sampled on that particular day, but is still a target site. Examples
might include a recent precipitation event that has caused unrepresentative conditions.
NO ACCESS TO SITE CATEGORIES
• Access Permission Denied: You are denied access to the site by the landowners.
• Permanently Inaccessible: Site is unlikely to be sampled by anyone due to physical barriers that
prevent access to the site (e.g., cliffs).
• Temporarily Inaccessible: Site cannot be reached due to barriers that may not be present at a
future date (e.g. forest fire, high water, road temporarily closed, unsafe weather conditions)
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National Rivers and Streams Assessment Final Manual
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5. Do not sample non-target or "Non-sampleable" or "No Access" sites. Fill in the "NO" circle for "Did you
sample this site?" and check the appropriate circle in the "Non-Sampleable" or "No Access" section of
the Verification Form; provide detailed explanation in comments section.
4.1.3 Sampling During or After Rain Events
Avoid sampling during high flow rainstorm events. It is often unsafe to be in the water
during such times. In addition, biological and chemical conditions during such episodes are
often quite different from those during baseflow. On the other hand, sampling cannot be
restricted to only strict baseflow conditions. It would be next to impossible to define "strict
baseflow" with any certainty at an unstudied site. Such a restriction would also greatly shorten
the index period when sampling activities can be conducted. Thus, some compromise is
necessary regarding whether to sample a given stream because of storm events. To a great
extent, this decision is based on the judgment of the field team. Some guidelines to help make
this decision are presented in Table 4-4. The major indicator of the influence of storm events will
be the condition of the stream itself. If you decide a site is unduly influenced by a storm event,
do not sample the site that day. Notify the Field Logistics Coordinator or other central contact
person to reschedule the stream for another visit.
Table 4-4. Guidelines to determine the influence of rain events
• If it is running at bank full discharge or the water seems much more turbid than typical for the class of
stream do not sample it that day.
• Do not sample that day if it is unsafe to be in the water.
• Keep an eye on the weather reports and rainfall patterns. Do not sample a stream during periods of
prolonged heavy rains.
• If the stream seems to be close to normal summer flows, and does not seem to be unduly influenced
by storm events, sample it even if it has recently rained or is raining.
4.1.4 Site Photographs
Taking site photographs is an optional activity, but should be considered if the site has
unusual natural or man-made features associated with it. If you do take photographs with a
digital camera at a site, date-stamp the photograph and include the site ID. Alternatively, start
the sequence with one photograph of an 8.5 x 11 inch piece of paper with the site ID, waterbody
name, and date printed in large, thick letters. After the photo of the site ID information, take at
least two photographs at the X-site, one in the upstream direction and one downstream. Take
any additional photos you find interesting after these first three pictures. Keep a log of your
photographs and briefly describe each one.
4.2 Laying out the sampling reach
Unlike chemistry, which can be measured at a point, most of the biological and habitat
structure measures require sampling a certain length of a stream to get a representative picture
of the ecological community. A length of 40 times the channel width is necessary to characterize
the habitat and several biotic assemblages associated with the sampling reach. Establish the
sampling reach about the X-site using the procedures described in Tables 4-5a (non-wadeable
sites) and 4-5b (wadeable sites). It is highly recommended that you lay out the sampling reach
for large, non-wadeable sites before you go in the field using maps, aerial photos, and/or GIS
software. This will save time on the field day.
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National Rivers and Streams Assessment Final Manual
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Page 41
Scout the sampling reach to make sure it is clear of obstacles that would prohibit
sampling and data collection activities. Record the channel width used to determine the reach
length, and the sampling reach length upstream and downstream of the X-site on page 2 of the
Verification Form as shown in Figure 4-4. Figures 4-5 and 4-6 illustrate the principal features of
the established sampling reach for both non-wadeable and wadeable sites, including the
location of 11 cross-section transects used for collecting samples and physical habitat
measurements. The figures also show the specific sampling stations on each cross-section tran-
sect at the two different types of sites for collection of sediment enzyme, periphyton, and benthic
macroinvertebrate samples.
Before leaving the stream, complete a rough sketch map of the stream reach you
sampled on page 2 of the Verification Form (Figure 4-4). In addition to any other interesting
features that should be marked on the map, note any landmarks/directions that can be used to
find the X-site for future visits.
Table 4-5a. Laying out the sampling reach at non-wadeable sites
Laying out the sampling reach at the base site (recommended at boatable sites)
1. On an aerial photo or a 1:100:000 topographic map, locate the X-site using the coordinates provided
for the site and the maps prepared in the site dossier for the site.
2. Determine the average wetted width of the channel at the X-site using maps and/or aerial
photographs. To get an average, determine the wetted width of the channel at 5 places of "typical"
width within approximately 5 channel widths upstream and downstream from the X-site. Average the 5
readings together and round to the nearest 1 m.
3. Multiply the average wetted width by 40 to determine the reach length. If the average width is <4 m,
use 150 m as a minimum reach length. If the average width is >100 m, use 4 km as a maximum
reach length.
4. From the X-site, measure a distance of 20 channel widths downstream using CIS software. Be careful
to measure all of the bends of the river/stream; do not artificially straighten out the line of
measurement. The downstream endpoint is marked as Transect K. Measure 20 channel widths
upstream from the X-site; the upstream end of the reach is marked as Transect A.
5. Measure 1/10 of the reach length downstream from Transect A, and mark this spot as Transect B.
Continue marking the 11 transects A- K in increments of 1/10 of the reach length. Enter the waypoints
for the transects into a GPS unit so the transects are easy to find on the sampling day.
6. Assign the sampling station at Transect A randomly (e.g., use the seconds display on a digital watch
to select the initial sampling station: 1 - 5 = Left Bank, 6 - 9 = Right Bank). From here, three stations
will be on the first (randomly selected) side of the river, then 2 on the other, then 2 on the first side,
and so on through Transect K (as shown in Figure 4-5).
7. When you are at the site, "ground truth" the wetted width measurements and proceed to Steps 9 & 10
to see if the layout needs to be adjusted.
Laying out the sampling reach in the field
8. Use a laser range finder to determine the wetted width of the channel at 5 places of "typical" width
within approximately 5 channel widths upstream and downstream from the X-site. Average the 5
readings together and round to the nearest 1 m. If the average width is <4 m, use 150 m as a
minimum reach length. If the average width is >100 m, use 4 km as a maximum reach length. Record
this width on page 2 of the Site Verification Form.
For channels with "interrupted flow", estimate the width based on the unvegetated width of
the channel (again, with a 150 m minimum and 4 km maximum).
9. Check the condition of the stream about the X-site by having one team member go upstream and one
downstream. Each person proceeds until they can see the stream to a distance of 20 times the
average channel width (equal to one-half the sampling reach length) determined in Step 1.
10. Determine if the reach needs to be adjusted about the X-site due to confluences with higher order
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streams (downstream), or a change to a lower order streams (upstream), impoundments (lakes,
reservoirs, ponds), physical barriers (e.g., falls, cliffs), or because of access restrictions to a portion of
the initially-determined sampling reach. Refer to Table 4-6 for specific instructions.
11. Starting at the X-site (or the new midpoint of the reach if it had to be adjusted as described in Step 10),
measure a distance of 20 channel widths downstream using a GPS unit, laser rangefinder, or tape
measure. Be careful to measure all of the bends of the river/stream; do not artificially straighten out the
line of measurement. Enter the channel to make measurements only when necessary to avoid
disturbing the stream channel prior to sampling activities. The downstream endpoint is flagged as
Transect K. The upstream end of the reach is flagged as Transect A.
12. Sampling Stations at non-wadeable sites
At Transect A, use the seconds display on a digital watch to select the initial sampling station for
transect samples: 1 - 5 = Left Bank, 6 - 9 = Right Bank. Mark "L" or "R" on the transect flagging.
13. Measure 1/10 of the reach length downstream from Transect A. Flag this spot as Transect B. Assign
the sampling station systematically after the first random selection as shown in Figure 4-5. Three
stations will be on the first side of the river, then 2 on the other, then 2 on the first side, and so on
through Transect K.
14. Proceed downstream with a GPS unit, laser rangefinder, or tape measure and flag the positions of 9
additional transects (labeled "C" through "K" as you move upstream) at intervals equal to 1/10 of the
reach length. Continue to assign the sampling stations systematically.
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Table 4-5b. Laying out the sampling reach at wadeable sites
1. Use a surveyor's rod, tape measure, or laser range finder to determine the wetted width of the
channel at 5 places of "typical" width within approximately 5 channel widths upstream and
downstream from the X-site. Average the 5 readings together and round to the nearest 1 m. If the
average width is <4 m, use 150 m as a minimum reach length. If the average width is >100 m, use 4
km as a maximum reach length. Record this width on page 2 of the Site Verification Form.
For channels with "interrupted flow", estimate the width based on the unvegetated width of
the channel (again, with a 150 m minimum and 4 km maximum).
2. Check the condition of the stream about the X-site by having one team member go upstream and one
downstream. Each person proceeds until they can see the stream to a distance of 20 times the
average channel width (equal to one-half the sampling reach length) determined in Step 1.
3. Determine if the reach needs to be adjusted about the X-site due to confluences with higher order
streams (downstream), a change to a lower order streams (upstream), impoundments (lakes,
reservoirs, ponds), physical barriers (e.g., falls, cliffs), or because of access restrictions to a portion of
the initially-determined sampling reach. Refer to Table 4-7.
4. Starting at the X-site (or the new midpoint of the reach if it had to be adjusted as described in Step 3),
measure a distance of 20 channel widths down one side of the stream using a GPS unit, laser
rangefinder, or tape measure. Be careful not to "cut corners". Enter the channel to make
measurements only when necessary to avoid disturbing the stream channel prior to sampling
activities. This endpoint is the downstream end of the reach, and is flagged as Transect "A".
5. Sampling Stations at wadeable sites:
At Transect A, use the seconds display on a digital watch to select the initial sampling station for
standard transect samples: 1-3="Left", 4-6="Center", 7-9=Right. Mark "L", "C", or "R" on the transect
flagging; the 3 potential collection points are roughly equivalent to 25%, 50%, and 75% of the channel
width, respectively.
6. Measure 1/10 of the required reach length upstream from transect A. Flag this spot as transect B.
Assign the sampling station systematically after the first random selection (Figure 4-6 & Table 4-6).
7. Proceed upstream with the tape measure and flag the positions of 9 additional transects (labeled "C"
through "K" as you move upstream) at intervals equal to 1/10 of the reach length. Continue to assign
the sampling stations systematically.
8. Benthic macroinvertebrates at "low gradient" streams: A second, separate composite is collected at
low gradient streams to include the edge habitats (0%, 50%, and 100% channel width). The initial
sampling station will be the first to the right of the one selected for the standard sample (Table 4-5).
For example, if the sampling station for transect A (standard), was "C", then the initial transect A
sampling station for the second sample would be "R". This second pattern would be R, L, C, R, ....
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STREAM VERIFICATION FORM - WADEABLE/BOATABLE (Back) "££'
SITE NAME:
fffvf/f
DATE:
I.O/I 2 °
VISIT: »1 O2 O3
SITE ID;
STREAMIRIVER REACH DETERMINATION
Channel Width
Used to Drfino
Reach (m)
DISTANCE (m) FROM X-SITE
Upstream
Length
Downstream
Length
Total Reach
Length trrtsneted
Comment
Z.H.O.
S.f.O.
-L
SKETCH MAP - Arrow Indcates North; Mark sit© L=taunch Xslmf&x T« Taka Out
NOTE: If &n outlin* map (s attachs-d here, usfi a ccmtimwws strip ol etear tape across She top e-dge,
You can also aitach a s*f|jaeiit@ sh*®l with the Gtrtiiifse map &n it.
¥w hostage s«Ses you can attseh Eopo map with reach, X-site and t*a^s»cl loeay-cmffi marfe&d
&
.frf
PERSONNEL
NAME
Blo/Chem
Sampling Habitat
* O
* O
Forms
O
o
Draft
83*96/2908 MRSA Stmm Verification
Figure 4-4. Verification Form (page 2)
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Upstream endpoint is "Transect A"
Downstream endpoint is "Transect K"
Distance between transects
= 4 x mean wetted width
Sampling Stations
• L = left; R = right
• 1 st station (at transect A)
determined randomly; subsequent
stations assigned systematically
* Stations extend 15m from bank
and 5rn up & downstream from
each transect (1 Om x 15m)
K
Total reach length = 40 x mean wetted width (min = 150 m; max = 4 km)
Figure 4-5. Sampling reach features for a non-wadeable site.
Distance between transects=4 times
mean wetted width at X -site
^ Total reach length=40 times mean wetted width at X -site (minimum=150 m)^
SAMPLING POINTS
• L=Left C=Center R=Right
• First point (transect A)
determined at random
• Subsequent points assigned in
order L, C, R
Figure 4-6. Sampling reach features for a wadeable site.
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Table 4-6. Sample point distribution in wadeable streams (the Transect A sample point for the
standard sample is randomly selected; the secondary sample point distribution is used only to collect the
second benthic macroinvertebrate sample in low gradient wadeable streams (L=left, C=center, R=right))
PRIMARY SAMPLE
Transect A
If you randomly
select
"LEFT"
Transect....
Then continue
sequence....
B
C
C
R
D
L
E
C
F
R
G
L
H
C
1
R
J
L
K
C
SECONDARY SAMPLE- Low gradient benthic macroinvertebrate only
Transect A
Select next in
sequence to start
2nd pattern
"CENTER"
Transect....
Then continue
sequence....
B
R
C
L
D
C
E
R
F
L
G
C
H
R
1
L
J
C
K
R
There are some conditions that may require sliding the reach about the X-site (i.e., the
X-site is no longer located at the midpoint of the reach) to avoid features we do not wish to or
physically cannot sample across. Sliding the reach involves noting the distance of the barrier,
confluence, or other restriction from the X-site, and flagging the restriction as the endpoint of the
reach. Add the distance to the other end of the reach, such that the total reach length remains
the same, but it is no longer centered about the X-site. Table 4-7 describes when you should
and should not slide the sampling reach.
Table 4-7. Sliding the sampling reach
1. Slide the reach if you run into an impoundment (lake, pond, or reservoir), so that the lake/stream
confluence is at one end.
2. Slide the reach if you run into an impassible barrier (e.g., waterfall, cliff, navigation dam) so that the
barrier is at one end.
3. When you are denied access permission to a portion of the reach, you can slide the reach to make it
entirely accessible; use the point of access restriction as the endpoint of the reach.
4. Note the distance of the barrier, confluence, or other restriction from the X-site, and flag the restriction
as the endpoint of the reach. Add the distance to the other end of the reach, so the total reach length
remains the same, but it is no longer centered about the X-site.
5. Do not slide the reach so that the X-site falls outside of the reach boundaries.
6. Do not proceed upstream into a lower order stream or downstream into a higher order stream
when laying out the stream reach (order is based on 1:100,000 scale maps).
7. Do not slide a reach to avoid man-made obstacles such as bridges, culverts, rip-rap, or channel-
ization. These represent important features and effects to study.
8. Do not slide a reach to gain more water to sample if the flow is interrupted (Section 4.3.1).
9. Do not slide a reach to gain better habitat for benthos or fish,
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4.3 Modifying Sample Protocols for High or Low Flows
4.3.1 Streams with Interrupted Flow
You cannot collect the full complement of field data and samples from streams that are
categorized as "Interrupted" (Table 4-8). Note that no data should be collected from streams
that are completely "Dry" as defined in Table 4-8. Interrupted streams will have some cross-
sections amenable to biological sampling and habitat measurements and some that are not. To
be considered target, streams must have greater than 50% water in the reach length within the
channel ( can be isolated pools). Modified procedures for interrupted streams are presented in
Table 4-8. Samples for water chemistry (Section 5) will be collected at the X-site (even if the
reach has been adjusted by "sliding" it). If the X-site is dry and there is water elsewhere in the
sample reach, collect the sample from a location having water with a surface area >1 m2 and a
depth >10 cm.
Collect data for the physical habitat indicator along the entire sample reach from
interrupted streams, regardless of the amount of water present at the transects. Obtain depth
measurements along the deepest part of the channel (the "thalweg") along the entire sampling
reach to provide a record of the "water" status of the stream for future comparisons (e.g., the
percent of length with intermittent pools or no water). Other measurements associated with
characterizing riparian condition, substrate type, etc., are useful to help infer conditions in the
stream when water is flowing.
Table 4-8. Reach layout modifications for interrupted streams
• Streams with less than 50% of reach length containing water (not necessarily continuous)
are considered dry and are not sampled.
• If more than 50% of the channel has water and if the X-site is dry but there is flowing water or
a pool of water having a surface area > 1 m2 and a depth > 10 cm somewhere along the
defined sampling reach, take the water sample at the pool or flowing water location that is
nearest to the X-site. Note that the sample was not collected at the X-site and where on the
reach the sample was collected on the field data form.
• Do not collect a water sample if there is no acceptable location within the sampling reach.
Record a "K" flag for the chemistry sample on the sample collection form and explain why the
sample was not collected in the comments section of the form.
Physical Habitat, Periphyton, Sediment Enzymes, and Benthic Macroinvertebrates
• Obtain a complete thalweg profile for the entire reach. At points where channel is dry, record
depth as 0 cm and wetted width as 0 m.
• At each of the transects (cross-sections), sample the stream depending on flow status:
DRY CHANNEL: No surface water anywhere in cross-section; collect all physical habitat data. Use
the unvegetated area of the channel to determine the channel width and the subsequent location of
substrate sampling points. Record the wetted width as 0 m. Record substrate data at the sampling
points located in the unvegetated, but dry, channel. Do not collect periphyton, sediment enzymes, or
benthic macroinvertebrates from this transect.
DAMP CHANNEL: No flowing water at transect, only puddles of water < 10 cm deep; collect all
physical habitat data. Do not collect periphyton, sediment enzymes, or benthic macroinvertebrates
from this transect.
WATER PRESENT: Transect has flow or pools > 10 cm deep; collect all data and measurements for
physical habitat, periphyton, sediment enzymes, benthic macroinvertebrate, and fish indicators, using
standard procedures.
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4.3.2 Partially Wadeable Sites
Some wadeable sites will have sections that are too deep or swift to wade safely, and it
will be impossible to do all of the wadeable sampling protocols at every transect. At these sites,
keeping safety in mind, try to do as much sampling and data collection as you can with the
wadeable procedures. The amount of sampling that can actually be done while wading will
depend on the extant conditions. Only sample or measure what can be done safely. Make
detailed comments on the Verification Form describing what the conditions were like and where
sampling occurred. Use the sketch map on the back of the Verification Form to indicate problem
areas and where samples were collected if you had to go off transect. If barriers prohibit
physically reaching the X-site, then the site is not a Sampleable site; it should be coded as "No
Access - Inaccessible" on the Verification Form.
4.3.3 Braided Rivers and Streams
Depending upon the geographic area and/or the time of the sampling visit, you may
encounter a stream having "braided" channels, which are characterized by numerous sub-
channels that are generally small and short, often with no obvious dominant channel. If you
encounter a braided stream, establish the sampling reach using the procedures presented in
Table 4-9. Figuring the mean width of extensively braided rivers and streams for purposes of
setting up the sample reach length is challenging. For braided channels, measure the mean
width and bankfull width as defined in the physical habitat protocols (Sections 5.2 and 6.2). For
relatively small streams (mean bankfull width <15 m) the sampling reach is defined as 40 times
the mean bankfull width. For larger streams (>15 m), sum the actual wetted width of all the
braids and use that as the width for calculating the 40 channel width reach length. If there is any
question regarding an appropriate reach length for the braided system, it is better to
overestimate. Make detailed notes and sketches on the Verification Form (Fig. 4-3 and Fig. 4-4)
about what you did. It is important to remember that the purpose of the 40 channel width reach
length is to sample enough stream to incorporate the variability in habitat types. Generally, the
objective is to sample a long enough stretch of a stream to include 2 to 3 meander cycles (about
6 pool-riffle habitat sequences). In the case of braided systems, the objective of this protocol
modification is to avoid sampling an excessively long stretch of stream. In a braided system
where there is a 100 m wide active channel (giving a 4 km reach length based on the standard
procedure) and only 10 m of wetted width (say five, 2 m wide braids), a 400 m long sample
reach length is likely to be sufficient, especially if the system has fairly homogenous habitat
throughout its length.
Table 4-9. Modifications for braided streams
1. Estimate the mean width as the bankfull channel width as defined in the physical habitat protocol.
• If the mean width is <15 m, set up a 40 x channel width sample reach in the normal manner.
• If >15 m, sum up the actual wetted width of all the braids and use that as the width for calculating
the 40 x channel width reach length. Remember the minimum reach length is always 150 m.
• If the reach length seems too short for the system in question, set up a longer sample reach,
taking into consideration that the objective is to sample a long enough stretch of a stream to
include at least 2 to 3 meander cycles (about 6 pool-riffle habitat sequences).
2. Make detailed notes and sketches on the Verification Form about what you did.
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5.0 NON-WADEABLE RIVERS
5.1 Water Quality
This section describes the procedures and methods for the field collection and analysis
of the water quality indicators (in-situ measurements, water chemistry, Secchi Disk
transparency, and sediment enzymes) from non-wadeable streams and rivers. Refer to
Appendix E for PPCP water sampling procedures at the designated urban river sites.
5.1.1 In Situ Measurements of Dissolved Oxygen, pH, Temperature, and Conductivity
5.1.1.1 Summary of Method
Measure dissolved oxygen (DO), pH, temperature, and conductivity using a calibrated
multi-parameter water quality meter (or sonde). Take the measurements mid-channel at the X-
site. Take the readings at 0.5 m depth. Measure the site depth accurately before taking the
measurements. Take care to avoid the probe contacting bottom sediments, as the instruments
are delicate.
5.1.1.2 Equipment and Supplies
Table 5.1-1 provides the equipment and supplies needed to measure dissolved oxygen,
pH, temperature, and conductivity. Record the measurements on the Field Measurement Form,
as seen in Figure 5.1-1.
Table 5.1-1. Equipment and supplies—DO, pH, temperature, and conductivity
For taking measurements and
calibrating the water quality meter
For recording measurements
• Multi-parameter water quality meter with pH, DO,
temperature, and conductivity probes.
• Extra batteries
• De-ionized and tap water
• Calibration cups and standards
• QCS calibration standard
• Barometer or elevation chart to use for calibration
• Field Measurement Form
• Pencils (for data forms)
5.1.1.3 Multi-Probe Sonde
Dissolved Oxygen Meter
Calibrate the DO meter prior to each sampling event. It is recommended that the probe
be calibrated in the field against an atmospheric standard (ambient air saturated with water)
prior to launching the boat. In addition, manufacturers typically recommend periodic
comparisons with a DO chemical analysis procedure (e.g., Winkler titration) to check accuracy
and linearity.
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FIELD MEASUREMENT FORM - BO AT ABLE
f\ J~M
SITE ID: FW08 / * 0 O
/ JU / 2009
M £ .iT S
Clear to Bottom?
Flag
Flag cgdas: K = No meaauramant
cww, Explain sfi fla^s in
MRS A Field Measurement Beatable
Figure 5.1 -1. Field Measurement Form.
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pH Meter
Calibrate the pH meter prior to each sampling event. Calibrate the meter in accordance
with the manufacturer's instructions and with the team agency's existing SOP. You must also
conduct a quality control check with the provided standard to verify the calibration and
periodically evaluate instrument precision (see Section 3.1.2). Once a week, each crew must
check their multi-probe against the QCS that was in each base kit. Any irregularities must be
reported to the Field logistics coordinator immediately.
Temperature Meter
Check the accuracy of the sensor against a thermometer that is traceable to the National
Institute of Standards (NIST) at least once per sampling season. The entire temperature range
encountered in the NRSA should be incorporated in the testing procedure and a record of test
results kept on file.
Conductivity Meter
Calibrate the conductivity meter prior to each sampling event. Calibrate the meter in
accordance with the manufacturer's instructions. The entire conductivity range encountered in
the NRSA should be incorporated in the testing procedure and a record of test results kept on
file. You must also conduct a quality control check with the provided standard to verify the
calibration and periodically evaluate instrument precision (see Section 3.1.2). Once a week,
each crew must check their multi-probe against the QCS that was in each base kit. Any
irregularities must be reported to the Field logistics coordinator immediately.
5.1.1.4 Sampling Procedure
Table 5.1-2 presents step-by-step procedures for measuring dissolved oxygen, pH,
temperature, and conductivity.
Table 5.1-2. Sampling procedure—temperature, pH, conductivity and dissolved oxygen.
1. Check meter and probes and calibrate according to manufacturer's specifications.
2. Check the calibration against the provided QCS solution for pH and conductivity and record the
results on the field sheet as the QCS Measured value. This should be done at least once a week.
3. Record the true value of the QCS solution from the stock solution container on the field sheet as QCS
True.
4. Samples are taken mid-channel, at the X site, at a depth of 0.5 meters or at a mid-depth if less than 1
meter deep.
5. Lower the sonde in the water and measure DO, pH, temperature, and conductivity at 0.5 m depth.
6. Record the measurements on the Field Measurement Form.
7. Flag any measurements that the team feels needs further comment or when a measurement cannot
be made.
8. If sampling at the X-site is not possible, move to another part of the reach to take the measurements
(as close to the X-site as possible), record the letter of the nearest transect in the "TRANSECT" box
and more detailed reasons and/or information in the Comments section.
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5.1.2 Water Chemistry Sample Collection and Preservation
5.1.2.1 Summary of Method
The water chemistry samples will be analyzed for total phosphorus (TP), total nitrogen
(TN), total ammonia-nitrogen (NH4), nitrate (NO3), basic anions, cations, total suspended solids
(TSS), turbidity, acid neutralizing capacity (ANC, alkalinity), dissolved organic carbon (DOC),
and total organic carbon (TOC). You will also collect a 2-L sample in an amber Nalgene bottle to
be filtered on shore for later analysis of chlorophyll a (See Section 7 for filtration procedure).
Store all samples in darkness on ice in a closed cooler. After you filter the chlorophyll a
samples, the filters must be kept frozen until ready to ship.
Collect the samples at mid-channel at the X-site of the river from a depth of 0.5 meters.
Use the 3 L Nalgene beaker to fill the individual sample bottles. The 3 L Nalgene beaker will be
rinsed and re-used at each sampling location.
5.1.2.2 Equipment and Supplies
Table 5.1-3 provides the equipment and supplies needed to collect water samples at the
index site. Record the Water Sample Collection and Preservation data on the Sample Collection
Form, Side 1 as seen in Figure 5.1-2.
Table 5.1-3. Equipment and supplies—water chemistry sample collection and preservation
For collecting samples
Laser Rangefinder
Nitrile gloves
one 2-L amber Nalgene bottle (chlorophyll)
4-L cube container
3 L Nalgene beaker
Cooler with ice
Field Operations Manual and/or laminated Quick Reference Guide
For recording
measurements
Sample Collection Form
Field Measurement Form
Pencils (for data forms)
fine-tipped indelible markers (for labels)
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SAMPLE COLLECTION FORM - BO AT ABLE (Front)
(Initials);
: .ri.
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5.1.2.3 Sampling Procedure
Table 5.1-4 describes the sampling procedures for collecting water chemistry samples in
non-wadeable streams and rivers. Refer to Appendix E for PPCP water sampling procedures at
the designated urban river sites.
Table 5.1-4. Sampling procedure for non-wadeable sites—water chemistry sample collection
1. Collect the water samples from the X-site in a flowing portion near the middle of the stream.
2. Put on nitrile gloves. Make sure not to handle sunscreen or other chemical contaminants until
after the sample is collected.
3. Rinse the 3-L Nalgene beaker three times with water, and discard the rinse downstream.
4. Remove the cube container lid and expand the cube container by pulling out the sides. NOTE:
DO NOT BLOW into the cube container to expand them, this will cause contamination.
5. Fill the 3-liter beaker with water and slowly pour 30 - 50 ml into the cube container. Cap the cube
container and rotate so that the water contacts all the surfaces. Discard the water downstream.
Repeat this rinsing procedure 2 more times.
6. Fill the beaker with water and pour into the cube container. Repeat as necessary to fill the cube
container. Let the weight of the water expand the cube container. Pour the water slowly as the
cube container expands. Fill the cube container to at least three-fourths of its maximum volume.
Rinse the cube container lid with water. Eliminate any airspace from the cube container, and cap
it tightly. Make sure the cap is tightly sealed and not on at an angle.
7. Fill the 3-liter beaker with water and slowly pour 30 - 50 mL into the 2 L amber Nalgene bottle.
Cap the bottle and rotate so that the water contacts all the surfaces. Discard the water
downstream. Repeat this rinsing procedure 2 more times.
8. Fill the beaker with water and pour into the 2 L amber Nalgene bottle. Cap the bottle tightly
9. Place the cube container and bottle in a cooler (on ice or water) and shut the lid. If a cooler is not
available, place the cube container in an opaque garbage bag and immerse it in the stream.
10. Record the Sample ID on the Sample Collection Form along with the pertinent stream information
(stream name, ID, date, etc.). Note anything that could influence sample chemistry (heavy rain,
potential contaminants) in the Comments section. If sampling at the X-site is not possible, move
to another part of the reach to collect the sample (as close to the X-site as possible), record the
letter of the nearest transect and more detailed reasons and/or information in the Comments
section.
5.1.3 Secchi Disk Transparency at Non-Wadeable Sites
5.1.3.1 Summary of Method
A Secchi disk is a black and white patterned disk used to measure water clarity (see
Figure 5.1-3). A Secchi disk transparency reading will be collected mid-channel at the X-site.
The Secchi disk will be affixed to the end of a solid metered rod (e.g., Schedule 80 PVC pipe, or
equivalent) and lowered into the water until it disappears from sight. Measurements are
recorded at the depth that the disk disappears and again when it reappears. The reading is
taken on the shady side of the boat, without sunglasses, hat or view aids.
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Metal or Plastic Disk
o | gye Bolt
' Metal Weight
Figure 5.1-3. Secchi disk diagram (EPA, 1991).
5.1.3.2 Equipment and Supplies
Table 5.1-5 lists the equipment and supplies needed to measure Secchi disc
transparency. Record the Secchi disk readings on the Field Measurement Form, Side 1 as seen
in Figure 5.1-1.
Table 5.1-5. Equipment and supplies—Secchi disc transparency
For taking measurements and
calibrating the water quality meter
For recording measurements
• 20 cm diameter Secchi disk and calibrated sounding
rod (marked in half centimeter intervals)
• Tape measure (in centimeters)
• Field Measurement Form
• Pencils (for data forms)
5.1.3.3 Sampling Procedure
Because different people measuring Secchi disk transparency at the same site may
obtain different results (due to differences in vision and interpreting disk disappearance and
reappearance), one team member will conduct Secchi disk measurements for all sites. Table
5.1-6 lists the procedure for Secchi disk transparency at non-wadeable sites.
If the water is shallow and clear, the Secchi disk might reach the bottom and still be
visible. If this is the case, it is important to not stir up the bottom sediments while anchoring the
boat. Be sure to move the boat away from the anchor before taking the reading. If the disk is
visible at the bottom, indicate this on the form.
Table 5.1-6. Sampling procedure at non-wadeable sites—Secchi disk transparency
1. Measure Secchi disk transparency mid-channel at theX-site.
2. Confirm that the lowering rod is firmly attached to the Secchi disk.
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3.
4.
5.
6.
Remove sunglasses and hats. Also, do not use view scopes or other visual aids. If wearing
prescription sunglasses, temporarily replace them with regular clear lens prescription glasses.
Lower the Secchi disk over the shaded side of the boat until it disappears.
Read the depth indicated on the lowering rod. If the disappearance depth is <1.0 meter, determine
the depth to the nearest 0.05 meter by marking the line at the nearest depth marker and measuring
the remaining length with a tape measure. Otherwise, estimate the disappearance depth to the
nearest 0.1 meter. Record the disappearance depth on the Sample Collection Form.
Lower the disk a bit farther and then slowly raise the disk until it reappears and record the
reappearance depth on the Field Measurement Form.
7. Note any conditions that might affect the accuracy of the measurement in the comments field.
5.1.4 Sediment Enzymes
5.1.4.1 Summary of Method
Collect sediment samples at the 11 sampling stations at each site and combine all
stations at a site, resulting in a single 500 mL sample per site. Collect fine surface sediments
(top 5 cm) using a spoon or dredge. Store samples on ice until shipment to the laboratory for
processing. Samples will be analyzed for available DIN, NH4, DIP, TP, TN, total carbon (TC),
and enzyme activity.
5.1.4.2 Equipment and Supplies
Table 5.1-7 lists the equipment and supplies needed to collect sediment enzyme
samples. Record collection data on Side 2 of the Sample Collection Form, as seen in Figure
5.1-4.
Table 5.1-7. Equipment and supplies—sediment enzymes
For collecting
samples
Petite Ponar sampler with plastic
tub, drop line, and spare pinch pin
Standard Ponar may substitute.
Graduated plastic bucket with lid
Large stainless steel spoon for collecting
& mixing sediment composite
500-mL plastic bottle for storing
sediment sample
For recording
measurements
Sample Collection Form
Sample labels
Pencils
Fine-tipped indelible markers (for labels)
Clear tape strips
5.1.4.3 Sampling Procedure
Near each of the macroinvertebrate and periphyton sampling locations, collect a fine-
grained sediment sample using a spoon. If the depth is too great to reach the bottom with the
spoon, a "petite Ponar" grab sampler can be used to collect sediment and the stainless steel
spoon can take the sample to be added to the composite bucket from the ponar. The objective
is to collect a 500-mL composite sample that is representative of depositional areas at the site.
The composite sample will be subsampled in the laboratory for multiple analyses. Table 5.1-8
presents step-by-step procedures for collecting sediment enzyme samples.
Table 5.1-8. Sampling procedure—sediment enzymes
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1. Collect a sediment sample at each of the 11 transect sampling stations, near the periphyton and
macroinvertebrate sample locations. Make sure each subsample comprises an approximately equal
portion of the total composite. You may collect sediment between stations to insure at least 500 ml
of composite volume (note any deviations from standard procedure in a comment.)
2. Locate sediment samples in areas or patches of fine-grained substrate (silty sand, silt, clay, muck) in
a zone bounded on the shore side by the apparent low-water mark from daily flow fluctuations (often
detected by the presence of periphyton or attached filamentous algae just below the low-water mark)
and bounded on the riverside by the 0.3-m depth contour (recommended maximum sample depth;
deeper sampling may be possible). If samples cannot be safely collected by wading at a station due
to vertical banks or other reason go to step 5.
3. Avoid the area that has just been kick sampled for macroinvertebrates. Sampling up-stream from the
kick sample location is recommended. If fine substrates are not present within 5 m up- or downstream
from the station, flag the station on the form.
4. If fine substrate is present, use the stainless steel spoon to collect a sample (approximately one
spoonful of sediment) from the top 5 cm of substrate. Place the sample in a clean bucket. Use gloves
for handling sediment. Do not assume rip rapped shorelines lack fine-grained sediment. Look for
fines between the large rocks.
5. If the littoral zone cannot be waded, use a petite Ponar (or similar) sampler deployed from the boat to
collect a sediment sample adjacent to the station. (Use caution with Ponar samplers. The jaws are
sharp and may close unexpectedly. Replace frayed lines and worn parts.) Raise the Ponar sampler
from the water and into a plastic tub rather than from the boat deck. This prevents feet from getting
under the sampler. Release the petite Ponar sample into a tub and use the scoop to collect about 15
x 15 cm (6x6 inches) of the top 5 cm of the sample. Using the stainless steel spoon, take a one
spoon grab from the top layer of sediment captured in the Ponar. Place this in the composite bucket
and discard the rest.
6. Repeat steps 2-5 at each of the 11 littoral stations. Record the total number of replicates (stations)
included in the composite. Note in a comment the stations at which sediment was collected using a
non-wading method.
7. It is important that a sufficient sediment (not less than 500 mL) composite sample for analysis be
collected. If multiple stations have no fine sediment, it is permissible to collect extra sample at
stations that do have fine sediment or between stations. Be sure to note this in a comment.
8. Using the stainless steel spoon, thoroughly mix the composite sample and transfer 500 mL into the
500 mL plastic bottle. Place in a in a cooler with ice for final labeling and preservation.
9. Prepare a label for the sample jar. Using a fine-point indelible marker, fill in the site # and sample
date. Place the label on the jar and cover it with clear tape. Record the sample ID and other data on
sampling form. Place the sample on ice or in a refrigerator. Do not freeze sediment samples. The
sediment enzyme sample has a two week holding time.
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• SAMPLE COLLECTION FORM - BOA!
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Figure 5.1-4. Sample Collection Form, Side 2.
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5.2 Physical Habitat Characterization in Non-Wadeable Rivers and Streams
Physical habitat in rivers includes all those physical attributes that influence or provide
sustenance to river organisms. Physical habitat varies naturally; thus, expectations differ even in
the absence of anthropogenic disturbance. Within a given physiographic-climatic region, river
drainage area and channel gradient are likely to be strong natural determinants of many aspects
of river habitat, because of their influence on discharge, flood stage, and stream power (the
product of discharge times gradient). Kaufmann (1993) identified 7 physical habitat attributes
important in influencing stream ecology that are likely applicable in rivers as well. They include:
Channel Dimensions
Channel Gradient
Channel Substrate Size and Type
Habitat Complexity and Cover
Riparian Vegetation Cover and
Structure
Anthropogenic Alterations
Channel-Riparian Interaction
The protocol defines the length of each sampling reach proportional to river wetted width
and then systematically places measurements to statistically represent the entire reach. Stream
thalweg depth measurements, habitat classification, and mid-channel substrate observations
are made at very tightly spaced intervals; whereas channel "littoral" and riparian stations for
measuring or observing substrate, fish cover, large woody debris, bank characteristics and
riparian vegetation structure are spaced further apart. The tightly spaced depth measures allow
calculation of indices of channel structural complexity, objective classification of channel units
such as pools, and quantification of residual pool depth, pool volume, and total stream volume.
5.2.1 Equipment and Supplies
Table 5.2-1 lists the equipment and supplies required to conduct all the activities
described for characterizing physical habitat. This checklist is similar to the checklist presented
in Appendix A, which is used at the base location (Section 3) to ensure that all of the required
equipment is brought to the river. Use this checklist to ensure that equipment and supplies are
organized and available at the river site in order to conduct the activities efficiently.
Table 5.2-1. Checklist of equipment and supplies for physical habitat
For making
measurements
Surveyor's telescoping leveling rod (round profile, metric scale, 7.5m extended)
Convex spherical canopy densiometer (Lemmon Model B), modified with taped "V"
GPS
1 roll each colored surveyor's flagging tape (2 colors)
2 pair chest waders
1 or 2 fisherman's vest with lots of pockets and snap fittings.
Digital camera with extra memory card & battery
50 m or 100 m measuring tape with reel
Meter stick for bank angle measurements
SONAR unit
Laser rangefinder (400 ft. distance range) and clear waterproof bag
Clinometer
Binoculars
Field Operations Manual and/or laminated quick reference guide
Laminated invasive species guide
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For recording
data
2 covered clipboards (lightweight, with strap or lanyard)
Soft (#2) lead pencils
11 plus extras Channel/Riparian Transect Forms
11 plus extras Thalweg Profile Forms
1+ extras field Form: Stream Verification Form
1+ extras field Form: Field Measurement Form
1+ extras field Form: Sample Collection Form
1+ extras field Form: Riparian "Legacy" Trees and Invasive Alien Plants
1+ extras field Form: Channel Constraint
1+ extras field Form: Fish Gear and Voucher/Tissue Information Form
1+ extras field Form: Fish Collection Form
1+ extras field Form: Visual Assessment Form
5.2.2 Components of the Field Habitat Assessment
Field data collection for the physical habitat assessment is accomplished in a single float
down each sampling reach. River sample reach lengths are defined as 40 x the wetted width at
the x-site, with a minimum of 150m and maximum of 4km. To characterize mid-channel habitat
(Table 5.2.2), they measure a longitudinal thalweg (or mid-channel) depth profile, record the
presence of snags and off-channel habitats, classify main channel habitat types, characterize
mid-channel substrate, and locate the 11 transect locations for littoral/riparian sampling and
other habitat observations. At each of the 11 transects (A-K), they measure channel wetted
width, bankfull channel dimensions, incision, GPS lat/long, and then assess near-shore,
shoreline, and riparian physical habitat characteristics by measuring or observing littoral depths,
riparian canopy cover, substrate, large woody debris, fish cover, bank characteristics, riparian
vegetation structure, presence of large ("legacy") riparian trees, non-native riparian and aquatic
species, and evidence of human activities. After all the thalweg and littoral/riparian
measurements and observations are completed, the crews estimate the extent and type of
channel constraint.
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Table 5.2-2. Components of river physical habitat protocol
Thalweg Profile:
At 10 equally spaced intervals between each of 11 transects (100 along entire reach):
• Classify habitat type, record presence of backwater and off-channel habitats.
Determine dominant substrate visually or using sounding rod.
At 10 equally spaced intervals between each of 11 transects (100 along entire reach):
Record the presence of mid-channel snags
Measure thalweg (maximum) depth using Sonar or rod
Littoral/Riparian Cross-Sections: @ 11 transects at equal intervals along reach length:
Measure/estimate from one chosen bank on 11 transects :
Wetted width and Mid-channel bar width (laser range finder).
Bankfull width (laser) and height (pole and clinometer used as level).
Incision height (pole and clinometer used as level).
Bank angle (estimate)
Riparian canopy cover (densiometer) in four directions from chosen bank.
Shoreline Substrate in the first 1m above waterline (dominant and subdominant size class).
In 20m long Littoral Plot extending streamward 10m from chosen bank : 1
Littoral depth at 5 locations systematically-spaced within plot (Sonar or sounding rod).
Dominant and Subdominant substrate size class at 5 systematically-spaced locations (visual or
sounding rod).
Tally large woody debris in littoral plot and in bankfull channel by size and length class.
Areal cover class of fish concealment and other features, including:
filamentous algae overhanging vegetation aquatic macrophytes
undercut banks large woody debris boulders and rock ledges
brush/small woody debris live trees or roots artificial structures
In 20m long Riparian Plot extending 10m landward starting at bankfull margin-both sides
of river:1
Estimate areal cover class and type (e.g., woody) of riparian vegetation in Canopy, Mid-Layer,
and Ground Cover layers
Observe and record human activities and disturbances and their proximity to the channel.
Record species of alien (non-native) trees, shrubs, grasses visible within riparian plot.
Looking upstream and downstream from each Transect (both sides of river):
Look for largest visible tree within 100m from the water's edge or as far as you can see, if less:
Estimate diameter (Dbh), height, species, and distance from river edge.
For the whole sampling reach, after completing thalweg and littoral/riparian measurements:*
• Classify channel type and degree of constraint, identify features causing constraint,
estimate the percentage of constrained channel margin for the whole reach, and
estimate the bankfull and valley widths.
*Note: Boundaries for visual observations are estimated by eye.
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5.2.3 Summary of Workflow
Table 5.2-3 lists the activities performed at and between each transect for the physical
habitat characterization. The activities are performed along the chosen river bank and mid-
channel (thalweg profile).
Table 5.2-3. Summary of workflow—river physical habitat characterization
A. At the chosen bank on first transect (farthest upstream):
Read GPS Lat./Long. and record it in the Transect (Shoreline) space on the field form.
Move boat in a "loop" within 10 x 20 m littoral plot, measuring 5 littoral depths and probing substrate.
Estimate dominant and subdominant littoral substrate, based on probing the 5 locations.
Estimate areal cover of fish concealment features in 10 x 20 meter littoral plot.
Tally LWD within or partially within the 10 x 20 meter littoral plot.
Do densiometer measurements at bank (facing upstream, downstream, left, right).
Choose bank angle class, estimate bankfull height, width and channel incision. (Note that width and incision
estimates incorporate both left and right banks.).
Tally LWD entirely out of water but at least partially within the bankfull channel.
Estimate and record distance to riparian vegetation on the chosen bank.
Make visual riparian vegetation cover estimates for the 10 x 20 meter riparian plot on both sides of the
channel. (Riparian plot starts where perennial vegetation begins or at bankfull channel margin, whichever is
closest to the wetted river margin. The plot continues 10m back from the bankfull line).
Identify taxa, height, diameter at breast height (Dbh), and distance from riverbank of largest tree as far as
you can see confidently upstream and downstream within 100m of the wetted river margin.
From a regional listing, record alien invasive tree, shrub, or grass taxa within in the 10m x 20m riparian plots
on either side of the river.
Make visual human disturbance tally on both sides of the river. Use the same plot dimensions as for riparian
vegetation ~ except that if a disturbance item is observed in the river or within the bankfull channel, the
proximity code is "B", the closest rating; "C" if within the riparian plot. If the item is only observed beyond
(outside) the riparian plot, the proximity code is "P".
Get out far enough from the bank so you can see downstream. Then use the laser rangefinder to sight and
record the distance to the intended position of the next downstream transect.
B. Thalweg Profile:
As soon as you get out from the bank after doing transect activities, take the first of 10 thalweg depth
measurements and substrate/snag probes using sonar and pole - also classify habitat type and record
presence of side-channels and backwaters.
Estimate thalweg measurement distance increments using the GPS course-tracking and trip-meter
functions. Alternatively, estimate these distances by keeping track of boat lengths or channel-width
distances traversed; each one is 1/1 Oth the distance between transects (also one-half channel-width, which
can help you keep track of your downstream progress).
C. Repeat the Whole Process (for the remaining 10 transects and spaces in between).
D. Channel Constraint Assessment
After completing the Thalweg Profile and Littoral-Riparian measurements and observations at all 11
Transects, complete the classification and estimation of channel constraint type, frequency of contact
with constraining features, and the width ratio of bankfull channel divided by valley width. You may
wish to refer to the individual transect assessments of incision and constraint.
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5.2.4 Habitat Sampling Locations on the Study Reach
Measurements are made at two scales of resolution along the mid-channel length of the
reach; the results are later aggregated and expressed for the entire reach, a third level of
resolution (Figure 5.2-1). Section 4 describes the procedures for locating the X-site, or the
midpoint of the sample reach. This sampling location is marked on the maps provided to the
field crews in the site dossiers prior to sampling. Sections 4.2 and 5.2.3 describe the protocol for
delineating a sample reach that is 40 times its width. Those sections also describe the protocol
for measuring out (with a laser range finder or GIS software) and locating the 11 littoral/riparian
stations where many habitat measurements will be made (Figure 5.2-3). The distance between
each of these transects is 1/1 Oth the total length of the sample reach.
The thalweg profile measurements are spaced as evenly as practicable over the entire
sample reach length. In addition, they must be sufficiently close together to not "miss" deep
areas and habitat units that are in a size range of about 1/3 to 1/2 of the average channel width.
To set the interval between thalweg profile measurements, measure the wetted channel width
with a laser rangefinder at 5 locations near the X-site and multiply the average width by 40 to
set the river sample reach length. Then divide that reach length by 100 to set the thalweg
increment distance. Following these guidelines, you will be making 100 evenly-spaced thalweg
profile measurements, 10 between each detailed channel cross-section where littoral/riparian
observations are made. If the thalweg is too deep or not physically possible to be measured to,
estimate the depth to the best of your ability and flag it on the field form.
UPSTREAM END
River Flow
Thalweg
Profile
Increments
DOWNSTREAM END
Figure 5.2-1. River reach sample layout.
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LEFT
UPSTREAM END
River Flow
aggilsSiw&i
Riparian
Plot
._~— ~
^TZ^
10m
• RIGHT
I BANK
20 rn
Thalweg
Profile
Increments
DOWNSTREAM END
Figure 5.2-2. Littoral-Riparian Plots for characterizing riparian vegetation, human influences,
fish cover, littoral substrate, and littoral depths.
5.2.5 Work Flow and Reach Marking
After finding adequate put-in and take-out locations, the team may opt to mark the
upstream end of the sample reach end with colored flagging. In a single midstream float down
the 40 channel-width reach, the 2-person habitat team accomplishes a reconnaissance, a
sonar/pole depth profile, and a pole-drag to tally snags and characterize mid-channel substrate.
The float is interrupted by stops at 11 transect locations for littoral/riparian observations. They
determine (and mark - optional, but recommended) the intended position of each successive
downstream transect using a global positioning system (GPS) or a laser range finder. Each
transect is located 4 channel-width's distance from the preceding transect immediately
upstream. The crew then floats downstream along the thalweg to the new transect location,
making thalweg profile measurements and observations at 10 evenly-spaced increments along
the way. When they reach the new downstream transect location, they stop to do cross-section,
littoral, and riparian measurements, recording the actual GPS latitude/longitude of the transect
position. In addition, while they are stopped at a cross-section station, the crew can fill out the
habitat "typing" entries retrospectively and prospectively for the portion of the stream distance
that is visible up- and downstream. They will also collect biological and sediment samples.
GPS coordinates are determined for the actual locations of each transect stop. If GPS
unit also has course tracking, trip-meter (accumulated distance and bearing), and
waypoint setting/navigation features, we recommend using it to locate thalweg measurement
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points (use course tracking and trip meter). Equipping the boat with a bow or stern anchor to
stop at transect locations can greatly ease the shore marking operation and shoreline
measurement activities, though such equipment can be dangerous in white-water rivers.
5.2.6 Reconnaissance
The habitat crew will also record reconnaissance and safety notes at this time. They will
inform the second boat of the route, craft, and safety precautions needed during its subsequent
electrofishing activities. They also assist the electrofishing boat crew over jams and help to
conduct shuttles (this can take considerable time where put-ins and take-outs are distant). As
the team floats downstream, they may choose and communicate to the electrofishing crew the
most practical path to be used when fishing with a less maneuverable boat, taking into
consideration multiple channels, blind channels, backwaters, alcoves, impassible riffles, rapids,
jams, and hazards such as dams, bridges and power lines. They determine if and where
tracking or portages are necessary.
5.2.7 Thalweg Profile
"Thalweg" refers to the flow path of the deepest water in a river channel. The thalweg
profile is a longitudinal survey of maximum depth and several other selected characteristics at
100 near-equally spaced points along the centerline of the river between the two ends of the
river reach (Figure 5.2-1). For practical reasons, field crews will approximate a thalweg profile
by sounding along the river course that they judge is deepest, but also safely navigable.
Locations for observations and measurements along the path of this profile are
determined using the GPS course-tracking and trip-meter features (recommended), or by
visually estimating distances based upon the river width. Data from the thalweg profile allows
calculation of indices of residual pool volume, river size, channel complexity, and the relative
proportions of habitat types such as riffles and pools. The procedure for obtaining thalweg
profile measurements is presented in Table 5.2-2. Record data on the Thalweg Profile Form as
shown in Figure 5.2-3.
5.2.7.1 Thalweg Depth Profile
A thalweg depth profile of the entire 40 channel-width reach is approximated by a sonar
or sounding rod depth profile while floating downstream along the deepest part of the channel
(or closest navigable path). In the absence of a recording fathometer (sonar depth sounder with
strip-chart output or electronic data recorder), the crew records depths at frequent, relatively
evenly-spaced downstream intervals while observing a sonar display and holding a surveyor's
rod off the side of the boat (see Section 5.2.7.2). The sonar screen is mounted so that the
crewmember can read depths on the sonar and the rod at the same time. The sonar sensor
may need to be mounted at the opposite end of the boat to avoid mistaking the rod's echo for
the bottom, though using a narrow beam (16 degree) sonar transducer minimizes this problem.
It is easy to hold the sounding rod vertically if you are going at the same speed as the water. If
the thalweg is too deep to safely be recorded, estimate the depth and note on comments form.
5.2.7.2 Pole Drag for Snags and Substrate Characteristics
The procedure for dragging the thalweg pole to detect underwater snags and substrate
characteristics is presented in Table 5.2-4. While floating downstream, one crewmember holds
a calibrated PVC sounding rod or surveying rod down vertically from the gunwale of the boat,
dragging it lightly on the bottom to simultaneously "feel" the substrate, detect snags, and
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measure depth with the aid of sonar. The crewmember shall record the dominant substrate type
sensed by dragging the rod along the bottom (bedrock/hardpan, boulder, cobble, gravel, sand,
silt & finer) on the Thalweg Profile Form (Figure 5.2-3). Substrate characteristics are recorded at
every thalweg depth measurement (e.g., 10 determinations between transects A and B). In
shallow, fast-water situations, where pole-dragging might be hazardous, crews will estimate
bottom conditions the best they can visually and by using paddles and oars. If unavoidable,
suspend measurements until out of Whitewater situations, but make notes and appropriately flag
observations concerning your best judgments of depth and substrate.
Table 5.2-4. Thalweg profile procedure
1. Determine the interval between transects based on the mean wetted width used to determine the reach
length. Transects are at 4 channel-width spacings; thalweg depth, snags, off-channel habitats and other
downstream longitudinal profile observations are recorded at intervals of 0.4 channel-width.
2. Complete header information on the Thalweg Profile Form, noting transect pair (up- to downstream).
3. Begin at the upstream transect (station "1" of "10"). Determine the locations to take measurements using
the course-tracking and trip-meter functions of the GPS. Alternatively, estimate your position.
Thalweg Depth Profile
a) While floating downstream along the thalweg, record depths at frequent, even-spaced intervals while
observing a sonar display and holding a surveyor's rod off the side of the boat.
b) A depth recording every 0.4 channel-width distance is required, yielding 10 measurements between
channel/riparian cross-section transects.
c) If the depth is >0.5 meters, or contains a lot of air bubbles, the sonar fathometer will not give reliable
depth estimates. In this case, record depths using a calibrated sounding rod. In shallow, fast-water
situations depths may have to be visually estimated to the nearest 0.5 m.
d) Measure depths to nearest 0.1 m and record in the "SONAR" or "POLE" column.
Pole Drag for Snags and Substrate Characteristics
From the gunwale of the boat, hold a surveying rod or calibrated PVC sounding rod down vertically into the
water. (CAUTION: Hold the rod over the side or stern of the raft; otherwise it could be jerked out of your
hands if it catches on an obstruction in fast water.)
Lightly drag the rod on the river bottom to "feel" the substrate and detect snags.
Record the presence of snags hit by the rod or seen visually, plus the dominant substrate type sensed by
dragging the rod along the bottom.
Circle the appropriate "SUBSTRATE" type and record the presence/absence of "SNAGS".
If it is too deep to safely measure the substrate type, estimate the type based on knowledge and
surrounding measurements and flag the date.
Channel Habitat Classification
Classify and record the channel habitat type at increments of every 0.4 channel width.
Check for off-channel and backwater habitat at increments of every 0.4 channel width.
If channel is split by a bar or island, navigate and survey the channel with the most flow.
When a side channel is encountered, circle "Y" in the "OFF-CHANNEL" column beginning with the point of
divergence from the main channel, continuing downriver until the side channel converges with the main
channel.
Circle the "CHANNEL HABITAT" and record side channels as described in (d) above.
Proceed downriver to the next station, and repeat the above procedures.
Record GPS waypoint (Lat/Long) midstream and at shoreline location on each transect in decimal degrees.
Repeat the above procedures until you reach next transect. Set a waypoint location for the transect location
midstream and at the adjacent bank. Record waypoints that you set for channel bends, transect mid-stream,
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and transect shoreline locations on the Channel-Riparian Transect Form corresponding to the downstream
end of the thalweg sub-reach you just traversed.
After completing activities at the shoreline, prepare a new Thalweg Profile Form, then repeat the above
procedures for each of the reach segments, until you reach the downriver end of the reach (Transect "K").
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PHAB: THALWEG PROFILE FORIVI - BOATABLE
^wed i ,
hmirgil Uf I
SITE ID: FVV08
DATE;
/ rt / / 2 0
TRANSECT: ft A-B . B-C " C-0 0-E O E-F 0 F-G 0 G-H H-l C I-J 0 J-K
SUBSTRATE CODES
BH = BESROCKiKASOPAiH jSMOOTH OR SOUSH^ - (LARGER THAN A CARj
81 = BOULPfeR (250 1O 40GO mm) - BASKETS ALL TO C AK]f
CB = COBStg |§4 lO2»nim| * (fENNSS BAit TO BAS-KETBALLJ
GR = COASSC TO RN£ GRAVEL (2 1Q 54 mm.\ - UAPYBllG IO ?E*«NIS BALL]
SA a SAJWD ^ M TO 2 mm! - (GRITTY . UP TO LADVBUO SHE)
F"N * Sttli CLAY / MUCK - ^OT GSEITYi
Of *> OTHER ICOMMENT ON OTHfcS SiD£i
CHANNEL HABITAT CODES
si
RI
Dry
OTHER
Off Channel = Off
REMEMBER; A ss Upsimam end ol Reach and K = Downstream sod of Weach.
TNALWEG PROFIiE
Figure 5.2-3. Thalweg Profile Form.
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5.2.7.3 Channel Habitat Classification
Classify and record channel habitat types shown in Table 5.2-5 at a spatial resolution of
about 0.5 channel-widths and check presence of off-channel and backwater habitat at every 0.4
channel-width increment. The procedures for classifying channel habitat are presented in Table
5.2-2. Designate side channels, backwaters and other off-channel areas independent of the
main-channel habitat type. Main channel habitat units are at least half as long as the channel is
wide, (e.g., if there is a small, deep, pool-like area at the thalweg within a large riffle area, don't
record it as a pool unless it occupies an area about half as wide or long as the channel is wide).
Table 5.2-5 Channel unit categories
Class (Code)3 Description
Pools (PO): Still water, low velocity, smooth, surface, deep compared to other parts of channel
Glide (GL) Water moving slowly, with a smooth, unbroken surface. Low turbulence.
Riffle (Rl) Water moving, with small ripples, waves and eddies—waves not breaking, surface
tension not broken. Sound: "babbling", "gurgling".
Rapid (RA) Water movement rapid and turbulent, surface with intermittent Whitewater with
breaking waves. Sound: continuous rushing, but not as loud as cascade.
Cascade (CA) Water movement rapid & very turbulent over steep channel bottom. Most of the water
surface is broken in short, irregular plunges, mostly Whitewater. Sound: roaring.
Falls (FA) Free falling water over vertical or near vertical drop into plunge, water turbulent and
white over high falls. Sound: splash to roar. (Do not navigate raft over a waterfall!).
Dry channel (DR) No water in the channel.
Off-channel Side-channels, sloughs, backwaters, and alcoves separated from the main channel.
a In order for a channel habitat unit to be distinguished, it must be at least half as wide or long as the channel is wide.
Mid-channel bars, islands, and side channels within a thalweg profile require some
guidance. Mid-channel bars are defined as channel features below the bankfull flow level that
are dry during baseflow conditions (Section 5.2.8.3 defines bankfull channel). Islands are
channel features that are dry even when the river is at bankfull flow. If a mid-channel feature is
as high as the surrounding flood plain, it is considered an island. Both mid-channel bars and
islands cause the river to split into side channels. If a bar or island is encountered along the
thalweg profile, navigate and survey the channel that carries the most flow. Note side channels
are present but do not sample them.
When side channels are present, on the Thalweg Profile form check the "Off-Channel"
column. These checkmarks will begin at the point of divergence from the main channel,
continuing downstream to the point of convergence with the main channel. In the case of a
slough or alcove, the "off-channel" checkmarks should continue from the point of divergence
downstream to where the off-channel feature is no longer evident. When major side channels
occur, flag the "Off-Channel" checkmarks and indicate in the comments section that the feature
is a side channel. For dry and intermittent rivers, record zeros for depth and wetted width in
places where no water is in the channel. Record habitat type as dry channel (DR).
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5.2.8 Channel Margin ("Littoral") and Riparian Measurements
This section covers channel margin depth and substrate, large woody debris, bank angle,
channel cross-section morphology, canopy cover, riparian vegetation structure, fish cover, and human
influences. Record measurements on the Channel/Riparian Transect Form (Figures 5.2-4 and 5.2-5).
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PHAB: CHANNEL/RIPARIAN TRANSECT FORM - BOATABLE (FRONT)
' '
SITE ID: FW08 tfX0C?O
TRANSECT: C A ' '
PI
DATE; ^ -
B C n D ng « F 0 G
Trzr .Y.
•LITTORAL
SHORE BOTTOM
DOM SEC DOM ' SFC
RS RS RS I RS
RR BK KR RR
XB XB XB XB
SB (SB^ SB (SB)
CB CB f£5) CB
(*5& GC GC OC
GF [ GF OF GF
SA SA SA SA
FN FN FN FN
HP HP HP HP
WD WD WD WO
OT | OT OT OT
E
CLASS CtJ
f & 1 t Z
?,/
<"••
7
Arrival Time Lsave Tim&
i-* r"i I fs i Oi t/ r^ v Chosen bank ^ide
* oft O HiuH
/ o, / ^7 J^ 3 J^
/ j^ r £»7 ¥ 5. /
' SUBSTRATF INFORMATION
OTTOM SUBSTRATA FROM ,X ONE) p( I j
udgement -of" • OBb ,i e Lfttoral Depths 3^ [ j
RS * Bedrock (Smooth) - (Larger than
•a c,*irf
RR = Bedrock ( Rough) - (Largsr than a earj
XB = Large Bouldor (1000 to 4000 mm
- ^Meterstick to carf
SB * Small Boylder J250 to 1000 mm) - (Basketball to Mcten.tick)
CB = Cobble (64 to 250 mm) - (Tennis ball to Basketball)
GC « Coarse Gravel (16 to 64 mm) - (Marble to Tennis ball)
GF - Fine Gravel (2 to 16 inm) - (Ladybuq to marble)
SA * Sand
0.06 to 2 mm) -{Gritty
* yp to Ladytiuy si/ej
FN = Silt; Clay' Muck (Not Gritty)
HP = Hartipan - (Firm, Consolidated Fine Substrate)
WD = Wood - lAny Sm>)
OT * Other (Write comment below)
LARGE WOODY OtBRIS ittiuftnnoi) TKirtACHnccc Flig 1 1
kV d Atl P <1 n Wasted CtwR
;„,„ """^
0 6 - 0 1 r
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f
* • ~' ir
^ IS m
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p
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-
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-
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BANK CHARACTERISTICS
X.XX (m) FLJ«3
Wetted Width ^jr* Q\
RM Wulili /j !
Bnnklull Width 7^"
Banhfull Height jj O
incised Height JK
BAMK
CIRCLE ONE \ ^«racal
i.^S
WTENO€Ofr.r««. ACfOSL!,™^
Y2fl 1 v?/5 I
ttd Riffl ss.s GPS L&n^ilud* - ddd mm s,s.a Tiag
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1 L ( • t t ( j
F Up Comments
k. /ve> TruA
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/**»Ct$ltt*s *>&T
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- .,, i. ,, ..
.' • « .u.,n™,,i. Draft
"•" * 1
Figure 5.2-4. Channel/Riparian Transect Form, page 1 (front side).
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PHAB: CHANNEL/RIPARIAN TRANSECT FORM - BOATABLE (Back) R..-d*,«.
SITE ID: FW08XXOOO
TRANSECT; • A OB O C
VISUAL RIPARIAN
ESTIMATES
R1PARIAW j
VfcGfcTAIfON COVFR .
W»0[|,V»,«,,,t«,t,^
SMALL Tre-es fTninfc
«,
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0 1
0 f
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00 OF
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n F ~< G ' H I
J ' Mssvy (« T5"fe M = Mixaii
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Canopy |>5 m hsyh)
E (M) N
(^ 3 4
") 2 3 4
Uf*d6FS!'Of¥
E M N
WuM> si';.^. o ' 20m Pint) ln Cha""e'^0¥fir f ,ag
rilnwnUM «s». (J) 1 2 3 4
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".oT^aiS ° ' © 3 4
BrU*^M°TdS«*U-1 ° 0 2 3 *
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°- .Tm o! L^ll 8 (p 2 3 4
UnJ.rvul8.nK 0 Q) 2 3 4
B~*"^.«(9« (J) 2 3 4
Art*,«St™,u™, ©1234
CHANNEL CONSTRAINT
DISTANCE FROM SHORE [ ^
TO RIP AR1AJJ VEGETATION ill) «» ^
CIRCLE ONE
C Ch.^...CM»Md
B Ch^iinsi •« in §r^a?a Vg'i^v {t*irt f»i3f*Sl/3'fiwJ £V (ftfit tn.
fS/ Chann®. •% in Nnrrnw Valley te^sl ^O! vwv cu^atrdssted
^"^
U chain* n Unrnrncwd in fitoitf VaMev
CHECK ONE
41 ¥ES t COULD ftfcAKLV SEE OVER THE BANK
O NO CO^LO HOf KiEADSLY SEE OVER THE BAWK
FLAG
Comn""!? c«K«OMSrry«.A»K
^^n*jr orNsiossETER ro TO vi IMX)
, W UM
UP 1 /^
DOWN
_J
$
KIOHT ^
-
'•''«' ' >
ruftG
v , * . ^Epwm* ^^
J-^l 1
Figure 5.2-5. Channel/Riparian Transect Form, page 2 (back side).
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5.2.8.1 Channel Margin Depth and Substrate
Channel margin depths are measured along the designated shoreline at each transect
within the 10m x 20m littoral plot that is centered on the transect. Dominant and sub-dominant
bottom substrates are determined and recorded at 5 systematically-spaced locations that are
located by eye within the 10m x 20m plot. The procedure for obtaining channel margin depth
and substrate measurements is described in more detail in Table 5.2-6. Record these
measurements on the Channel/Riparian Transect Form as shown in Figure 5.2-4. Identify the
dominant and subdominant substrate present along a shoreline swath 20 meters long and 1
meter back from the waterline. The substrate size class choices are as shown in Table 5.2-6.
Table 5.2-6. Channel margin depth and substrate procedure
1.
2.
3.
4.
5. f
Fill in the header information on page 1 of a Channel/Riparian Transect Form. Be sure to indicate the
letter designating the transect location.
Measure depth and observe bottom substrates within the 10m x 20 m littoral plot that is centered on
each transect location.
Determine and record the depth and the dominant and subdominant substrate size class at 5
systematically-spaced locations estimated by eye within this 1 0m x 20m plot and 1 m back from the
waterline. If the substrate particle is "artificial" (e.g. concrete, asphalt), choose the appropriate
size class, flag the observation and note that it is artificial in the comment space.
Code
RS
RR
XB
SB
CB
GC
GF
SA
FN
HP
WD
OT
Size Class
Bedrock (Smooth)
Bedrock (Rough)
Large Boulders
Small Boulders
Cobbles
Gravel (Coarse)
Gravel (Fine)
Sand
Fines
Hard pan
Wood
Other
Size Range (mm)
>4000
>4000
>1 000 to 4000
>250to 1000
>64 to 250
>16to64
> 2 to 16
>0.06to2
<0.06
Regardless of Size
Regardless of Size
Description
Smooth surface rock bigger than a car
Rough surface rock bigger than a car
Meter stick to Car size
Basketball to Meter stick size
Tennis ball to basketball size
Marble to tennis ball size
Ladybug to marble size
Gritty - up to ladybug size,
Silt Clay Muck (not gritty between fingers)
Firm, consolidated fine substrate
Wood & other organic particles
Concrete, metal, tires, etc. (note in comments)
On page 1 of the Channel/Riparian Transect Form, circle the appropriate shore and bottom substrate
type and record the depth measurements ("SONAR" or "POLE" columns).
Repeat Steps 1 through 4 at each new cross-section transect.
5.2.8.2 Large Woody Debris
Large Woody Debris (LWD) is defined as woody material with small end diameter of >30
cm (1ft) and length of >5 m (15 ft). These size criteria are larger than those used in wadeable
streams because of the lesser role that small wood plays in controlling velocity and morphology
of larger rivers. The procedure for tallying LWD is presented in Table 5.2-7. For each tally
(Wood All/Part in Wetted Channel and Dry but All/Part in Bankfull Channel), the field form
(Figure 5.2-4) provides 12 entry boxes for tallying debris pieces visually estimated within three
length and four diameter class combinations. Tally each LWD piece in only one box. Do not tally
woody debris in the area between channel cross-sections, but the presence and location of
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large debris dams and accumulations should be mapped (sketched) and noted in the thalweg
profile comments.
For each LWD piece, first visually estimate its length and its large and small end
diameters and place it in one of the diameter and length categories. The diameter classes on
the field form (Figure 5.2-4) refer to the large end diameter. Sometimes LWD is not cylindrical,
so it has no clear "diameter". In these cases visually estimate what the diameter would be for a
piece of wood with circular cross-section that would have the same volume. When evaluating
length, include only the part of the LWD piece that has a diameter >0.3m (1 ft). Count each of
the LWD pieces as one tally entry and include the whole piece when assessing dimensions,
even if part of it is outside of the bankfull channel. If you encounter massive, complex debris
jams, estimate their length, width, and height. Estimate the diameter and length of large "key"
pieces and judge the average diameter and length of the other pieces making up the jam.
Record this information in the comments section of the form.
Table 5.2-7. Procedure for tallying large woody debris
Note: Tally pieces of large woody debris (L WD) within the 11 transects of the river reach at the same time
the shoreline measurements are being determined. Include all pieces whose large end is located within
the transect plot in the tally. Tally wood that is at least partially within the wetted channel separately from
that that is not presently wetted, but still within or directly above (bridging) the bankfull channel
1. LWD is tallied in 11 "plots" systematically spaced over the entire length of the stream reach. These
plots are each 20 m long in the upstream-downstream direction (10m up, 10m down). They are
positioned along the chosen bank and extend from the shore in 10m towards mid-channel and then all
the way to the bankfull margin.
2. Tally all LWD pieces within the plot that are at least partially within the presently wetted (baseflow)
channel. First, determine if a piece is large enough to be classified as LWD (small end diameter 30
cm [1ft.]; length 5m [15 ft.])
3. For each piece of LWD, determine its diameter class based on the diameter of the large end (0.3 m
to < 0.6 m, 0.6 m to <0.8 m, 0.8 m to <1.0 m, or >1.0 m), and the length class of the LWD pieces
based on the part of its length that has diameter >30 cm. Length classes are 5m to <15m, 15m to
<30m, or>30m.
• If the piece is not cylindrical, visually estimate what the diameter would be fora piece of wood
with circular cross-section that would have the same volume.
• When estimating length, include only the part of the LWD piece that has a diameter >0.3 m (1 ft.)
4. Place a tally mark in the appropriate diameter x length class tally box in the "WOOD ALL/PART IN
WETTED CHANNEL" section of the Channel/Riparian Transect Form.
5. Tally all shoreline LWD pieces along the littoral plot that are at least partially within or above (bridging)
the bankfull channel, but not in the wetted channel. For each piece, determine the diameter class based
on the diameter of the large end (0.3 m to < 0.6 m, 0.6 m to <0.8 m, 0.8 m to <1.0 m, or >1.0 m), and
the length class based on the length of the piece that has diameter >30 cm. Length classes are
5m to <15m, 15m to <30m, or >30m.
6. Place a tally mark for each piece in the appropriate diameter x length class tally box in the "DRY BUT
ALL/PART IN BANKFULL CHANNEL" section of the Channel/Riparian Transect Form.
7. After all pieces within the segment have been tallied, write the total number of pieces for each diameter
x length class in the small box at the lower right-hand corner of each tally box.
8. Repeat Steps 1 through 7 for the next river transect, using a new Channel/Riparian Transect Form.
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5.2.8.3 Bank Angle and Channel Cross-Section Morphology
Bank angles of undercut, vertical, steep, and gradual are visually estimated as defined
on the field form (Figure 5.2-4). Observations are made from the wetted channel margin up 5 m
(a canoe's length) into the bankfull channel margin on the previously chosen side of the stream.
You will measure or estimate the wetted width, mid-channel bar width, bankfull height
and width, the amount of incision, and the degree of channel constraint. These are assessed for
the whole channel (left and right banks) at each of the 11 cross-section transects. Record
each on the Channel/Riparian Transect Form (Figure 5.2-4). The procedures for obtaining bank
angle and measurements of channel cross-section morphology are presented in Table 5.2-8.
Wetted width is the width of the channel containing free-standing water; if >15 m, it can
be measured with a laser rangefinder. Mid-channel bar width, the width of exposed mid-
channel gravel or sand bars, is included within the wetted width, but is also recorded separately.
In channel cross-section measurements, the wetted and bankfull channel boundaries include
mid-channel bars. Therefore, the wetted width is measured as the distance between wetted left
and right banks. Measure across and over mid-channel bars and boulders. If islands are
present, treat them like bars, but flag these measurements and indicate in the comments that
the "bar" is an island. If you are unable to see across the full width of the river when an island
separates a side channel from the main channel, record the width of the main channel, flag the
observation, and note in the comments section that the width pertains only to the main channel.
Table 5.2-8. Procedure for bank angle and channel cross-section
1. Visually estimate the bank angle (undercut, vertical, steep, gradual), as defined on the field form.
Bank angle observations refer to the area from the wetted channel margin up 5 m (canoe's length)
into the bankfull channel margin on the previously chosen side of the river. Circle the angle in the
"BANK ANGLES" section of the Channel/Riparian Transect Form.
2. Hold the surveyor's rod vertically, with its base planted at the water's edge. Examine both banks, then
determine the channel incision as the height up from the water surface to elevation of the first
terrace of the valley floodplain (Note this is at or above the bankfull channel height). Whenever
possible, use the clinometer as a level (positioned so it reads 0% slope) to measure this height by
transferring (backsighting) it onto the surveyor's rod. Record this value in the INCISED HEIGHT field of
the bank characteristics section on the field data form.
3. While still holding the surveyor's rod as a guide, and sighting with the clinometer as a level, examine
both banks to measure and record the height of bankfull flow above the present water level. Look for
evidence on one or both banks such as:
• An obvious slope break that differentiates the channel from a relatively flat floodplain terrace
higher than the channel.
• A transition from exposed stream sediments to terrestrial vegetation.
• Moss growth on rocks along the banks.
• Presence of drift material caught on overhanging vegetation.
• A transition from flood- and scour-tolerant vegetation to that which is relatively intolerant of these
conditions.
4. Record the wetted width value determined when locating substrate sampling points in the BANK
CHARACTERISTICS section of the field data form. Also determine the bankfull channel width and the
width of exposed mid-channel bars (if present).
5. Repeat Steps 1 through 6 at each cross-section transect, (including any additional side channel
transects established when islands are present). Record data for each transect on a separate field
data form.
Bankfull flows are large enough to erode the stream bottom and banks, but frequent
enough (every 1 to 2 years) to not allow substantial growth of upland terrestrial vegetation.
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Consequently, in many regions, it is these flows that have determined the width and depth of the
channel. Estimates of the bankfull dimensions of stream channels are extremely important in
EMAP surveys. They are used to calculate shear stress and bed stability (see Kaufmann et al.,
1999). Unfortunately, we have to depend upon evidence visible during the low-flow sampling
season. If available, consult published rating curves relating expected bankfull channel
dimensions to stream drainage area within the region of interest. Graphs of these rating curves
can help you get a rough idea of where to look for field evidence to determine the level of
bankfull flows. Curves such as these are available from the USGS for streams in most regions
of the U.S. (e.g., Dunne and Leopold 1978; Harrelson et al. 1994, Leopold 1994). To use them,
you need to know the contributing drainage area to your sample site. Interpret the expected
bankfull levels from these curves as a height above the streambed in a riffle, but remember that
your field measurement will be a height above the present water surface of the stream. Useful
resources to aid your determination of bankfull flow levels in streams in the United States are
video presentations produced by the USDA Forest Service for western streams (USDA Forest
Service 1995) and eastern streams (USDA Forest Service 2002).
After consulting rating curves that show where to expect bankfull levels in a given size of
stream, estimate the bankfull flow level by looking at the following indicators:
• First look at the stream and its valley to determine the active floodplain. This is a
depositional surface that frequently is flooded and experiences sediment deposition
under the current climate and hydrological regime.
• Then look specifically for:
• An obvious break in the slope of the banks.
• A change from water-loving and scour-tolerant vegetation to more drought-tolerant
vegetation.
• A change from well-sorted stream sediments to unsorted soil materials.
In the absence of clear bankfull indications, consider the previous season's flooding as the best
evidence available (note: you could be wrong if very large floods or prolonged droughts have
occurred in recent years.). Look for:
• Drift debris ("sticky wickets" left by the previous seasons flooding).
• The level where deciduous leaf-fall is absent on the ground (carried away by
previous winter flooding).
• Unvegetated sand, gravel or mud deposits from previous year's flooding.
In years that have experienced large floods, drift material and other recent high flow
markers may be much higher than other bankfull indicators. In such cases, base your
determination on less-transient indicators such as channel form, perennial vegetation, and
depositional features. In these cases, flag your data entry and also record the height of drift
material in the comments section of the field data form.
We use the vertical distance (height) from the observed water surface up to the level of
the first major valley depositional surface (Figure 5.2-6) as a measure of the degree of incision
or downcutting of the stream below the general level of its valley. This value is recorded in the
incised height field. It may not be evident at the time of sampling whether the channel is
downcutting, stable, or aggrading (raising its bed by depositing sediment). However, by
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recording incision heights measured in this way and monitoring them over time, we will be able
to tell if streams are incising or aggrading.
If the channel is not greatly incised, bankfull channel height and incision height will be
the same. However, if the channel is incised greatly, the bankfull level will be below the level of
the first terrace of the valley floodplain, making "Bankfull Height" smaller than "Incision" (Figure
5.2-6). Bankfull height is never greater than incision height. Look for evidence of recent
flows (within about 1 year) to distinguish bankfull and incision heights, though recent flooding of
extraordinary magnitude may be misleading. In cases where the channel is cutting a valley
sideslope and has oversteepened and destabilized that slope, the bare "cutbank" against the
steep hillside at the edge of the valley is not necessarily an indication of recent incision. In such
a case, the opposite bank may be lower, with a more obvious terrace above bankfull height;
choose that bank for your measurement of incised height. Examine both banks to determine
incision height and bankfull height. Remember that incision height is measured as vertical
distance to the first terrace above bankfull; if terrace heights differ on left and right
banks, choose the lower of the two terraces. Even when quite constrained by their valley
sideslopes, large rivers often have flood terraces above bankfull height. In some cases, though,
your sample reach may be in a steep "V" shaped valley or gorge formed over eons, and the
slopes of the channel banks simply extend uphill indefinitely, not reaching a terrace before
reaching the top of a ridge. In such cases, record incision height values equal to bankfull values
and make appropriate comments that no terrace is evident. Similarly, when the river is
extremely incised below an ancient terrace or plateau,(e.g., the Colorado River in the Grand
Canyon), you may crudely estimate the terrace height if it is the first one above bankfull level. If
you cannot estimate the terrace height, make appropriate comments describing the situation.
Finally, assess the local degree of river channel constraint (i.e., at the transect) by
following the guidelines on the form (Figure 5.2-5) regarding the relationships among channel
incision, valley sideslope, and width of the valley floodplain. You will also do an overall
assessment of channel constraint for the whole river reach; see Section 5.2.9 for a discussion of
constraint concepts and assessment procedures.
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A. Channel not Incised
Downcutting over
geologic time
Active
floodplain at or near
valley bottom elevation
(Record this height)
First terrace on
valley bottom
above bankfull
level
Second
terrace
No recent incision- bankfull
level at valley bottom
Valley Fill
B. Incised Channel
Downcutting over
geologic time
Former second
terrace becomes
Former active floodplain Former first third terrace
no longer connected— terrace becomes
becomes new first terrace second terrace
above bankfull level
(Record this height),
Recent incision-
bankfull level below
first terrace of valley
bottom
Valley Fill
Figure 5.2-6. Schematic showing bankfull channel and incision for channels. (A) not recently
incised, and (B) recently incised into valley bottom. Note level of bankfull stage relative to elevation of first
terrace on valley bottom (stick figure included for scale)
5.2.8.4 Canopy Cover (Densiometer)
Measure vegetative cover over the reach at the chosen bank at each of the 11 transects
(A-K). with a Convex Spherical Densiometer. Tape the densitometer exactly as shown in Figure
5.2-7 to limit the number of grid intersections to 17. Densiometer readings can range from 0 (no
canopy cover) to 17 (maximum canopy cover). Four measurements are obtained at each cross-
section transect (upriver, downriver, left, and right). The procedure for obtaining canopy cover
data is presented in Table 5.2-8. Record the counts in the "Canopy Density @ Bank" section of
the Channel/Riparian Transect Form as shown in Figure 5.2-4.
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TAPE
BUBBLE LEVELED-
Figure 5.2-7. Schematic of modified convex spherical canopy densiometer (From Mulvey et al.,
1992). In this example, 10 of the 17 intersections show canopy cover, giving a densiometer reading of 10.
Note proper positioning with the bubble leveled and face reflected at the apex of the "V."
Table 5.2-9. Procedure for canopy cover measurements
1. Take densiometer readings at a cross-section transect while anchored or tied up at the river margin.
2. Hold the densiometer 0.3 m (1 ft) above the surface of the river. Holding the densiometer level using
the bubble level, move it in front of you so your face is just below the apex of the taped "V".
3. At the channel margin measurement locations, count the number of grid intersection points within the
"V" that are covered by either a tree, a leaf, a high branch, or the bank itself.
4. Take 1 reading each facing upstream (UP), downstream (DOWN), left bank (LEFT), and right bank
(RIGHT). Right and left banks are defined with reference to an observer facing downstream.
5. Record the UP, DOWN, LEFT, and RIGHT values (0 to 17) in the "CANOPY COVER @ BANK"
section of the Channel/Riparian Transect Form.
6. Repeat Steps 1 through 5 at each cross-section transect. Record data for each transect on a separate
field data form.
5.2.8.5 Riparian Vegetation Structure
Riparian vegetation observations apply to the riparian area upstream 10m and
downstream 10m from each of the 11 transects. They include the visible area from the river
bankfull margin back a distance of 10 m (30 ft) shoreward from both the left and right banks,
creating a 10m X 20m riparian plot on each side of the river (Figure 5.2-2). The riparian plot
dimensions are estimated, not measured. Table 5.2-9 presents the procedure for characterizing
riparian vegetation structure and composition. Figure 5.2-5 illustrates how measurement data
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are recorded in the "Visual Riparian Estimates" section of the Channel/Riparian Transect Form,
side 2.
Table 5.2-10. Procedure for characterizing riparian vegetation structure
1. Anchor or tie up at the river margin at a cross-section transect; then make the following observations
to characterize riparian vegetation structure.
2. Estimate the distance from the shore to the edge of the riparian vegetation plot; record it just below
the title "Channel Constraint" on the Channel/Riparian Transect Form, side 2.
3. Facing the left bank (left as you face downstream), estimate a distance of 10 m back into the riparian
vegetation, beginning at the bankfull channel margin. Estimate the cover and structure of riparian
vegetation within an estimated 10 m x 20 m plot centered on the transect, and starting where
perennial vegetation begins or at the bankfull river margin (whichever is closest to the river
shoreline). On steeply-sloping channel margins, estimate the riparian plot dimensions as if they were
projected down from an aerial view.
4. Within this 10 m x 20 m area, conceptually divide the riparian vegetation into 3 layers: a CANOPY
(>5m high), an UNDERSTORY (0.5 to 5 m high), and a GROUND COVER layer (<0.5 m high).
5. Within this 10 m x 20 m area, determine the dominant woody vegetation type for the CANOPY
LAYER (vegetation > 5 m high) as either Deciduous, Coniferous, broadleaf Evergreen, Mixed, or
None. Consider the layer "Mixed" if more than 10% of the areal coverage is made up of the alternate
vegetation type. If the dominant vegetation type in the canopy layer is not woody, record the
vegetation type as "hJone". Indicate the appropriate vegetation type in the "VISUAL RIPARIAN
ESTIMATES" section of the Channel/Riparian Cross-section and Thalweg Profile Form.
6. Determine separately the areal cover class of large trees (> 0.3 m [1 ft] diameter at breast height
[DBH]) and small trees (< 0.3 m DBH) within the canopy layer. Estimate areal cover as the amount of
shadow that would be cast by a particular layer alone if the sun were directly overhead. Record the
appropriate cover class on the field data form ("0" = absent, zero cover; "1" = sparse, <10%; "2" =
moderate, 10-40%; "3" = heavy, 40-75%; or "4" = very heavy, >75%).
7. Look at the UNDERSTORY layer (vegetation between 0.5 and 5 m high). Determine the dominant
woody vegetation type for the understory layer as described in Step 5 for the canopy layer. If the
dominant vegetation type in the understory is not woody (e.g., herbaceous), record the vegetation
type as "hJone".
8. Determine the areal cover class for woody shrubs and saplings separately from non-woody
vegetation within the understory, as described in Step 6 for the canopy layer.
9. Look at the GROUND COVER layer (vegetation < 0.5 m high). Determine the areal cover class for
woody shrubs and seedlings, non-woody vegetation, and the amount of bare ground or duff (dead
organic material) present as described in Step 6 for large canopy trees.
10. Repeat Steps 1-9 for all transects, using a separate field data form for each transect.
You will estimate the areal cover separately in each of the three vegetation layers. Note
that the areal cover can be thought of as the amount of shadow cast by a particular layer alone
when the sun is directly overhead. The maximum cover in each layer is 100%, so the sum of the
areal covers for the combined three layers could add up to 300%. When rating vegetation cover
types, mixtures of two or more subdominant classes might all be given sparse ("1") moderate
("2") or heavy ("3") rankings. One very heavy cover class with no clear subdominant class might
be ranked "4" with all the remaining classes either moderate ("2"), sparse ("1") or absent ("0").
Two heavy classes with 40-75% cover can both be ranked "3".
5.2.8.6 Fish Cover, Algae, Aquatic Macrophytes
Over a defined length and distance from shore at the sampling locations, crews shall
estimate by eye and by sounding the proportional cover of fish cover features and trophic level
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indicators including large woody debris, rootwads and snags, brush, live trees in the wetted
channel, undercut banks, overhanging vegetation, rock ledges, aquatic macrophytes,
filamentous algae, and artificial structures.
The procedure to estimate the types and amounts of fish cover is outlined in Table 5.2-
10. Record data in the "Fish Cover/Other" section of the Channel/Riparian Transect Form as
shown in Figure 5.2-5. Crews will estimate the areal cover of all of the fish cover and other listed
features that are in the water and on the banks within the 10m x 20m plot (refer to Figure 5.2-2).
Table 5.2.11. Procedure for estimating fish cover
1. Stop at the designated shoreline at a cross-section transect and estimate a 10 m distance upstream
and downstream (20 m total length), and a 10 m distance out from the banks to define a 20 m x 10 m
littoral plot.
2. Examine the water and the banks within the 20 m x 10 m littoral plot for the following features and
types offish cover: filamentous algae, aquatic macrophytes, large woody debris, in-channel live trees
or roots, brush and small woody debris, overhanging vegetation, undercut banks, boulders, and
artificial structures.
3. For each cover type, estimate its areal cover by eye and/or by sounding with a pole. Record the
appropriate cover class in the "FISH COVER/OTHER" section of the Channel/Riparian Transect Form
("0"=absent: zero cover, "1"=sparse: <10%, "2"=moderate: 10-40%, "3"=heavy: 40-75%, or"4"=very
heavy: >75%).
4. Repeat Steps 1 through 3 at each cross-section transect, recording data from each transect on a
separate field data form.
Filamentous algae pertains to long streaming algae that often occur in slow moving
waters. Aquatic macrophytes are water loving plants in the river, including mosses, that could
provide cover for fish or macroinvertebrates. If the river channel contains live wetland grasses,
include these as macrophytes. Woody debris are the larger pieces of wood that can provide
cover and influence stream morphology (i.e., those pieces that would be included in the large
woody debris tally [Section 5.2.8.2]). Brush/woody debris pertains to the smaller wood that
primarily affects cover but not morphology. The entry for trees or brush within one meter of the
surface is the amount of brush, twigs, small debris etc. that is not in the water but is close to the
stream and provides cover. "Live Trees or Roots" are living trees that are within the channel -
estimate the areal cover provided by the parts of these trees or roots that are inundated. For
ephemeral channels, estimate the proportional cover of these trees that is inundated during
bankfull flows. Boulders are typically basketball to car sized particles. Many streams contain
artificial structures designed for fish habitat enhancement. Streams may also have in-channel
structures discarded (e.g., cars or tires) or purposefully placed for diversion, impoundment,
channel stabilization, or other purposes. Record the cover of these structures on the form.
5.2.8.7 Human Influences
For the left and right banks at each of the 11 detailed Channel/Riparian Cross-Sections,
evaluate the presence/absence and the proximity of 11 categories of human influences outlined
in Table 5.2-11. Record human influences on the Channel/Riparian Transect Form (Figure 5.2-
5). You may mark "P" more than once for the same human influence observed outside of more
than one riparian observation plot (e.g. at both Transect D and E). The rule is that you count
human disturbance items as often as you see them, BUT NOT IF you have to site through
a previously counted transect or its 10x20 meter riparian plot.
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Table 5.2-12. Procedure for estimating human influence
1. Stop at the designated shoreline at a cross-section transect, look toward the left bank (left when facing
downstream), and estimate a 10m distance upstream and downstream (20 m total length). Also,
estimate a distance of 10 m back into the riparian zone to define a riparian plot area.
2. Examine the channel, bank and riparian plot area adjacent to the defined river segment for the following
human influences: (1) walls, dikes, revetments, riprap, &dams; (2) buildings; (3) cleared lot, pavement
(e.g., paved, graveled, dirt parking lot, foundation); (4) roads or railroads, (5) inlet or outlet pipes; (6)
landfills or trash (e.g., cans, bottles, trash heaps); (7) parks or maintained lawns; (8) row crops; (9)
pastures, rangeland, or hay fields; (10) logging; and (11) mining (include gravel mining).
3. For each type of influence, determine if it is present and what its proximity is to the river and riparian plot
area. Consider human disturbance items as present if you can see them from the cross-section transect.
Do not include them if you have to site through another transect or its 10 m x 20 m riparian plot.
4. For each type of influence, record the proximity class in the "HUMAN INFLUENCE" part of the "VISUAL
RIPARIAN ESTIMATES" section of the Channel/Riparian Transect Form. Proximity classes are:
•B ("Bank") Present within the defined 20 m river segment and located in the stream or on the
wetted or bankfull bank.
•C ("Close") Present within the 10 x 20 m riparian plot area, but above the bankfull level.
•P ("Present") Present, but observed outside the riparian plot area.
•O ("Absent") Not present within or adjacent to the 20 m river segment or the riparian plot area
at the transect
5. Repeat Steps 1 through 4 for the opposite bank.
6. Repeat Steps 1 through 5 for each cross-section transect, recording data for each transect on a
separate field form.
5.2.8.8 Riparian "Legacy" Trees and Invasive Alien Species
At each littoral-riparian station (A-K), search for the largest tree visible. Confine your
search to within 100m (or as far as you can see) from the wetted bank on either side of the river
from each transect upstream and downstream. Classify this tree as broadleaf deciduous,
coniferous, or broadleaf evergreen (classify western larch as coniferous). Identify, if possible,
the species or the taxonomic group of this tree from the list provided in Table 5.2-12 (also on
field form) and estimate its height, diameter at breast height (dbh) and distance from the wetted
margin of the river. You may need to use binoculars to make these determinations. Enter this
information on the left hand column of the field form for Riparian "Legacy" Trees and Invasive
Alien Plants (Figure 5.2-8). If the largest tree is a dead "snag", enter "Snag" as the taxonomic
group. Note that the tree you choose may not truly be a "Legacy" tree; we use this data to
determine if there are Legacy Trees along the stream reach.
Search in the 10 m x 20 m riparian and littoral plots on both banks for the presence of
any invasive alien species listed in the NRSA Invasive Species Guide provided to each field
crew. Document the species observed on the Riparian "Legacy" Trees and Invasive Alien Plants
form (Figure 5.2-8), answering the question of whether each of the target species is present in
the plot. If you have a camera, document the species with a photograph. If you observe no alien
taxa within the riparian and littoral plots, but can confidently identify them outside of the plots,
include your observations in the comments portion of the form. If the river is too wide to
effectively observe the far bank at a transect, record what you observe for the plot on the near
bank, record a "U" flag, and explain in the comments section of the form.
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Table 5.2-13. Procedure for identifying riparian legacy trees and alien invasive species
Legacy Trees:
Beginning at Transect A, look upstream and downstream as far as you can see within the 100m
of the wetted bank but look no further downstream than half of the distance to the next transect.
Locate the legacy tree from within that area.
Classify this tree as broadleaf deciduous, coniferous, or broadleaf evergreen (classify western
larch as coniferous). Identify, if possible, the species or the taxonomic group of this tree from
the list below.
1. Acacia/Mesquite 10. Poplar/Cottonwood
2. Alder/Birch 11. Snag (Dead Tree of Any Species)
3. Ash 12. Spruce
4. Cedar/Cypress/Sequoia 13. Sycamore
5. Fir (including Douglas Fir, 14. Willow
Hemlock)
6. Juniper 15. Unknown, other Broadleaf Evergreen
7. Maple/Boxelder 16. Unknown or Other Conifer
8. Oak 17. Unknown or Other Deciduous
9. Pine 18. Elm
NOTE: If the largest tree is a dead "snag", enter "Snag" as the taxonomic group.
Estimate the height of the potential legacy tree, its diameter at breast height (dbh) and its
distance from the wetted margin of the stream. Enter this information on the left hand column
of the Riparian "Legacy" Trees and Invasive Alien Plants field form.
Alien Invasive Species:
Examine the 10m x 20m riparian and littoral plots on both banks for the presence of alien
species. (Species lists will be provided)
Record the presence of any species listed within the plots on either the left or right bank on the
Riparian "Legacy" Trees and Invasive Alien Species field form. If none of the species listed is
present in the plots at a given transect, fill in the circle indicating "None" for this transect.
Repeat for each remaining transect (B through K). At transect "K", look upstream a distance of 4
channel widths) when locating the legacy tree.
Any invasive species seen but not included on this list should be written in the comments
section.
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National Rivers and Streams Assessment
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Date: April 2009
Page 84
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5.2.9 Channel Constraint Assessment
After completing the thalweg profile and littoral-riparian measurements and observations,
visualize the stream at bankfull flow and evaluate the degree, extent and type of channel
constraint, following the procedure in Table 5.2-12. Figure 5.2-9 illustrates anastomosing and
braided channel types. Use the definitions on the Channel Constraint Assessment form (Figure
5.2-10) to classify the channel. Estimate the percent of the channel margin in contact with
constraining features (for unconstrained channels, this is 0%). To aid in this estimate, you may
wish to refer to the individual transect assessments of incision and constraint. Finally, estimate
the "typical" bankfull channel width and visually estimate the average width of the valley floor.
(valley floor width can often be determined from 1:24,000-scale topographic maps).
Table 5.2-14. Procedures for assessing channel constraint
NOTE: These activities are conducted after completing the thalweg profile and littoral-riparian
measurements and observations, and represent an evaluation of the entire stream reach.
Record this information on the Channel Constraint Form.
CHANNEL CONSTRAINT: Determine the degree, extent, and type of channel constraint based on
envisioning the stream at bankfull flow.
Classify the stream reach channel pattern as predominantly one channel, an anastomosing
channel, or a braided channel.
One channel may have occasional in-channel bars or islands with side channels, but
feature a predominant single channel, or a dominant main channel with a subordinate side
channel.
Anastomosing channels have relatively long major and minor channels branching and
rejoining in a complex network separated by vegetated islands, with no obvious dominant
channel.
Braided channels also have multiple branching and rejoining channels, separated by
unvegetated bars. Subchannels are generally small, short, and numerous, often with no
obvious dominant channel.
After classifying the channel pattern, determine whether the channel is constrained within a
narrow valley, constrained by local features within a broad valley, unconstrained and free to
move about within a broad floodplain, or free to move about, but within a relatively narrow valley
floor.
Then examine the channel to ascertain the bank and valley features that constrain the stream.
Entry choices for the type of constraining features are bedrock, hillslopes, terraces/alluvial fans,
and human land use (e.g., a road, a dike, landfill, rip-rap, etc.).
Estimate the percent of the channel margin in contact with constraining features (for uncon-
strained channels, this is 0%).
Finally, estimate the "typical" bankfull channel width. To aid in this estimate, you may wish to
refer to the individual transect assessments of incision and constraint that were recorded on the
Channel/Riparian Cross-Section Forms.
Visually estimate the average width of the valley floor. If the valley is wider than you can directly
estimate, record the distance you can see and mark the box on the field form.
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A) Anastomosing channel pattern
Vegetated islands above bankfull flow. Multiple
channels remain during major flood events.
B) Braided channel pattern
Unvegetated bars below bankfull flow. Multiple
channel pattern disappears during major flood events.
DVP
Figure 5.2-9. Types of multiple channel patterns.
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CHANNEL CONSTRAINT FORM - WADEABLE/BQATABLE
SITE
DATE
CHANNEL CONSTRAINT
CHANNEL PATTERN (Fill in one)
• One channel
U Anastomosing (complex) channel - (Relatively long major ano minor channels branching and rejoining.)
O Braided channel - (Multiple short channels branching and rejoining • mainly one channel broken up by
numerous mid-channe! bars.)
CHANNEL CONSTRAiNTfFill in one)
O Channel very constrained in V-shaped valley (i.e it is very unlikely to spread out over valley or erode a
new channel tiurtng flood)
9 Channel is in Broad Valley but channel movemenl by erosion during floods is constrained by Incision (Flood
flows do not commonly spread over valley floor or into multiple channels.)
U Channel is in Narrow Valley but is not very constrained, but limited in movement by relatively narrow
valley floor K -10 x bankfull width)
O Channel is Unconstrained in Broad Valley (i.e during flood it can fill off-channel areas and side channels,
spread out over flood plain, or easily cut new channels by erosion)
CONSTRAINING FEATURES (Fill In one)
-' Bedrock (i 6. channel is a b^drock-dominalad qorge)
'-.-' HHIslope (i.e. channel constrained in narrow V-shaped valley)
9 Terrace (i e. channel is constrained by its own incision into river/stream gravel/soil deposits')
O Human Bank Alterations (i e. constrained by rip-rap, landfill, rjike, road, etc.)
O No constraining features
Percent of channel length with margin
in contact with constraining feature:
Bankfutl widtn
(0-100%)
tf you cannot s« tht valley b'0«ters, record the
distance you cart see and irtaj'fc iMt too*.
Percent of Channel Margin
Comments
OliBS'2008 2Bfli Ctan Constf iinl
Figure 5.2-10. Channel Constraint Form.
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5.2.10 Debris Torrents and Recent Major Floods
Debris torrents, or lahars, differ from conventional floods in that they are flood waves of
higher magnitude and shorter duration, and their flow consists of a dense mixture of water and
debris. Their high flows of dense material exert tremendous scouring forces on streambeds. For
example, in the Pacific Northwest, flood waves from debris torrents can exceed 5 meters deep
in small streams normally 3 m wide and 15 cm deep. These torrents move boulders in excess of
1 m diameter and logs >1 m diameter and >10 m long. In temperate regions, debris torrents
occur primarily in steep drainages and are relatively infrequent, occurring typically less than
once in several centuries. They are usually set into motion by the sudden release of large
volumes of water upon the breaching of a natural or human-constructed impoundment, a
process often initiated by mass hillslope failures (landslides) during high intensity rainfall or
snowmelt. Debris torrents course downstream until the slope of the stream channel can no
longer keep their viscous sediment suspension in motion (typically <3% for small streams); at
this point, they "set up", depositing large amounts of sediment, boulders, logs, and whatever
else they were transporting. Upstream, the torrent track is severely scoured, often reduced in
channel complexity and devoid of near-bank riparian vegetation. As with floods, the massive
disruption of the stream channel and its biota are transient, and these intense, infrequent events
will often lead to a high-quality complex habitat within years or decades, as long as natural
delivery of large wood and sediment from riparian and upland areas remains intact.
In arid areas with high runoff potential, debris torrents can occur in conjunction with flash
flooding from extremely high-intensity rainfall. They may be nearly annual events in some steep
ephemeral channels where drainage area is sufficient to guarantee isolated thunderstorms
somewhere within their boundaries, but small enough that the effect of such storms is not
dampened out by the portion of the watershed not receiving rainfall during a given storm.
Because they may alter habitat and biota substantially, infrequent major floods and
torrents can confuse the interpretation of measurements of stream biota and habitat in regional
surveys and monitoring programs. Therefore, it is important to determine if a debris torrent or
major flood has occurred within the recent past. After completing the thalweg profile and
channel/riparian measurements and observations, examine the stream channel along the entire
sample reach, including its substrate, banks, and riparian corridor, checking the presence of
features described on the Torrent Evidence Assessment Form (Figure 5.2-11). It may be
advantageous to look at the channel upstream and downstream of the actual sample reach to
look for areas of torrent scour and massive deposition to answer some of the questions on the
field form. For example, you may more clearly recognize the sample reach as a torrent
deposition area if you find extensive channel scouring upstream. Conversely, you may more
clearly recognize the sample reach as a torrent scour reach if you see massive deposits of
sediment, logs, and other debris downstream.
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TORRENT EVIDENCE ASSESSMENT FORM
SITE ID FW08XX£>OO
DATE O 7 / O f I 2 0
(TORRENT EVIDENCE
Please fif! in any of the following that are evident
EVIDENCE OF TORRENT SCOURING:
I 01 - Stream channel has a recently d«vegetate«l corridor two or more times the width of (tie low flow channel. This
O corridor lacks riparian vegetation with possible exception of fireweed, even-aged alder or eottonwood seedlings,
grasses, or other herbaceous plants.
02 - Stream substrate cobbles or large gravel particles am NOT IMBRICATED, jlmbncatid means that they lie with flat
sidas horizontal and that they are stacked like roof shingles -imagine the upstream direction as the top erf the "roof.") In
| a torrent scour or deposition channel, the stones are laying in unorganized patterns, lying "every which way " In addition
J many ot tlM substrate partMstes are angular {not "water-worn.")
O 03 - Channel has little evidence of pool-riffle structure. (For eiample, could you ride a mountain bike down the channel?)
O
04 - Th© stream channel Is scoured down to bedrock for substantial portion of reach.
05 - There arc gravel or cobble terms (little levees) above bankfyll level.
O
Ofi - Downstream of the scoured reach (possibly several mites), there are massive deposits of sediment, logs, and other
debris
Q 07 - Riparian trees have fresh bar* scars at many points along the stream at seemingly unbelievable heights above the
| channel bed.
08 - Riparian trees have fatten into the channel as a result of scouring near their roots.
EVIDENCE OF TORRENT DEPOSITS:
O
09 - There are massive deposits of sediment, logs, and other debris in the reach- They may contain wood and boulders
that in your judgement, could not have been movod by the stream at even extreme flood stage.
10 - rf the stream has begun to erode newly laid deposits, it is evident that these deposits are "MATRIX SUPPORTED,"
I This means that the large particles, like boulders arid cobbles, are often not touching each other, but have silt sand, and
other fine particles between them (their weight is supported by these fine particles - in contrast to a normal stream
deposit, where fines, if present, normally Till-in" th® interstices between coarser particles.)
NO EVIDENCE;
11 - Mo evidence of torrent sccnirmg of torrent deposits.
COMMENTS
NRSA 10fr?«mt Evidence
Figure 5.2-11. Torrent Evidence Form.
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5.3 Periphyton
5.3.1 Summary of Method
Collect periphyton from the near-shore shallows at each of the sampling stations located
on the 11 cross-section transects ("A" through "K") established within the sampling reach.
Collect periphyton samples at the same time as sediment enzyme samples (Section 5.1.4) and
benthic macroinvertebrate samples (Section 5.4). Prepare one composite sample of periphyton
for each site. At the completion of the day's sampling activities, but before leaving the site,
prepare four types of laboratory samples (an ID/enumeration sample to determine taxonomic
composition and relative abundances, a chlorophyll sample, a biomass sample (for ash-free dry
mass [AFDM]), and an acid/alkaline phosphatase activity [APA] sample) from the composite
periphyton sample.
5.3.2 Equipment and Supplies
Table 5.3-1 is a checklist of equipment and supplies required to conduct periphyton
sample collection and processing activities. This checklist is similar to the checklist presented in
Appendix A, which is used at the base location (Section 3) to ensure that all of the required
equipment is brought to the river.
Table 5.3-1. Equipment and supplies list for periphyton at non-wadeable sites
For collecting
samples
Large Funnel (15-20 cm diameter)
12-cm2 area delimiter (3.8 cm diameter pipe, 3 cm tall)
Stiff-bristle toothbrush with handle bent at 90° angle
1-L wash bottle for stream water
500-mL plastic bottle for the composite sample with marked volume
gradations
60-mL plastic syringe with 3/8" hole bored into the end
Aspirator
Cooler with bags of ice
Field Operations Manual or laminated Quick Reference Guide
For recording
measurements
Sample Collection Form
Soft (#2) lead pencils for recording data on field forms
Fine-tipped indelible markers for sample labels
Sample labels (4 per set) with the sample ID number
Clear tape strips for covering labels
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5.3.3 Sampling Procedure
At each of the 11 transects, collect samples from the sampling station assigned during
the layout of the reach. Collect the substrate selected for sampling from a depth no deeper than
0.5 m. If you cannot collect a sample because the location is too deep, skip the transect. The
procedure for collecting samples and preparing a composite sample is presented in Table 5.3-2.
Collect one sample from each of the transects and composite in one bottle to produce one
composite sample for each site. Record the volume of the sample on the Sample Collection
Form as shown in Figure 5.1-4.
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Table 5.3-2. Procedure for collecting composite index samples of periphyton at non-wadeable
sites
1. Starting with Transect "A", collect a single sample from the assigned sampling station using the
procedure below.
a) Collect a sample of hard substrate (rock or wood) that is small enough (< 15 cm diameter) and
can be easily removed from the river. Place the substrate in a plastic funnel which drains into a
500-mL plastic bottle with volume graduations marked on it.
b) Use the area delimiter to define a 12-cm2 area on the upper surface of the substrate. Dislodge
attached periphyton from the substrate within the delimiter into the funnel by brushing with a stiff-
bristled toothbrush for 30 seconds. Take care to ensure that the upper surface of the substrate is
the surface that is being scrubbed, and that the entire surface within the delimiter is scrubbed.
c) Fill a wash bottle with river water. Wash the dislodged periphyton from the piece of substrate,
brush, delimiter and funnel into the 500-mL bottle. Use an appropriate amount of water to bring
the sample up to the next gradation. Doing so should result in collecting approximately 45ml_ of
sample at each transect.
d) If no coarse sediment (cobbles or larger) are present:
• Use the area delimiter to confine a 12-cm2 area of soft sediments.
• Either:
Vacuum the top 1 cm of sediment from within the delimited area into a de-tipped
60- ml syringe.
Use an aspirator to suction the top 1 cm of sediment from within the delimited area
into the sample bottle.
• Empty the syringe into the same 500-mL plastic bottle as above.
e) Put the bottle in a cooler on ice while you travel between transects and collect the
subsequent samples. (The samples need to be kept cool and dark because a chlorophyll
sample will be filtered from the composite.)
2. Repeat Step 1 for transects "B" through "K". Place the sample collected at each sampling site into the
single 500-mL bottle to produce the composite index sample.
3. After samples have been collected from all 11 transects, thoroughly mix the 500-mL bottle regardless
of substrate type.
4. Record the total volume of the composite sample in the periphyton section of the Sample Collection
Form.
5. If you are unable to collect a sample at any location, mark it on the field form and record the volume of
overall sample collected.
5.3.4 Sample Processing in the Field
You will prepare four different types of laboratory samples from the composite index
samples: an ID/enumeration sample (to determine taxonomic composition and relative
abundances), a chlorophyll sample, a biomass sample (for ash-free dry mass [AFDM]), and
an acid/alkaline phosphatase activity (APA) sample. All the sample containers required for an
individual site should be sealed in plastic bags until use to avoid external sources of
contamination (e.g., dust, dirt, or mud) that are present at site shorelines. Please refer to
Sections 7.2.5 and 7.2.6 processing the periphyton samples.
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5.4 Benthic Macroinvertebrates
5.4.1 Summary of Method
Collect benthic macroinvertebrate composite samples using a D-frame net with 500 urn
mesh openings. Take the samples from the sampling stations at the 11 transects equally
distributed along the targeted reach. Composite all sample material and field-preserve with
-95% ethanol.
5.4.2 Equipment and Supplies
Table 5.4-1 shows the checklist of equipment and supplies required to complete the
collection of benthic macroinvertebrates at non-wadeable sites. This checklist is similar to the
checklist presented in Appendix A, which is used at the base location to ensure that all of the
required equipment is brought to the site.
Table 5.4-1. Equipment and supplies list for benthic macroinvertebrate collection at non-
wadeable sites
For collecting
samples
Modified kick net (D-frame with 500
|jm mesh) and 4-5 ft handle
Spare net(s) and/or spare bucket
assembly for end of net
Buckets, plastic, 8- to 10-qt
Sieve bucket with 500 urn mesh
openings (U.S. std No. 35)
Watchmakers' forceps
Wash bottle, 1-L capacity labeled
"STREAM WATER"
Funnel, with large bore spout
Small spatula, spoon, or scoop to
transfer sample
Sample jars, 1-L HOPE plastic
suitable for use with ethanol
95% ethanol, in a proper container
Cooler (with absorbent material) for
transporting ethanol & samples
Plastic electrical tape
Scissors
Field Operations Manual or
laminated Quick Reference Guide
For recording
measurements
Composite benthic sample labels with
& without preprinted ID numbers
Blank labels on waterproof paper for
inside of jars
Soft (#2) lead pencils
Fine-tip indelible markers
Clear tape strips
Sample Collection Form
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5.4.3 Sampling Procedure
Collect benthic macroinvertebrate samples at the 11 transects and within the sampling
stations for non-wadeable streams. The process for selecting the sample stations is described
in the Initial Site Procedures Section (Section 4). Collect all benthic samples at non-wadeable
sites from the dominant habitat type within the 10 m x 15 m randomly selected sampling station
at each transect (Figure 5.4-1). Take 1 linear meter sweep at the dominant habitat type. Record
the benthic macroinvertebrate collection data on the Sample Collection Form, Side 1 as seen in
Figure 5.1-2.
The sampling process for collecting benthic samples from non-wadeable sites is
illustrated in Figure 5.4-2 and described in Table 5.4-2.
10m
A
Continue collecting samples
through Transect K
Figure 5.4-1.
sites.
Transect sample design for collecting benthic macroinvertebrates at non-wadeable
5.4.4 Sample Processing in Field
Use a 500 |o,m mesh sieve bucket placed inside a larger bucket full of site water while
sampling to carry the composite sample as you travel around the site. It is recommended that
teams carry a sample bottle containing a small amount of ethanol with them to enable them to
immediately preserve larger predaceous invertebrates such as helgramites and water beetles.
Doing so will help reduce the chance that other specimens will be consumed or damaged prior
to the end of the field day. Once the sample from all stations is composited, sieved and reduced
in volume, store in a 1-liter jar and preserve with 95% ethanol. Multiple jars may be required if
detritus is heavy (Table 5.4-3). It is suggested that no more than 5 1-L jars be used at any site.
If more than one jar is used for a composite sample, use the "extra jar" label provided; record
the SAME sample ID number on this "extra jar" label. DO NOT use two different sample
numbers on two jars containing one single sample. Remove any inorganic material (rocks,
debris, etc) before preserving sample. Cover the labels with clear tape. The sample ID number
is also recorded with a No. 2 lead pencil on a waterproof label that is placed inside each jar. Be
sure the inside label and outside label describe the same sample. If there is a large amount of
organic material in the sample, or there are adverse field conditions (i.e. hot, humid weather),
place sample in a 1-L jar with ethanol after each station.
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Record information for each composite sample on the Sample Collection Form as shown
in Figure 5.1-2. If a sample requires more than one jar, make sure the correct number of jars for
the sample is recorded on the Sample Collection Form. Do not fill out the collection form
until you have collected (or confirmed at the site that you will collect) samples. If forms
are filled out before you arrive at the site, and then no samples are collected, a lot of time is
wasted by others later trying to find samples that do not exist. If you are unable to collect a
sample at any station, make note of it on the sample collection form. Place the samples in a
cooler or other secure container for transporting and/or shipping to the laboratory (see Appendix
C).
NON-WADEABLE
At Transect "A", locate the first sampling station &
determine the dominant habitat type.
Sweep 1 linear meter of
dominant habitat type at the sampling station.
Transfer sample into sieve bucket.
Mark the habitat, substrate, and
channel type on the Sample Collection
Form.
Thoroughly rinse net into the sieve bucket.
Immediately preserve large predaceous
invertebrates in ethanol.
Proceed to sampling station on Transect "B" and
collect next sample.
Proceed to sampling station on Transect "C" and
collect next sample; continue collecting samples
throuah Transect "K".
The samples from all stations are composited to
create a single sample for the site.
Figure 5.4-2. Benthic macroinvertebrate collection at non-wadeable sites.
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Table 5.4-2. Procedure for benthic macroinvertebrate sampling at non-wadeable sites
1. After locating the sampling station site according to procedures described in the physical habitat
section, identify the dominant habitat type within the plot:
• Rocky/cobble/gravel/large woody debris • Organic fine mud or sand
• Macrophyte beds • Leaf Pack
2. Use the D-frame dip net (equipped with 500 urn mesh) to sweep through 1 linear meter of the most
dominant habitat type within the 10m x 15m sampling station, making sure to disturb the substrate
enough to dislodge organisms.
• If the dominant habitat is rocky/cobble/large woody debris it may be necessary to exit the boat
and disturb the substrate (e.g., overturn rocks, logs) using your feet while sweeping the net
through the disturbed area.
• Because a dip-net is being used for sampling, the maximum depth for sampling will be
approximately 0.5 m; therefore, in cases in which the depth of the river quickly drops off it may be
necessary to sample in the nearest several meters to the shore.
3. After completing the 1 linear meter sweep, remove all organisms and debris from net and place them
in a bucket following sample processing procedures described in the following section.
4. Record the sampled habitat type on the Sample Collection Form.
a) Fine/sand: not gritty (silt/clay/muck <0.06 mm diam.) to gritty (up to ladybug sized 2 mm diam.)
b) Gravel: fine to coarse gravel (ladybug to tennis ball sized; 2 mm to 64 mm diam.)
c) Coarse: Cobble to boulder (tennis ball to car sized; 64 mm to 4000 mm)
d) Other: bedrock (larger than car sized; > 4000 mm), hardpan (firm, consolidated fine substrate),
wood of any size, aquatic vegetation, etc.). Note "other" substrate in comments on field form.
5. Identify the channel habitat type where the sampling sweep was located. Mark the appropriate
channel habitat type for the transect on the Sample collection Form.
a) Pool; Still water; low velocity; smooth, glassy surface; usually deep compared to other parts of
the channel
b) GLide: Water moving slowly, with smooth, unbroken surface; low turbulence
c) Riffle: Water moving, with small ripples, waves, and eddies; waves not breaking, and surface
tension is not broken; "babbling" or "gurgling" sound.
d) RApid: Water movement is rapid and turbulent; surface with intermittent "white water" with
breaking waves; continuous rushing sound.
6. Proceed to the next sampling station and repeat steps 1-5. The organisms and detritus collected at
each station on the river should be combined in a single bucket to create a single composite sample
for the river. After sampling at all 11 stations is completed, process the composite sample in the
bucket according to procedures described in the following section.
7. If the sample contains primarily organic material, or if adverse weather conditions exist (i.e. hot humid
weather) process the sample at each station by placing it in a 1-L nalgene jar with ethanol. Follow
instructions in Table 5.4-3.
8. Immediately preserve larger predaceous invertebrates such as helgramites and water beetles in
ethanol.
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Table 5.4-3. Procedure for compositing samples for benthic macroinvertebrates at non-wadeable
sites
Estimate the total volume of the sample in the sieve and determine how large a jar will be
needed for the sample (500-mL or 1-L) and how many jars will be required. It is suggested that
no more than 5 1-L jars are used at each site.
Fill in a sample label with the Sample ID and date of collection. Attach the completed label to
the jar and cover it with a clear tape strip. Record the Sample ID for the composite sample on
the Sample Collection Form. For each composite sample, make sure the number on the form
matches the number on the label.
Wash the contents of the sieve to one side by gently agitating the sieve in the water. Wash the
sample into a jar using as little water from the wash bottle as possible. Use a large-bore funnel if
necessary. If the jar is too full pour off some water through the sieve until the jar is not more
than 1/3 full, or use a second jar if a larger one is not available. Carefully examine the sieve for
any remaining organisms and use watchmakers' forceps to place them into the sample jar.
Remove any inorganic material, such as gravel, by rinsing the material, examining it and
removing it from the sample.
• If a 2nd jar is needed, fill in a label that does not have a pre-printed ID # on it. Record the
ID # from the pre-printed label prepared above in the "SAMPLE ID" field of the label.
Attach the label to the 2nd jar and cover it with a strip of clear tape. Record the number of
jars on the Sample Collection Form. Make sure the number you record matches the
actual number of jars used. Write "Jar N of X' on each sample label using a waterproof
marker. Try to use no more than 5 jars per site.
Place a waterproof label inside each jar with the following information written with a #2 lead
pencil:
Site ID • Collectors initials
Type of sampler and mesh size used • Number of stations sampled
Name of site
Date of collection • Jar"N"of"X"
Completely fill the jar with 95% ethanol (no headspace). It is very important that sufficient
ethanol be used, or the organisms will not be properly preserved. Existing water in the jar
should not dilute the concentration of ethanol below 70%.
• NOTE: Composite samples can be transported back to the vehicle before adding ethanol
if necessary. In this case, fill the jar with stream water, then drain using the net (or sieve)
across the opening to prevent loss of organisms, and replace with ethanol at the vehicle.
Replace the cap on each jar. Slowly tip the jar to a horizontal position, then gently rotate the jar
to mix the preservative. Do not invert or shake the jar. After mixing, seal each jar with plastic
tape.
Store labeled composite samples in a container with absorbent material that is suitable for use
with 70% ethanol until transport or shipment to the laboratory.
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5.5 Fish
5.5.1 Summary of Method
The fish sampling method is designed to provide a representative sample of the fish
community, collecting all but the rarest fish inhabiting the site. It is assumed to accurately
represent species richness, species guilds, relative abundance, size, and anomalies. The goal
is to collect fish community data that will allow the calculation of an Index of Biotic Integrity (IBI)
and Observed/Expected (O/E) models. Boat electrofishing is the preferred method of sampling.
If electrofishing is not possible due to safety concerns, high turbidity, or extremes in
conductivity, complete the "Not Fished" section of the field form and comment why.
The time and effort necessary to sample the reach in its entirety is prohibitive in the
context of the survey, thus sub-sampling is required. Electrofishing will occur in a downstream
direction at all habitats along alternating banks (see section 5.5.3), over a length of 20 times the
mean channel width (Transects A through F). Collection of a minimum of 500 fish is required. If
this target is not attained, sampling will continue until 500 individuals are captured or the
downstream extent of the site (transect K) is reached. Identification and processing offish
should occur at the completion of each transect. If sampling cannot happen at any individual
transect, record it on the field collection form.
5.5.2 Equipment and Supplies
Table 5.5-1 shows the checklist of equipment and supplies required to complete the non-
wadeable fish assessment. This checklist is similar to the one presented in Appendix A, which is
used at the base location to ensure that all of the required equipment is brought to the site.
Record fish collection data on the Fish Collection Form, Side 1 (Fig. 5.5-1). Additional sheets
may be necessary - remember to indicate the transect on each form.
Table 5.5-1. Equipment and supplies — fish assessment at non-wadeable sites.
For collecting
samples
Boat, motor, and trailer (and
necessary safety equipment)
Gasoline and oil (if using a 2 cycle)
Boat electrofishing equipment
• Pulsator Control Box
• Foot Pedal
• Anode Droppers
• Generator
• Linesman's Gloves
• Hearing Protection
Tow barge electrofishing equipment
• Probes with extensions.
• Appropriate switching box
Dip nets (non-conductive handles)
%" mesh
Scientific collection permit
GPS with transect waypoints preloaded
Several Leak-proof HOPE jars for fish
voucher specimens (various sizes from
250 mL - 4L)
1 scalpel for slitting open large fish before
preservation
1 container of 10% buffered formalin
1 Minnow net for dipping small fish from
live well
2 measuring boards (3 cm size classes)
1 set Fish ID keys
Field Operations Manual and/or
laminated Quick Reference Guide
Digital camera with extra memory card &
battery
For recording
measurements
• Sample labels
• Sample Collection Form
• Soft (#2) lead pencils
• Fine-tip indelible markers
Clear tape strips
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O
o!
O
"S 1
si
>o
1§
o o
O
5
O
tt
o
n
0
0
0
I
MS
Nl
Figure 5.5-1. Fish Collection Form, Side 1.
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5.5.3 Sampling Procedure
Sampling will begin at the upstream half of the overall site, representing 20 times the
mean channel width. The total distance fished will depend upon the number of individuals
captured. Shoreline electrofishing will begin at transect A and proceed in a downstream
direction, alternating banks and terminating with the completion of subreach E-F (Figure 5.5-2).
Determination of the initial stream bank sampling location at transect A (i.e., right or left bank)
corresponds to the sequence established for physical habitat sampling and is determined at
random. Subreaches A-B, B-C, and C-D are sampled along the same bank before alternating to
the opposite bank to complete subreaches D-E and E-F. Each subreach is sampled for a
maximum of 700 seconds per subreach. Identification and processing of the sample should be
completed prior to beginning the next subreach. A minimum of 500 specimens is required. If
fewer than 500 individuals are captured, sampling must continue on alternating banks (again
following the pattern laid out for physical habitat sampling) until the minimum number is attained
or the downstream extent of the site (transect K) is reached (Figure 5.5-2).
Continue fishing on opposite bank for
2 subreaches; if 500 fish are
collected at this point, STOP
FLOW
B
Start fishing on the same
bank as the 1st randomly
selected sampling station
Reach length = 20 x mean wetted width
unless <500 fish collected
If <500 fish, continue fishing 1
subreach at a time on opposite
banks until 500 fish are collected
Figure 5.5-2. Transect sampling design for fish sampling at non-wadeable sites.
The sampling crew should consist of one boat operator (also controlling the
electrofishing unit) and one dip-netter (1/4" mesh dip nets) situated at the bow. Prior to sampling
each subreach, the crew should determine the most appropriate gear for the segment (e.g.,
boat or barge electrofishing units). Electrofishing should proceed downstream at a pace equal to
or slightly greater than the prevailing current to maximize capture efficiency. It may be
necessary to maneuver the electrofishing unit in and around complex habitat; however,
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discretion should be used in sampling these areas in order to maintain equal effort between
subreaches. Total effort expended (i.e., button time) over the five subreaches should be
approximately 3500 seconds. If additional subreaches are sampled, additional time will be
spent. To reduce stress and mortality, immobilized fish should be netted immediately and
deposited into a live-well for processing. For safety, all crew members are required to wear
personal floatation devices and insulated gloves. Polarized sunglasses and caps to aid vision
are also required. Table 5.5-2 provides the procedure for electrofishing in non-wadeable
streams.
Table 5.5-2. Procedure for electrofishing at non-wadeable sites.
1. Review all collecting permits to determine if any sampling restrictions are in effect for the site. In some
cases, you may have to cease sampling if you encounter certain State- or Federally-listed species.
2. Boat electrofishing will be used in non-wadeable streams, and the direction of fishing will be downstream.
If conductivity, turbidity, or safety precludes electrofishing, complete the "NOT FISHED" field on the Fish
Collection Form and comment why.
3. The sampling reach is defined as 20 times the mean channel width, corresponding to transects A through
F unless < 500 individuals are captured.
4. Shoreline electrofishing between each transect will occur on alternating banks following the sequence
established in the physical habitat procedures. Sampling will begin on the bank selected at random and
continue from transect A downstream for 700 seconds or until the next transect is reached. Subreaches
B-C and C-D are fished similarly; subreaches D-E and E-F will then be sampled on the opposite bank. If
fewer than 500 individuals are captured, sampling should continue until the minimum catch is attained or
the last subreach (J-K) is fished. Follow the systematic rotation of banks such that up to two subreaches
would be fished on the same bank prior to switching to the opposite bank. Crews must complete each of
the additional subreaches as described above, do not stop in the middle of any subreach, even if the 500
fish minimum is attained before the end of the subreach.
5. Set unit to pulsed DC and test settings outside of the sampling area. Start the electrofisher, set the timer,
and depress the switch to begin fishing. Typical settings are: 500-1OOOVDC; 8-20A; and 120 Hz. If fishing
success is poor, increase the pulse width first and then the voltage. Increase the pulse rate last to
minimize mortality or injury to large fish. If mortalities occur, first decrease pulse rate, then voltage, then
pulse width.
6. Once the settings on the electrofisher are adjusted to sample effectively and minimize injury and
mortality, begin sampling at the upstream reach (Transect A). Electrofishing proceeds downstream in
close proximity to the bank and at a pace equal to or slightly greater than the prevailing current to
maximize capture efficiency. Crews may "nose in" to habitat to effectively sample but should not remain in
that habitat for too long. Generally effort (i.e., button time) should be 700 seconds per subreach. At sites
with maximum reach length (4km) it is likely that the entire subreach (400m) will not be fished. Depending
upon the habitat complexity, variable distances may be fished in the time allotted. Distance sampled is
recorded on the Fish Collection Form.
7. Recommended mesh size on dip nets is 6mm (1/4"). Dip netters should actively capture stunned fish,
removing them from the electric field and immediately placing them in the livewell. Special attention
should be devoted to netting small and benthic fishes as well as fishes that may respond differently to the
current.
8. Process fish at the completion of each subreach to reduce mortality and track sampling effort. Release
fish in a location that eliminates the likelihood of recapture.
9. Complete header information on the Fish Collection Form. Record the number of seconds fished and the
estimated distance fished (as tracked by GPS or measured by range finder).
10. Repeat Steps 6 through 8 until subreach E-F and 500 individuals are captured or at a maximum,
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| subreach J-K is finished.
5.5.4 Processing Fish
Process fish when fish show signs of stress (e.g., loss of righting response, gaping,
gulping air, excessive mucus). Change water or stop fishing and initiate processing as soon as
possible. Similarly, State- and Federally-listed threatened or endangered species or large game
fish should be processed and released as they are captured. If periodic processing is required,
fish should be released in a location that prevents the likelihood of their recapture.
Use the Fish Collection Form - Large Wadeable/Boatable/Raftable. If several forms are
needed, use an extra form and note the page number on the top of the form as well as the
subreach sampled (i.e. Page 1 of 3). Taxonomic identification and processing should only be
completed on specimens greater than 25 mm total length and by crew members designated as
"fish taxonomic specialists" by EPA regional coordinators. Fish are tallied by species, evaluated
for maximum and minimum length, and examined for the presence of DELT (Deformities,
Eroded Fins, Lesions and Tumors) anomalies. Common names of species should follow those
established under the American Fisheries Society's publication, "Common and Scientific Names
of Fishes from the United States, Canada and Mexico" (Nelson, et al. 2004). A list of species
common to freshwater systems of the United States is presented in Appendix D.
Species not positively identified in the field should be separately retained (up to 20
individuals per species) for laboratory identification. Common names for retained species should
be assigned as "unknown", followed by its common family name and sequential lettering to
designate separate species (e.g., UNKNOWN SCULPIN A). Following positive laboratory
identification, field form information should be updated to reflect the actual species count and
number in the Final Count field. For example, if a sample of 20 specimens of species A is later
identified as 15 individuals of species A and 5 of species B, the Final Count of species A should
be corrected by assigning 25% to species B and 75% to species A. Table 5.5-3 presents the
procedure for processing fish.
Table 5.5-3. Procedure for processing fish at non-wadeable sites.
1. Complete all header information accurately and completely. If no fish were collected, complete the
"NONE COLLECTED" field on the Fish Collection Form.
2. Complete the information on the Fish Gear and Voucher/Tissue Sample Information Form.
3. Only identify and process individuals > 25mm in total length, ideally handling specimens only once.
Record the common name on the first blank line in the "COMMON NAME" Field of the Fish
Collection Form.
4. Fill in the Tag Number. The tag number is a number starting with 01 and continuing sequentially to
a number equal to the total number of species collected within the entire sample reach. Each
reoccurrence of a species within the entire reach should be assigned the same tag number as it
was assigned initially. For example, if a bluegill is assigned tag number 01 when processing fish
from the first subreach, all bluegills from the other subreaches will also be assigned tag number 01.
The purpose of the tag number is to connect species identifications with subsequent verification
and voucher collections.
5. If a species cannot be positively identified, assign it a sequential tag number in the Tag Number
Field and leave the "COMMON NAME" Field Blank. Flag this line and indicate in the "COMMENT"
field its common family name (e.g., UNKNOWN SCULPIN A). Retain a maximum subsample of 20
individuals for in-house laboratory identification of Unknowns. Do not include the number of each
species retained solely for in-house lab verification in the Voucher Count column of the fish
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collection form. This column is reserved only for those fish that are to be sent in for independent
re-identification as part of a complete voucher collection.
6. Process species listed as threatened and endangered first and return individuals immediately to the
stream. Photograph specimens for verification purposes if conditions permit and stress to
individuals will be minimal. Indicate if photographed on Fish Collection Form. If individuals are
killed, prepare them as verification specimens and preserve them in field.
7. Tally the number of individuals of each species collected in the "TALLY" box on the Fish Collection
Form and record the total number in the "TOTAL COUNT" field on the form. Do not enter a total for
fishes that must be identified in the laboratory.
8. Measure the total length of the largest and smallest individual to provide a size range for the
species. Record these values in the "LENGTH" area of the Fish Collection Form. If only one fish is
collected, leave the maximum field blank.
9. Examine each individual for external anomalies and tally those observed. Readily identify external
anomalies including missing organs (eye, fin), skeletal deformities, shortened operculum, eroded
fins, irregular fin rays or scales, tumors, lesions, ulcerous sores, blisters, cysts, blackening, white
spots, bleeding or reddening, excessive mucus, and fungus. After all of the individuals of a species
have been processed, record the total number of individuals affected in the "ANOMALIES" Field of
the Fish Collection Form.
10. Record the total number of mortalities due to electrofishing or handling on the Fish Collection Form.
11. Follow the appropriate procedure to prepare voucher specimens and/or to select specimens for
tissue samples. Release all remaining individuals so as to avoid their recapture.
12. For any line with a fish name, ensure that all spaces on that line are filled in with a number, even if
it is zero.
5.5.5 Taxonomic Quality Assurance/Quality Control
5.5.5.1 Sample Preservation
Fish retained for laboratory identification or voucher purposes should be placed in a
large sample jar containing a 10% buffered formalin solution in a volume equal to or greater
than the total volume of specimens. Individuals larger than 200 mm in total length should be slit
along the right side of the fish in the lower abdominal cavity to allow penetration of the solution.
Fish retained for laboratory identification or as vouchers should be preserved in the field
following the precautions outlined in the MSDS. All personnel handling 10% buffered formalin
must read the MSDS (Appendix D). Formalin is a potential carcinogen and should be used
with extreme caution, as vapors and solution are highly caustic and may cause severe
irritation on contact with skin, eyes, or mucus membranes. Wear vinyl or nitrile gloves
and safety glasses, and always work in a well-ventilated area.
5.5.5.2 Laboratory Identification of Fish
Fish that are difficult to identify in the field should be kept for laboratory identification or
to verify difficult field identifications. Table 5.5-4 outlines the laboratory identification process
and completing the Fish Collection Form. Field crews may use a supplemental Fish
Identification Lab sheet such as that shown in Figure 6.5-4 for internal laboratory use only.
Crews should retain the Fish verification sample - contact your regional EPA coordinator if you
cannot store the samples at your facility.
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Do not include the number of each species retained solely for in-house lab verification in
the Voucher Count column of the fish collection form. This column is reserved only for those
fish that are to be sent in for independent re-identification as part of a complete voucher
collection.
Field crews should not retain the Fish Collection Form(s) if the laboratory identification
process cannot be completed within a short period of time. If the time needed to complete the
identification/verification is expected to exceed two weeks, make copies of the Fish Collection
Form(s) and send the entire pack of original data forms to the Information Management
Coordinator. When the identification/verification process is complete, make the necessary
changes to the copied Fish Collection Form(s) and send them as soon as possible to the
Information Management Coordinator as well.
Table 5.5-4. Procedure for laboratory identification offish samples.
1. Fish may be retained for routine laboratory identification and verification purposes. Fish tags are provided
with each site kit. Crews may use these tags at their discretion in order to identify fish at their laboratory.
2. Retained fish should be placed in a large sample jar containing a 10% buffered formalin solution in a
volume equal to or greater than the total volume of specimens. Individuals larger than 200mm in total
length should be slit along the right side of the fish in the lower abdominal cavity to allow penetration of the
solution.
3. Following fixation for 5 to 7 days, the volume of formalin should be properly discarded and replaced with
tap water for soaking specimens over a 4-5 day period. Soaking may require periodic water changes and
should continue until the odor of formalin is barely detectable. Final storage of specimens is done in 45%-
50% isopropyl alcohol or 70% ethanol. Formalin is a potential carcinogen and should be used with
extreme caution, as vapors and solution are highly caustic and may cause severe irritation on contact with
skin, eyes, or mucus membranes. Wear vinyl or nitrile gloves and safety glasses, and always work in a
well-ventilated area.
4. Formalin must be disposed of properly. Contact your regional EPA coordinator if your laboratory does not
have the capability of handling waste formalin.
5. Unknown fish are identified to species in the laboratory. You may use a Fish Identification Lab Sheet such
as the one presented in Figure 6.5-4.
6. Fill in the Unknown species name in the "COMMON NAME" field of the Fish Collection Form and make
certain the "FINAL COUNT" field is correct.
7. If species field identifications were incorrect, correct the "COMMON NAME" Field by completely erasing
the Common Name and replacing the correct name. Add an additional Common Name if needed. Make
certain the "FINAL COUNT" field is correct. If the "COMMON NAME" Field was incorrect or cannot be
cleanly erased, cross out the line of data and fill out a new line with the correct "COMMON NAME" and
"FINAL COUNT".
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5.5.5.3 Voucher Specimens
Approximately 10% of each field crews' sites will be randomly pre-selected for re-
identification by an independent taxonomist. A minimum of one complete voucher is required for
each person performing field taxonomy and will consist of either preserved specimen(s) or
digital images representative of all species in the sample, including common species. Multiple
specimens per species can be used as vouchers, if necessary (i.e., to document different life or
growth stages, or sexes). Note that a complete sample voucher does not mean that all
individuals of each species will be vouchered, only enough so that independent verification can
be achieved.
Digital images should be taken as voucher documentation for species that are
recognized as Rare, Threatened, or Endangered - they should not be killed. Digital images
should also be taken of fish specimens too large for preservation.
Certain states or regions may require that more fish vouchers are taken. Check with
your state/regional coordinators to determine if your team will be required to collect complete
vouchers at more than 10% or your sites.
For the sample voucher, specimen containers should be labeled with the sample
number, site ID number, site name, and collection date. There should be no taxonomic
identification labels in or on the container, or in any of the digital photos.
Choose individual specimens that are intact and in good condition, such that re-
identification will be possible. Fish that are damaged, have significant scale loss or those that
have been dead for a significant amount of time prior to preservation should be avoided if
possible. Fish in pristine condition and those possessing clear identification characteristics are
preferred. Additionally, fish that are preserved while still live will typically flare their fins and gills
and will allow for easier re-identification in the laboratory.
Place one or more representative specimens of each species in plastic mesh sleeves
along with one of the corresponding tag number labels provided in your site kit. (Several fish
may be placed in a single mesh sleeve, as long as they are of the same species). Ensure that
the tag numbers in the voucher collection match the tag numbers on the fish collection data
forms. Seal both ends of the mesh sleeve with zip ties and place it inside the voucher collection
jar with the appropriate preservative. Unknown fish may be identified in the laboratory as
described in section 5.5.5.2 and subsequently included in the voucher collection.
Record the total number of each fish species retained for voucher purposes in each
subreach on the fish collection form. Record the voucher sample ID number on the fish gear /
voucher / fish tissue collection form. If no voucher is prepared for the site, fill in the "no
vouchers preserved" circle on the fish gear form.
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Table 5.5-5. Procedure for vouchering offish samples.
1. Approximately 10% of each field crews' sites will be randomly pre-selected for re-identification by an
independent taxonomist. A minimum of one complete voucher is required for each person performing field
taxonomy and will consist of either preserved specimen(s) and/or digital images representative of all
species in the sample, even common species.
2. Take digital images as voucher documentation for species that are recognized as Rare, Threatened, or
Endangered; or when fish specimens are too large for preservation.
3. For the sample voucher, label the specimen containers with the sample number, site ID number, site
name, and collection date. Do not put taxonomic identification labels in or on the container.
4. Place one or more representative specimens of each species in plastic mesh sleeves along with one of the
corresponding tag number labels provided in your site kit. (Several fish may be placed in a single mesh
sleeve, as long as they are of the same species).
5. Ensure that the tag numbers in the voucher collection match the tag numbers on the fish collection data
forms.
6. Seal both ends of the mesh sleeve with zip ties and place it inside the voucher collection jar with the
appropriate preservative.
7. Unknown fish may be identified in the laboratory as described in section 5.5.5.2 and subsequently
included in the voucher collection.
8. Record the total number of each fish species retained for voucher purposes in each subreach on the fish
collection form.
9. Record the voucher sample ID number on the fish gear / voucher / fish tissue collection form.
10. If no voucher is prepared for the site, fill in the "no vouchers preserved" circle on the fish gear form.
5.5.5.4 Photovouchering
Digital imagery should be used for fish species that cannot be retained as preserved
specimens (e.g., RTE species; or very large bodied fish). Views appropriate and necessary for
an independent taxonomist to accurately identify the specimen should be the primary goal of the
photography. Additional detail for these guidelines is provided in Staufferet al. (2001), and is
provided to all field crews as a handout.
The recommended specifications for digital images to be used for photovouchering
include: 16-bit color at a minimum resolution of 1024x768 pixels; macro lens capability allowing
for images to be recorded at a distance of less than 4 cm; and built-in or external flash for use in
low-light conditions. Specimens should occupy as much of the field of view as possible, and the
use of a fish board is recommended to provide a reference to scale (i.e., ruler or some
calibrated device) and an adequate background color for photographs. Information on Station
ID, Date and TAG NUMBER should also be captured in the photograph, so that photos can be
identified if file names become corrupted. All photovouchered species should have at least a
full-body photo (preferably of the left side of the fish) and other zoom images as necessary for
individual species, such as lateral line, ocular/oral orientation, fin rays, gill arches, or others. It
may also be necessary to photograph males, females, or juveniles.
Images should be saved in medium- to high-quality jpeg format, with the resulting file
name of each picture noted one the Fish Collection Form. It is important that time and date
stamps are accurate, as this information can also be useful in tracking the origin of photographs.
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Because close-up photography is difficult in the best of conditions with typical point and shoot
cameras, it might be best to take high quality pictures at a greater distance so that the image
can be zoomed with a PC. It is recommended that images stored in the camera be transferred
to a PC or storage device at the first available opportunity. At this time the original file should be
renamed to follow the logic presented below:
F01_CT003_20080326_A.jpg
Where:
F = fish
01 = TAG NUMBER
CT003 = state (Connecticut) and site number
20080326 = date (yyyymmdd)
A = first of several pictures of same fish (e.g., A, B, C)
Field crews should maintain files for the duration of the sampling season. Notification
regarding the transfer of all images to the existing database will be provided at the conclusion of
the sampling. Only keep photos that are useful for identifications. If photos are to be submitted
as vouchers, burn a CD of those photos that can be submitted along with the voucher jar.
5.6 Fish Tissue
5.6.1 Summary of Method
You will collect one predator species composite from each target site for human health
related analyses. The focus is on fish species that commonly occur throughout the region of
interest, and that are sufficiently abundant within a sampling reach. Each composite sample will
consist of five adult fish of the same species that are similar in size (the smallest individual in
the composite is no less than 75% of the total length of the largest individual). Collection occurs
in the sampling reach.
5.6.2 Equipment and Supplies
Table 5.6-1 lists the equipment and supplies necessary for field crews to collect fish
tissue samples. This list is comparable to the checklist presented in Appendix A, which provides
information to ensure that field teams bring all of the required equipment to the site. Record the
fish tissue sampling data on the Fish Gear and Voucher/Tissue Sample Information Form
(Figure 5.6-1).
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Table 5.6-1. Equipment and supplies—fish tissue collection at non-wadeable sites
For collecting fish
composite sample
Electrofishing equipment (including
variable voltage pulsator unit, wiring
cables, generator, electrodes, dip nets,
protective gloves, boots, and necessary
safety equipment)
Scientific collection permit
Sampling vessel (including boat, motor,
trailer, oars, gas, and all required safety
equipment)
Coast Guard-approved personal
floatation devices
Maps of target sites & access routes
Global Positioning System (GPS)
unit
Livewell and/or buckets
Measuring board (millimeter scale)
Clean nitrile gloves
For storing and
preserving fish
composite sample
Aluminum foil (solvent-rinsed and
baked)
Heavy-duty food grade polyethylene
tubing
Large plastic (composite) bags
Knife or scissors
Dry Ice
Plastic cable ties
Coolers
For documenting the
fish composite
sample
Fish Collection Forms
Clipboard
Sample Identification Labels
#2 pencils
Fine tipped indelible markers
For shipping the fish
composite samples
Preaddressed FedEx airbill
Coolers
Tracking Form
Chain-of-custody labels
Packing/strapping tape
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5.6.3 Sampling Procedure
The fish tissue indicator will be collected using the same gear and procedures used to
collect the fish community assemblage. Collection of individuals for fish tissue occurs in the
sample reach during the fish community assemblage sampling. If the five fish are not collected
during the community sampling, sample for up to one additional hour. If the sample is still not
collected, call the Logistics Coordinator at the end of the day and record on the field collection
form. If the target species are unavailable, the fisheries biologist will select an alternative
species (i.e., a species that is commonly consumed in the study area, with specimens of
harvestable or consumable size, and in sufficient numbers to yield a composite) to obtain a fish
composite sample from the species that are available. Recommended target species, listed in
order of preference, are given in Table 5.6-2.
Table 5.6-2. Recommended target species for fish tissue collection (in order of preference) at
non-wadeable sites
Predator/Gamefish Species
(in order of preference)
Family name
Centrarchidae
Ictaluridae
Percidae
Percichthyidae
Esocidae
Salmonidae
Common name
Large mouth bass
Smallmouth bass
Black crappie
White crappie
Channel Catfish
Blue Catfish
Flathead Catfish
Walleye/sauger
Yellow perch
White bass
Northern pike
Brown trout
Rainbow trout
Brook trout
Scientific name
Micropterus salmoides
Micro pter us dolomieu
Pomoxis nigromaculatus
Pomoxis annularis
Ictalurus punctatus
Ictalurus furcatus
Pylodictis olivaris
Sander vitreus /S.
canadensis
Perca flavescens
Morone chrysops
Esox lucius
Salmo trutta
Oncorhynchus mykiss
Salvelinus fontinalis
Length Guideline
(Estimated Minimum)
-280 mm
-300 mm
-330 mm
-330 mm
-300 mm
-300 mm
-350 mm
-380 mm
-330 mm
-330 mm
-430 mm
-300 mm
-300 mm
-330 mm
The procedures for collecting and processing fish composite samples are presented in
Table 5.6-3.
Table 5.6-3. Sampling procedure for fish composite samples at non-wadeable sites
1. Put on clean nitrile gloves before handling the fish. Do not handle any food, drink, sunscreen, or
insect repellant until after the composite sample has been collected, measured, and wrapped.
2. Rinse potential target species/individuals in ambient water to remove any foreign material from the
external surface and place in clean holding containers (e.g., livewells, buckets). Return non-target
fishes or small specimens to the river or stream.
3. Retain one predator species composite from each site. The composite must consist of 5 fish of
adequate size to provide a total of 500 grams of edible tissue for analysis (refer to Table 5.6-2 for
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minimum species length guidelines). Select fish for each composite based on the following
criteria:
• all are of the same species,
• all satisfy legal requirements of harvestable size (or weight) for the sampled river, or at least be
of consumable size if no legal harvest requirements are in effect,
• all are of similar size, so that the smallest individual in a composite is no less than 75% of the
total length of the largest individual, and
• all are collected at the same time, i.e., collected as close to the same time as possible, but no
more than one week apart (Note: Individual fish may have to be frozen until all fish to be
included in the composite are available for delivery to the designated laboratory).
Accurate taxonomic identification is essential in assuring and defining the organisms that have been
composited and submitted for analysis. Under no circumstances should individuals from different
species be used in a single composite sample.
4. Measure each individual fish to determine total body length. Measure total length of each
specimen in millimeters, from the anterior-most part of the fish to the tip of the longest caudal fin
ray (when the lobes of the caudal fin are depressed dorsoventrally).
5. Record sample number, species retained, specimen length, site ID, and sampling date on the Fish
Collection Form (Figure 5.5-1) in black ink. Mark site type ("Urban" or "Non-urban") next to the site
identification number at the top left of the fish form, and write primary or duplicate in the comment
section. Make sure the sample identification numbers recorded on the collection form match those
on the sample labels.
6. Remove each fish retained for analysis from the clean holding container(s) (e.g., livewell) using
clean nitrile gloves. Dispatch each fish using a clean wooden bat (or equivalent wooden device).
7. Wrap each fish in extra heavy-duty aluminum foil with the dull side in (foil provided by EPA as
solvent-rinsed, oven-baked sheets).
8. Prepare a Sample Identification Label for each sample, ensuring that the label information matches
the information recorded on the Fish Collection Form. Be sure to include fish species and
specimen length on each label.
9. Cut a length of food grade tubing (provided by EPA) that is long enough to contain each individual
fish and to allow extra length on each end to secure with cable ties. Place each foil-wrapped
specimen into the appropriate length of tubing. Seal each end of the tubing with a plastic cable tie.
Attach the fish sample label to the outside of the food-grade tubing with clear tape and secure the
label by taping around the entire fish (so that tape sticks to tape).
10. Place all the wrapped fish in the composite from each site in a large plastic bag and seal with
another cable tie.
11. After each sample is packaged, place it immediately on dry ice for shipment. If samples will be
carried back to a laboratory or other facility to be frozen before shipment, wet ice can be used to
transport wrapped and bagged fish samples in the coolers to a laboratory or other interim facility.
12. If possible, keep all (five) specimens designated for a particular composite in the same shipping
container (ice chest) for transport.
13. Samples may be stored temporarily on dry ice (replenishing the dry ice daily). You have the
option, depending on site logistics, of:
• shipping the samples packed on dry ice in sufficient quantities to keep samples frozen for up to
48 hours (50 pounds are recommended), via priority overnight delivery service (e.g., Federal
Express), so that they arrive at the sample preparation laboratory within less than 24 hours
from the time of sample collection, or
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• freezing the samples within 24 hours of collection at <-20°C, and storing the frozen samples
until shipment within 2 weeks of sample collection (frozen samples will subsequently be
packed on dry ice and shipped to the sample preparation laboratory via priority overnight
delivery service).
14. Ship fish tissue samples from urban sites to the EPA NERL lab in Cincinnati, OH and from non-
urban sites to the GLEG lab in Traverse City, Ml on Monday through Thursday.
5.7 Fecal Indicator (Enterococci)
5.7.1 Summary of Method
Collect a fecal indicator sample at the last transect (Transect K) after all other sampling
is completed. Samples must be filtered and the filters must be frozen within 6 hours of
collection. Use a pre-sterilized, 250 ml bottle and collect the sample approximately 1 m off the
bank at about 0.3 meter (12 inches) below the water surface. Following collection, place the
sample in a cooler, chill for at least 15 minutes, and maintain on ice prior to filtration of four 50
ml_ volumes. (Samples must be filtered and frozen on dry ice within 6 hours of collection). In
addition to collecting the sample, look for signs of disturbance throughout the reach that would
contribute to the presence of fecal contamination to the waterbody. Record these disturbances
on the Site Assessment Form (Figure 7-2).
5.7.2 Equipment and Supplies
Table 5.7-1 provides the equipment and supplies needed to collect the fecal indicator
sample. Record the sample data on the Sample Collection Form, Side 2 (Figure 5.1-4).
Table 5.7-1. Equipment and supplies list for fecal indicator sampling at non-wadeable sites
For collecting samples
nitrile gloves
pre-sterilized, 250 ml sample
bottle
sodium thiosulfate tablet
Wet ice
cooler
For recording
measurements
Sample Collection Form
Fecal Indicator sample labels
(4 vial labels and 1 bag label)
Pencils (for data forms)
Fine tipped indelible markers
(for labels)
Clear tape strips
5.7.3 Sampling Procedure
The procedure for collecting the fecal indicator sample is presented in table 5.7-2.
Table 5.7-2. Procedure for fecal indicator (Enterococci) sample collection at non-wadeable sites
1. Put on nitrile gloves.
2. Select a sampling location at transect K that is approximately 1 m from the bank and approximately
0.3m deep. Approach the sampling location slowly from downstream or downwind.
3. Lower the un-capped, inverted 250 ml sample bottle to a depth of 1 foot below the water surface,
avoiding surface scum, vegetation, and substrates. Point the mouth of the container away from the
body or boat. Right the bottle and raise it through the water column, allowing bottle to fill completely.
If the depth does not reach 0.3m along the transect at 1 m from the bank, take the sample and flag it
on the field form.
4. After removing the container from the water, discard a small portion of the sample to allow for proper
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mixing before analyses.
5. Add the sodium thiosulfate tablet, cap, and shake bottle 25 times.
6. Store the sample in a cooler on ice to chill (not freeze). Chill for at least 15 minutes and do not hold
samples longer than 6 hours before filtration and freezing.
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6.0 WADEABLE STREAMS
6.1 Water Quality
This section describes the procedures and methods for the field collection and analysis
of the water quality indicators (in-situ measurements, water chemistry, and sediment enzymes)
from wadeable streams and rivers.
6.1.1 In Situ Measurements of Dissolved Oxygen, pH, Temperature, and Conductivity
6.1.1.1 Summary of Method
You will measure dissolved oxygen (DO), pH, temperature, and conductivity by using a
multi-parameter water quality meter (or sonde). Take all measurements at the X site at 0.5 m
depth, or mid-depth if depth is <1 m. The site depth must be accurately measured before taking
the measurements, and care should be taken to avoid the probe contacting bottom sediments.
6.1.1.2 Equipment and Supplies
Table 6.1-1 provides the equipment and supplies needed to measure dissolved oxygen,
pH, temperature, and conductivity. Record the measurements on the Field Measurement Form,
as seen in Figure 6.1-1.
Table 6.1-1. Equipment and supplies—DO, pH, temperature, and conductivity
For taking measurements and
calibrating the water quality meter
For recording measurements
• Multi-parameter water quality meter with DO, pH,
temperature, and conductivity probes.
• Extra batteries
• De-ionized and tap water
• Calibration cups and standards
• QC calibration standard
• Barometer or elevation chart to use for calibration
• Field Measurement Form
• Pencils (for data forms)
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FIELD MEASUREMENT FORM - WADEABLE
SITE ID FW08XX0OC?
DATE:
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CALIBRATION INFORMATION
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6.1.1.3 Multi-Probe Sonde
Dissolved Oxygen Meter
Calibrate the DO meter prior to each sampling event. We recommend that the probe be
calibrated in the field against an atmospheric standard (ambient air saturated with water) prior to
sampling. In addition, manufacturers typically recommend periodic comparisons with a DO
chemical analysis procedure (e.g., Winkler titration) to check accuracy and linearity.
pH Meter
Calibrate the pH meter prior to each sampling event. Calibrate the meter in accordance
with the manufacturer's instructions and with the team agency's existing SOP. You must also
conduct a quality control check with the provided standard to verify the calibration and
periodically evaluate instrument precision (see Section 3.1.2). Crews must check their probe
once a week against the provided Quality Control Standard (QCS) and record the information
on the data forms.
Temperature Meter
You must check the accuracy of the sensor against a thermometer that is traceable to
the National Institute of Standards (NIST) at least once per sampling season. The entire
temperature range encountered in the NRSA should be incorporated in the testing procedure
and a record of test results kept on file.
Conductivity Meter
Calibrate the conductivity meter prior to each sampling event. Calibrate the meter in
accordance with the manufacturer's instructions. The entire conductivity range encountered in
the NRSA should be incorporated in the testing procedure and a record of test results kept on
file. You must also conduct a quality control check with the provided standard to verify the
calibration and periodically evaluate instrument precision (see Section 3.1.2). Crews must check
their probe once a week against the provided QCS and record the information on the data
forms.
6.1.1.4 Sampling Procedure
Table 6.1-2 presents step-by-step procedures for measuring dissolved oxygen, pH,
temperature, and conductivity.
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Table 6.1-2. Sampling procedure—temperature, pH, conductivity and dissolved oxygen
1. Check meter and probes and calibrate according to manufacturer's specifications.
2. Wadeable Sites: Measurements are taken at the X site at a depth of 0.5 meters or at mid-depth if
less than 1 meter deep.
3. Lower the sonde in the water and measure DO, pH, temperature, and conductivity at 0.5 m depth.
4. Record the measurements on the Field Measurement Form.
5. If sampling at the X-site is not possible, move to another part of the reach to collect the sample (as
close to the X-site as possible), record the letter of the nearest transect in the "TRANSECT" box and
more detailed reasons and/or information in the Comments section.
6. Flag any measurements that need further comment (or when a measurement cannot be made).
6.1.2 Water Chemistry Sample Collection and Preservation
6.1.2.1 Summary of Method
The water chemistry samples will be analyzed for total phosphorus (TP), total nitrogen
(TN), total ammonia-nitrogen (NH4), nitrate (NO3), basic anions, cations, total suspended solids
(TSS), turbidity, acid neutralizing capacity (ANC, alkalinity), dissolved organic carbon (DOC),
and total organic carbon (TOC). You will collect a grab sample in one 4-L cube container and in
one 2-L amber Nalgene bottle from the X site at the center of the reach. Store all samples on ice
in a closed cooler.
6.1.2.2 Equipment and Supplies
Table 6.1-3 provides the equipment and supplies needed to collect water samples at the
index site. Record the Water Sample Collection and Preservation data on the Sample Collection
Form, as seen in Figure 6.1-2.
Table 6.1-3. Equipment and supplies—water chemistry sample collection and preservation
For collecting samples
Nitrile gloves
4-L cube container for wadeable sites
2-L amber Nalgene bottle
3 L Nalgene beaker
Cooler with ice
Dl water (for cleaning beaker and carboy between sites)
Field Operations Manual and/or laminated Quick Reference Guide
For recording
measurements
Sample Collection Form
Field Measurement Form
Pencils (for data forms)
Fine tipped indelible markers
6.1.2.3 Sampling Procedure
Table 6.1-4 presents step-by-step procedures for collecting water chemistry samples at
wadeable sites.
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Table 6.1-4. Sampling procedure for wadeable sites—water chemistry sample collection
1. Collect the water samples from the X-site in a flowing portion near the middle of the stream.
2. Put on nitrile gloves. Make sure not to handle sunscreen or other chemical contaminants until after
the sample is collected.
3. Rinse the 3-L Nalgene beaker three times with water, and discard the rinse downstream.
4. Remove the cube container lid and expand the cube container by pulling out the sides. NOTE: DO
NOT BLOW into the cube container to expand them, this will cause contamination.
5. Fill the 3-liter beaker with water and slowly pour 30 - 50 ml into the cube container. Cap the cube
container and rotate so that the water contacts all the surfaces. Discard the water downstream.
Repeat this rinsing procedure 2 more times.
6. Fill the beaker with water and pour into the cube container. Repeat as necessary to fill the cube
container. Let the weight of the water expand the cube container. Pour the water slowly as the cube
container expands. Fill the cube container to at least three-fourths of its maximum volume. Rinse the
cube container lid with water. Eliminate any air space from the cube container, and cap it tightly.
Make sure the cap is tightly sealed and not on at an angle.
7. Fill the 3-liter beaker with water and slowly pour 30 - 50 mL into the 2 L amber Nalgene bottle. Cap
the bottle and rotate so that the water contacts all the surfaces. Discard the water downstream.
Repeat this rinsing procedure 2 more times.
8. Fill the beaker with water and pour into the 2 L amber Nalgene bottle. Cap the bottle tightly
9. Place the cube container and bottle in a cooler (on ice or water) and shut the lid. If a cooler is not
available, place the cube container in an opaque garbage bag and immerse it in the stream.
10. Record the Sample ID on the Sample Collection Form along with the pertinent stream information
(stream name, ID, date, etc.). Note anything that could influence sample chemistry (heavy rain,
potential contaminants) in the Comments section. If sampling at the X-site is not possible, move to
another part of the reach to collect the sample (as close to the X-site as possible), record the letter
of the nearest transect and more detailed reasons and/or information in the Comments section.
6.1.3 Sediment Enzymes
6.1.3.1 Summary of Method
Collect sediment samples at the 11 sampling stations along each reach and combine for
all stations at a site, resulting in a single 500 mL sample per site. Collect fine surface sediments
(top 5 cm) using a scoop, spoon or dredge. Store samples on ice until shipment to the
laboratory. Samples will be analyzed for available DIN, NH4, DIP, TP, TN, total carbon (TC) and
enzyme activity.
6.1.3.2 Equipment and Supplies
Table 6.1-5 lists the equipment and supplies needed to collect sediment enzyme
samples. Record collection data on the Sample Collection Form, as seen in Figure 6.1-2.
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Table 6.1-5. Equipment and supplies—sediment enzymes
For collecting samples
4 L graduated plastic bucket
Large stainless steel spoon for mixing sediment composite
500 ml plastic jar for storing sediment sample
For recording measurements
Sample Collection Form
Sample labels
Pencils
Fine tipped indelible markers
Clear tape strips
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46387
SAMPLE COLLECTION FORM - WADEABLE (Front) -
2 0
Edge:
U - Undercut
L = Leaf Litter
S = Snag R
OG = Ofganic
= Rootwad
deposits OT
M * Macrophyte bed
= Otber or Co- dominant
i n ooti
Substrate:
F - Fine/Sand C * Coarse substrate
G = Sravsl OT = Oftar (Explain in
•it s&etron belcw}
Channel:
P = Pool Rl = Rime GL ' Glide
RA = Rapid OT = Olhit {Explain in
3«cdcin
Flag code s: K - No meBaurarnerrt or observaticr mad E; U - St^pect meaau n^meji" or nbqeivfllion; F1, F2l«c. -misc. flags e
flags in comm«nl £«cUti!1&. ^3lmpl« Categories P - Primary, D = Field Duplicate
04/07.12009 NRSA Sam pla CollsctiDn WadsablB 20D9
crew. E xplei n all
Figure 6.1-2. Sample Collection Form, Side 1.
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Draft
SAMPLE COLLECTION FORM - WADEABLE (Back)
Reviev/ed lav
f initial):
SITE
ID: FW08
DATE:
COMPOSITE PERIPHYTON SAMPLE -
Sample ID
i i • ' * • •
Assemblage ID(.1)
(SO-mLtube)
Sample Vol. (mL)
Flag Preserved
O
Sampfe
Category *
OP
OD
Composite Volume (mL)
Chlorophyll (.2)
(GF/F filter)
Sample Vol. (mL)
Flag
Froz9fi
O
/ / 2 0
Primary ^° Sample Co lected O
Number of transects sampled (0-11):
Biomass (.3)
(GF/F Filter)
Sample Vol. (mL}
Flag
COMPOSITE PERIPHYTON SAMPLE
Sample ID
Assemblage I D(.1)
(50-mL tube)
Sample Vol. (mL)
Flag
Flag Preserved
O
Sample
Category *
OP
OD
Composite Volume (mL)
Chlorophyll (.2)
(GF/F filter)
Sample Vol. (mL)
Flag
Frozen
O
Frozen
O
APA (.4)
(50-mL tube)
Sample Vol. (mL)
Flag
Frozen
O
- Duplicate No Sample Collected Q
Number of transects sampled (0-11):
Biomass (.3) ,
(GF/F Filter) .
Sample Vol. (mL!
Flag
- ',
,F«zen
O
APA (.4)
(50-mL tube)
Sample- Vol. (mL)
Comments • - ••
Flag
Frozen
O
I
Flag codes: K = No measurement or observation made; U = S
flags in; comrngnt SQdiQns.
it or observation; F1, F2. etc. = flags assigned by field crew. Explain all
Sample ID
Sampte
Category *
OP
OD
OP
OD
Composite Volume
•„
stfll
TrAmacts
f
WENT CHEMISTRY / ENZYMES No Sample Collected Q
Chilled
O
o
Comments
ENTEROCOCCI (Target Volume = 250 mL) No Sample Collected Q
Sample ID
One unique ID per line
Flag
Sample
Cate-
gory*
QP
OD
OP
OD
OP
OD
OF
Time"
Collected
(hhmm)
Depth
Collected
(m)
Sample
Volume
(mL)
Filt Start
Time
(hhmm)
Volume Filtered
(Target = 50 mL) "
Flit. 1
Flit. 2
Flit. 3
Flit. 4
Filt. End
Time
(hhmm)
Time
Frozen
(hhmm)
Flag
Comment
* Sample Categories: P = Primary; D = Duplicate: F = Filter Blank (Enterococci sample only) Filter blank is coliected at visit '/.tiere field duplicate sample is NOT taken.
** ff <2S ml of buffer solution '/vas used to rinse filter, indicate with an F flag and note in comment section tttiich filters) v*ere affected along 'Aith the
approximate volume(s) of buffer solution used.
^™ NRSA Sample Collection - Wadeable 03/06/2008
Figure 6.1-3. Sample Collection Form, Side 2.
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6.1.3.3 Sampling Procedure
Near each of the macroinvertebrate and periphyton sampling locations, collect a fine-
grained sediment sample using either a hand scoop or spoon sampler. The objective is to
collect a 500-mL composite sample that is representative of depositional areas at the site. The
composite sample will be subsampled in the lab for multiple analyses. Table 6.1-6 presents
step-by-step procedures for collecting sediment enzyme samples.
Table 6.1-6. Sampling procedure—sediment enzymes
1. Collect a sediment sample at each of the macroinvertebrate and periphyton sample locations. Make
sure each of the subsamples comprises an approximately equal portion of the total composite. It is
permissible to collect sediment between stations to insure a composite volume of at least 500 ml.
(Note any deviations from standard procedure in a comment.)
2. Locate sediment samples in areas or patches of fine-grained substrate (silty sand, silt, clay, muck) in
a zone bounded on the shore side by the apparent low-water mark from daily flow fluctuations and
bounded on the riverside by the 0.3-m (usually about mid-biceps) depth contour (recommended
maximum sample depth; deeper sampling may be possible). The low-water mark at a site can often
be detected by the presence of periphyton or attached filamentous algae just below the low-water
mark. If samples cannot be safely collected by wading at a station due to vertical banks or other
reason go to step 5.
3. Be sure to avoid the area that has just been kick sampled for macroinvertebrates. Sampling up-
stream from the kick sample location is recommended. If fine substrates are not present within 5 m
up- or downstream from the station, flag the station on the form.
4. If fine substrate is present, use a stainless steel spoon to collect a sample of about 50ml or one
spoonful from the top 5 cm of substrate. Place the sample in a clean bucket. Use gloves for handling
sediment. Do not assume rip rapped shorelines lack fine-grained sediment. Look for fines between
the large rocks.
5. Repeat steps 2-4 at each of the 11 littoral stations. Record the total number of replicates (stations)
included in the composite. Note in a comment the stations at which sediment was collected using a
non-wading method.
6. It is important that a sufficient sediment (not less than 500 mL) sample for analysis be collected. If
multiple stations have no fine sediment, it is permissible to collect extra sample at stations that do
have fine sediment or between stations. Be sure to note this in a comment.
7. Using the stainless steel spoon, thoroughly mix the composite sample and transfer 500 mL into the
500 mL plastic bottle. Place in a cooler with ice for final labeling and preservation.
8. Prepare a label for the sample jar. Using a fine-point indelible marker, fill in the site # and sample
date. Place the label on the jar and cover it with clear tape. Record the sample ID and other data on
sampling form. Place the sample on ice or in a refrigerator. Do not freeze sediment samples. The
sediment enzyme samples have a 2 week holding time.
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6.2 Physical Habitat Characterization—Wadeable Streams
Physical habitat in streams includes all those physical attributes that influence or provide
sustenance to organisms within the stream. The physical habitat of a stream varies naturally,
thus expectations differ even in the absence of anthropogenic disturbance. Within a given
physiographic-climatic region, stream drainage area and overall stream gradient are likely to be
strong natural determinants of many aspects of stream habitat. This is because of their
influence on discharge, flood stage, and stream power (the product of discharge times gradient).
Kaufmann (1993) identified seven general physical habitat attributes important in influencing
stream ecology:
Channel Dimensions
Channel Gradient
Channel Substrate Size and Type
Habitat Complexity and Cover
Riparian Vegetation Cover and
Structure
Anthropogenic Alterations
Channel-Riparian Interaction
The procedures are employed on a support reach length 40 times its baseflow wetted
width, as described in Section 4. Measurement points are systematically placed to statistically
represent the entire reach. Stream depth and wetted width are measured at very tightly spaced
intervals, whereas channel cross-section profiles, substrate, bank characteristics and riparian
vegetation structure are measured at larger intervals. Woody debris is tallied along the full
length of the sampling reach, and discharge is measured at one location. The tightly spaced
depth and width measures allow calculation of indices of channel structural complexity,
objective classification of channel units such as pools, and quantification of residual pool depth,
pool volume, and total stream volume.
6.2.1 Components of the Habitat Characterization
There are five components of the physical habitat characterization (Table 6.2-1).
Measurements are recorded on 11 copies of a two-sided field form, and separate forms for
recording slope and bearing measurements, recording observations concerning riparian legacy
(large) trees and alien invasive riparian plants, assessing the degree of channel constraint, and
recording evidence of debris torrents or recent major flooding. The thalweg profile is a
longitudinal survey of depth, habitat class, presence of deposits of soft/small sediments, and
presence of off-channel habitats at 100 equally spaced stations (150 in streams less than 2.5 m
wide) along the centerline between the two ends of the sampling reach. Thalweg refers to the
flow path of the deepest water in a stream channel. Wetted width is measured and substrate
size is evaluated at 21 equally spaced cross-sections (at 11 regular transects [A through K], and
10 supplemental cross-sections spaced midway between each of these). Data for the second
component, the woody debris tally, are recorded for each of 10 segments of stream located
between the 11 regular transects. The third component, the channel and riparian
characterization, includes measures and/or visual estimates of channel dimensions, substrate,
fish cover, bank characteristics, riparian vegetation structure, presence of large (legacy) riparian
trees, nonnative (alien) riparian plants, and evidence of human disturbances. These data are
obtained at each of the 11 equally-spaced transects established within the sampling reach. In
addition, measurements of the stream slope and compass bearing between stations are
obtained, providing information necessary for calculating reach gradient, residual pool volume,
and channel sinuosity. The fourth component, assessment of channel constraint, debris
torrents, and major floods, is an overall assessment of these characteristics for the whole reach,
and is undertaken after the other components are completed.
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Table 6.2-1. Components of physical habitat characterization
Component
Description
Thalweg Profile
(Section 6.2.4.1)
Measure maximum depth, classify habitat and pool-forming features, and
check presence of backwaters, side channels and loose, soft deposits of
sediment particles at 10-15 equally spaced intervals between each of 11
transects (100 or 150 individual measurements along entire reach).
Measure wetted width and evaluate substrate particle size classes at 11
cross-section transects and midway between them (21 width measurements
and substrate cross-sections).
Woody Debris Tally
(Section 6.2.4.2)
Between each of the channel cross-sections, tally large woody debris
numbers within and above the bankfull channel according to specified length
and diameter classes (10 separate tallies).
Channel and Riparian
Characterization
(Section 6.2.5)
At 11 transects (21 for substrate size) placed at equal intervals along reach:
Measure: channel cross-section dimensions, bank height, bank undercut
distance, bank angle, slope and compass bearing (backsight), and riparian
canopy density (densiometer).
Visually Estimate3: substrate size class and embeddedness; areal cover
class and type (e.g., woody trees) of riparian vegetation in Canopy, Mid-Layer
and Ground Cover; areal cover class offish concealment features, aquatic
macrophytes and filamentous algae.
Observe & Record3: Presence and proximity of human disturbances,
presence of large trees, and presence of invasive riparian plants.
Assessment of
Channel Constraint,
Debris Torrents, and
Major Floods
(Section 6.2.6)
After completing thalweg and transect measurements and observations,
identify features causing channel constraint, estimate the percentage of the
channel margin that is constrained for the whole reach, and estimate the ratio
of bankfull/valley width. Check evidence of recent major floods and debris
torrent scour or deposition.
Discharge
(Section 6.2.6.3)
Measure water depth and velocity at 0.6 depth at 15 to 20 equally spaced
intervals across one carefully chosen channel cross-section.
In very small streams, measure discharge by timing the passage of a
neutrally buoyant object through a segment whose cross-sectional area has
been estimated or by timing the filling of a bucket.
Substrate size class is estimated for a total of 105 particles taken at 5 equally-spaced points along each of 21 cross-
sections. Depth is measured and embeddedness estimated for the 55 particles located along the 11 regular transects A
through K. Cross-sections are defined by laying the surveyor's rod or tape to span the wetted channel. Woody debris is
tallied over the distance between each cross-section and the next cross-section upstream. Riparian vegetation and
human disturbances are observed 5m upstream and 5m downstream from the cross-section transect. They extend
shoreward 10m from left and right banks. Fish cover types, aquatic macrophytes, and algae are observed within the
channel 5m upstream and 5m downstream from the cross-section stations. These boundaries for visual observations
are estimated by eye.
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6.2.2 Habitat Sampling Locations within the Reach
Measurements are made at two scales of resolution along the length of the reach; the
results are later aggregated and expressed for the entire reach, a third level of resolution. Figure
6.2-1 illustrates the locations within the reach where data for the different components of the
physical habitat characterization are obtained. Many channel and riparian features are
characterized on 11 cross-sections and pairs of riparian plots spaced at 4 channel-width
intervals (i.e., transect spacing = 1/10th the total reach length). The thalweg profile
measurements must be spaced evenly over the entire support reach. In addition, they must be
sufficiently close together that they do not miss deep areas and major habitat units. Follow
these guidelines for choosing the increment between thalweg profile measurements:
• Channel Width < 2.5 m — increment = 1.0 m
• Channel Width 2.5 to 3.5 m — increment = 1.5 m
• Channel Width > 3.5 m — increment = 0.01 x (reach length)
Following these guidelines, make 150 evenly spaced thalweg profile measurements in
the smallest category of streams, 15 between each detailed channel cross-section. In all of the
larger stream sizes, you will make 100 measurements, 10 between each cross-section.
6.2.3 Logistics and Work Flow
The five components (Table 6.2-1) of the habitat characterization are organized into four
grouped activities:
1. Thalweg Profile and Large Woody Debris Tally (Section 6.2.4). Two people proceed
upstream from the downstream end of the sampling reach (see Figure 6.2-1) making
observations and measurements at the chosen increment spacing. One person is in
the channel making width and depth measurements, and determining whether
soft/small sediment deposits are present under his/her staff. The other person records
these measurements, classifies the channel habitat, records presence/absence of side
channels and off-channel habitats (e.g., backwater pools, sloughs, alcoves), and tallies
large woody debris. Each time this team reaches a flag marking a new cross-section
transect, they start filling out a new copy of the Thalweg Profile and Woody Debris
Form. They interrupt the thalweg profile and woody debris tallying activities to complete
data collection at each cross-section transect as it comes. When the crew member in
the water makes a width measurement at channel locations midway between regular
transects (i.e., A, B, K), she or he also locates and estimates the size class of the
substrate particles on the left channel margin and at positions 25%, 50%, 75%, and
100% of the distance across the wetted channel. Procedures for this substrate tally are
the same as for those at regular cross-sections, but data are recorded on the thalweg
profile side of the field form.
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Thalweg
profile
stations
Channel/Riparian
Cross-section..
ran sect
Intermediate transects (width and
substrate measurements only
Woody
Debris
Tally
(bet ween
transects)
Downstream end
of sampling reach
PRKOVP 8/06
Figure 6.2-1. Reach layout for physical habitat measurements (plan view).
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2. Channel/Riparian Cross-Sections (Section 6.2.5). One person proceeds with the
channel cross-section dimension, substrate, bank, and canopy cover measurements.
The second person records those measurements on the Channel/ Riparian Cross-
section Form while making visual estimates of riparian vegetation structure, instream
fish cover, and human disturbance specified on that form. They also make
observations to complete the riparian "legacy" tree field form. Slope is measured by
measuring the difference in elevation between each transect and bearing is determined
by backsighting to the previous transect. Supplementary points may need to be located
and flagged (using a different color) if the stream is extremely brushy, sinuous, or steep
to the point that you cannot sight for slope and bearing measures between two
adjacent transects.
The work flow for the thalweg profile and channel cross described above can be
modified by delaying the measurements for slope and bearing and the woody
debris tally until after reaching the upstream end of the reach. Backsighting and
wood tallies can be done on the way back down (Note that in this case, the slope
and bearing data form would have to be completed in reverse order).
3. Channel Constraint and Torrent Evidence (Section 6.2.6). After completing
observations and measurements along the thalweg and at all 11 transects, the field
crew completes the overall reach assessments of channel constraint and evidence of
debris torrents and major floods.
4. Stream Discharge. Discharge measurements are made after collecting the water
chemistry sample. They are done at a chosen optimal cross-section (but not
necessarily at a transect) near the X-site. However, do not use the electromagnetic
current meter close to where electrofishing is taking place. Furthermore, if a lot of
channel disruption is necessary and sediment must be stirred up, wait on this activity
until all chemical and biological sampling has been completed.
6.2.4 Thalweg Profile and Large Woody Debris Measurements
6.2.4.1 Thalweg Profile
Thalweg refers to the flow path of the deepest water in a stream channel. The thalweg
profile is a longitudinal survey of maximum flow path depth and several other selected
characteristics at 100 or 150 equally spaced points (termed stations) along the length of the
reach measured along the centerline of the channel. Data from the thalweg profile allows
calculation of indices of residual pool volume, stream size, channel complexity, and the relative
proportions of habitat types such as riffles and pools. One person walks upstream carrying a
fiberglass telescoping (1.5 to 7.5 m) surveyor's rod and a 1-m metric ruler (or a calibrated rod or
pole, such as a ski pole, shovel handle, wooden dowel, or old billiard cue). A second person on
the bank or in the stream carries a clipboard with 11 copies of the field data form.
The procedure for obtaining thalweg profile measurements is presented in Table 6.2-2. Record
data on the Thalweg Profile and Woody Debris Data Form as shown in Figure 6.2-2. Use the
surveyor's rod and a metric ruler or calibrated rod or pole to make the required depth and width
measurements at each station, and to measure off the distance between stations as you
proceed upstream. You may need to make minor adjustments to align each 10th measurement
to be one increment short of the next transect. In streams with average widths less than 2.5 m,
make thalweg measurements at 1-meter increments. Because the minimum reach length is set
at 150 meters, there will be 15 measurements on a field data form: Station 0 at the transect plus
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14 additional stations between it and the next transect upstream. Use the five extra lines on the
thalweg profile portion of the data form (Figure 6.2-2) to record these measurements.
Table 6.2-2. Thalweg profile procedure
1. Determine the increment distance between measurement stations based on the wetted width used to
determine the length of the reach. Using a laser rangefinder or surveyor's rod:
• For widths < 2.5 m, establish stations every 1 m (150 total).
• For widths > 2.5 and <3.5 m, establish stations every 1.5 m (100 total).
• For widths > 3.5 m, establish stations at increments equal to 0.01 times the reach length (100 total).
2. Complete the header information on the Thalweg Profile and Woody Debris Form, noting the transect
pair (downstream to upstream). Record the increment distance determined in Step 1 in the
INCREMENT field on the field data form.
3. Begin at the downstream end (station 0) of the first transect (transect A).
4. Measure the wetted width at station 0, and at either station 5 (if the stream width defining the reach
length is > 2.5 m), or station 7 (if the stream width defining the reach length is < 2.5 m). Wetted width
is measured across and over mid-channel bars and boulders. Record the width on the field data form
to the nearest 0.1 m. For streams with interrupted flow, where no water is in the channel at the station
or transect, record zeros for wetted width.
NOTE: If a mid-channel bar is present at a station where wetted width is measured, measure the wetted
width across and including the bar, but also measure the bar width and record it on the field data
form.
5. At station 5 or 7 (see above) classify the size of the bed surface particle at the tip of your depth
measuring rod at the left wetted margin and at positions 25%, 50%, 75%, and 100% of the distance
across the wetted width of the stream. This procedure is identical to the substrate size evaluation
procedure described for regular channel cross-sections (transects A - K), except that for these
midway supplemental cross-sections, substrate size is entered on the thalweg profile side of the field
form.
6. At each thalweg profile station, use a calibrated pole or rod to locate the deepest point within the
deepest flow path (the thalweg), which may not always be found at mid-channel (and may not always
be the absolute deepest point in every channel cross-section). Measure the thalweg depth to the
nearest cm from the substrate surface to the water surface, and record it on the thalweg profile form.
Read the depth on the side of the rod to avoid inaccuracies due to the wave formed by the rod in
moving water.
NOTE: For streams with interrupted flow - if there is no water at a transect, record zeros for depth.
NOTE: Obtain thalweg depths at all stations. If the thalweg is too deep to measure directly, stand in
shallower water and extend the surveyor's rod or pole at an angle to reach the thalweg.
Determine the angle by resting the clinometer on the upper surface of the rod and reading the
angle on the external scale of the clinometer. Leave the depth reading for the station blank, and
record a U flag to indicate a non-standard procedure was used. Record the water level on the rod
and the rod angle in the comments section of the field data form. For deeper depths, use the
same procedure with a taut string as the measuring device. Tie a weight to one end of a length of
string or fishing line, and toss the weight into the deepest channel location. Draw the string up
tight and measure the length of the line that is under water. Measure the string angle with the
clinometer exactly as done for the surveyor's rod. If a direct measurement cannot be obtained,
make the best estimate you can of the thalweg depth, and use a U flag to identify it as an
estimated measurement.
7. At the point where the thalweg depth is determined, observe if unconsolidated, loose (soft) deposits
of small diameter (<16mm) sediments are present directly beneath your ruler, rod, or pole. Soft/small
sediments are defined here as fine gravel, sand, silt, clay or muck readily apparent by "feeling" the
bottom with the rod. Record presence or absence in the SOFT/SMALL SEDIMENT field on the field data
form. Note: A thin coating of fine sediment or silty algae coating the surface of cobbles should not be
considered soft/small sediment. However, fine sediment coatings should be identified in the
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comments section of the field form when determining substrate size and type.
8. Determine the channel unit code and pool forming element codes for the station. Record these on the
field data form using the standard codes provided. For dry and intermittent streams, where no water
is in the channel, record habitat type as dry channel (DR).
9. If the station cross-section intersects a mid-channel bar, indicate the presence of the bar in the BAR
WIDTH field on the field data form.
10. Record the presence or absence of a side channel at the station's cross-section in the SIDE CHANNEL
field on the field data form.
Record the presence or absence of quiescent off-channel aquatic habitats, including sloughs, alcoves
and backwater pools in the BACKWATER column of the field form.
11. Proceed upstream to the next station, and repeat Steps 2 through 11.
12. Repeat Steps 2 through 12 until you reach the next transect. At this point complete Channel/ Riparian
measurements at the new transect (Section 6.2.5). Then prepare a new Thalweg Profile and Woody
Debris Form and repeat Steps 2 through 12 for each of the reach segments, until you reach the
upstream end of the sampling reach (transect K). At transect K, you will have completed 10 copies of
the Thalweg Profile and Woody Debris Form, one for each segment (A to 8, 8 to C, etc.).
Measure thalweg depths at all stations. Missing depths at the end of the reach (e.g., due
to the stream flowing into or out of a culvert or under a large pile of debris) can be tolerated, but
those in the middle of the reach are more difficult to deal with. Flag any missing measurements
using a K code and explain the reason in the comments section of the field data form. At points
where a direct depth measurement cannot be made, make your best estimate of the depth,
record it on the field form, and flag the value using a U code (nonstandard measurement),
explaining that it is an estimated value in the comments section of the field data form. Where the
thalweg points are too deep for wading, measure the depth by extending the surveyor's rod at
an angle to reach the thalweg point. Record the water level on the rod, and the rod angle, as
determined using the external scale on the clinometer (vertical = 90°). In analyzing these data
we calculate the thalweg depth as the length of the rod (or string) under water multiplied by the
trigonometric sine of the rod angle. (For example, if 3 meters of the rod are under water when
the rod held at 30 degrees (s/ne=0.5), the actual thalweg depth is 1.5 meters.) These
calculations are done after field forms are returned for data analysis. On the field form, crews
are required only to record the wetted length of the rod under the water, a U code in the flag
field (to indicate a nonstandard technique), and a comment to the right saying "depth taken at
an angle ofxx degrees." If a direct measurement of the thalweg depth is not possible, make the
best estimate you can of the depth, record it, and use a U flag and a comment to note it is an
estimated value.
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National Rivers and Streams Assessment
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Date: April 2009
Page 131
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page 132
At every thalweg station, determine by sight or feel whether deposits of soft/small
sediments are present on the channel bottom. These particles are defined as substrate equal to
or smaller than fine gravel (< 16 mm diameter). These soft/small sediments are different from
Fines described when determining the substrate particle sizes at the cross-section transects
(Section 6.2.5.2). If the channel bottom is not visible, determine if soft/small sediment deposits
are readily obvious by feeling the bottom with your boot, the surveyor's rod, or a calibrated rod
or pole.
Measure wetted width at each transect (station 0), and midway between transects
(station 5 for larger streams having 100 measurement points, or station 7 for smaller streams
having 150 measurement points). The wetted width boundary is the point at which substrate
particles are no longer surrounded by free water. Estimate substrate size for five particles
evenly spaced across each midway cross-section using procedures described for substrate at
regular cross-sections (Section 6.2.5.2), but at the supplemental cross-sections, only the size
class (not distance and depth) data are recorded.
While recording the width and depth measurements and the presence of soft/small
sediments, the second person evaluates and records the habitat class and the pool forming
element (Table 6.2-3) applicable to each of the 100 (or 150) measurement points along the
length of the reach. Make channel unit scale habitat classifications at the thalweg of the cross-
section. The habitat unit itself must meet a minimum size criteria in addition to the qualitative
criteria listed in Table 6.2-3. Before being considered large enough to be identified as a
channel-unit scale habitat feature, the unit should be at least as long as the channel is wide. For
instance, if there is a small deep (pool-like) area at the thalweg within a large riffle area, do not
record it as a pool unless it occupies an area about as wide or long as the channel is wide. If a
backwater pool dominates the channel, record PB as the dominant habitat unit class. If the
backwater is a pool that does not dominate the main channel, or if it is an off-channel alcove
or slough (large enough to offer refuge to small fishes), circle Yto indicate presence of a
backwater in the BACKWATER column of the field form, but classify the main channel habitat unit
type according to characteristics of the main channel. Sloughs are backwater areas having
marsh-like characteristics such as vegetation, and alcoves (or side pools) are deeper areas off
the main channel that are typically wide and shallow (Helm 1985, Bain and Stevenson 1999).
When trying to identify the pool forming element for a particular pool, remember that most pools
are formed at high flows, so you may need to look for elements that are dry at baseflow, but still
within the bankfull channel (e.g., boulders or large woody debris).
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Table 6.2-3. Channel unit and pool forming element categories
Class (Code)
Channel Unit Habitat Classes a
Description
Pools: Still water, low velocity, a smooth, glassy surface, usually deep compared to other parts of the
channel:
Plunge Pool (PP)
Trench Pool (PT)
Lateral Scour Pool (PL)
Backwater Pool (PB)
Impoundment Pool(PD)
Pool (P)
Glide (GL)
Riffle (Rl)
Rapid (RA)
Cascade (CA)
Falls (FA)
Dry Channel (DR)
Pool at base of plunging cascade or falls
Pool-like trench in the center of the stream
Pool scoured along a bank
Pool separated from main flow off the side of the channel (large enough to offer
refuge to small fishes). Includes sloughs (backwater with marsh characteristics
such as vegetation), and alcoves (a deeper area off a wide and shallow main
channel)
Pool formed by impoundment above dam or constriction.
Pool (unspecified type)
Water moving slowly, with a smooth, unbroken surface. Low turbulence.
Water moving, with small ripples, waves and eddies - waves not breaking,
surface tension not broken. Sound: babbling, gurgling.
Water movement rapid and turbulent, surface with intermittent Whitewater with
breaking waves. Sound: continuous rushing, but not as loud as cascade.
Water movement rapid and very turbulent over steep channel bottom. Much of
the water surface is broken in short, irregular plunges, mostly Whitewater.
Sound: roaring.
Free falling water over a vertical or near vertical drop into plunge, water turbulent
and white over high falls. Sound: from splash to roar.
No water in the channel, or flow is submerged under the substrate (hyporheic
flow).
a Note that in order for a channel habitat unit to be distinguished, it must be at least as wide or long as the channel is
wide (except for off channel backwater pools, which are noted as present regardless of size).
Code
N
W
R
B
F
WR, RW, RBW
OT
Categories of Pool-forming Elements0
Category
Not Applicable, Habitat Unit is not a pool
Large Woody Debris.
Rootwad
Boulder or Bedrock
Unknown cause (unseen fluvial processes)
Combinations
Other (describe in the comments section of field form)
In determining the pool forming element, remember that most pools are formed at high flows, so you may need to
look at features, such as large woody debris, that are dry at baseflow, but still within the bankfull channel.
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6.2.4.2 Large Woody Debris Tally
Large Woody Debris is defined here as woody material with a small end diameter of at
least 10 cm (4 in.) and a length of at least 1.5 m (5 ft.). The procedure for tallying LWD is
presented in Table 6.2-4. The tally includes all pieces of LWD that are at least partially in the
baseflow channel (Zone 1), in the bankfull channel (Zone 2, flood channel up to bankfull stage),
or spanning above the bankfull channel (Zone 3), as shown in Figure 6.2-3. The bankfull
channel is defined as the channel that is filled by moderate sized flood events that typically
recur every one to two years. LWD in or above the bankfull channel is tallied over the entire
length of the reach, including the area between the channel cross-section transects. Pieces of
LWD that are not at least partially within Zones 1, 2, or 3 are not tallied.
Table 6.2-4. Procedure for tallying large woody debris
Note: Tally pieces of large woody debris (LWD) within each segment of stream while the thalweg profile is
being determined. Include all pieces in the tally whose large end is found within the segment.
1. Scan the stream segment between the two cross-section transects where thalweg profile
measurements are being made.
2. Tally all LWD pieces within the segment that are at least partially within the bankfull channel. Determine
if a piece is LWD (small end diameter >10 cm [4 in.], and length >1.5m [5 ft.})
3. For each piece of LWD, determine the class based on the diameter of the large end (0.1 m to < 0.3 m,
0.3 m to <0.6 m, 0.6 m to <0.8 m, or >0.8 m), and the class based on the length of the piece (1.5m to
<5.0m, 5m to <15m, or>15m).
• If the piece is not cylindrical, visually estimate what the diameter would be fora piece of wood with
circular cross-section that would have the same volume.
• When estimating length, include only the part of the LWD piece that has a diameter >10 cm (4 in)
4. Place a tally mark in the appropriate diameter x length class tally box in the PIECES ALL/PART IN
BANKFULL CHANNEL section of the Thalweg Profile and Woody Debris Form.
5. Tally all LWD pieces within the segment that are not actually within the bankfull channel, but are at
least partially spanning (bridging) the bankfull channel. For each piece, determine the class based on
the diameter of the large end (0.1 m to < 0.3 m, 0.3 mto<0.6 m, 0.6 mto<0.8 m, or>0.8 m), and the
class based on the length of the piece (1.5 m to <5.0 m, 5 m to <15 m, or >15 m).
6. Place a tally mark for each piece in the appropriate diameter x length class tally box in the PIECES
BRIDGE ABOVE BANKFULL CHANNEL section of the Thalweg Profile and Woody Debris Form.
7. After all pieces within the segment have been tallied, write the total number of pieces for each diameter
x length class in the small box at the lower right-hand corner of each tally box.
8. Repeat Steps 1 through 7 for the next stream segment, using a new Thalweg Profile and Woody Debris
Form.
6.2.5 Channel and Riparian Measurements at Cross-Section Transects
6.2.5.1 Slope and Bearing
Measure bearing by sighting between transects (e.g., transect 6 and A, C and 6, etc.) as
shown in Figure 6.2-4. To measure the bearing between adjacent transects, follow the
procedure presented in Table 6.2-5. Record bearing data on the Slope and Bearing Form as
shown in Figure 6.2-5.
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Slope is typically measured by two people, one holding a surveyor's rod and the second
sighting through the surveyor's level. Be sure that the person is standing (or holding the marked
pole) at the water's edge holding the rod at the surface of the water. The intent is to get a
measure of the water surface slope, which may not necessarily be the same as the bottom
slope. The surveyor's level is leveled according to the manufacturer's recommendations which
is generally to adjust the three screw leveling feet until the bubble is centered. Level is checked
in all planes to be measured. If the level does not "self level" in all measured planes the user
should check the instruction manual for suggested options. Elevation readings are made at
each transect and the difference between each elevation reading is recorded as the change in
elevation. NOTE: Multiple transect elevations can often be made for each setup of the level, but
every time the transit is moved requires re-measuring the last transect elevation from the last
setup. You cannot use elevations from previous setups because the relative height of the transit
has changed.
BANKFULL CHANNEL WIDTH
ZONE
ZONE 4
WATER SURFACE AT
BANKFULLFLOW
WATER SURFACE
ZONE 2 ATBASEFLOW
PRK/DVP 8/06
Figure 6.2-3. Large woody debris influence zones (modified from Robison and Beschta, 1990).
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To calculate sinuosity from bearing measurements, it does not matter whether or not you
adjust your compass bearings for magnetic declination, but it is important that you are
consistent in the use of magnetic or true bearings throughout all the measurements you make
on a given reach. Note in the comments section of the Slope and Bearing Form which type of
bearings you are taking, so the measurements can be used to describe reach aspect. Also,
guard against recording reciprocal bearings (erroneous bearings 180 degrees from what they
should be). The best way to do this is to know where the primary (cardinal) directions are in the
field: (north [0 degrees], east [90 degrees], south [180 degrees], and west [270 degrees]), and
insure that your bearings "make sense."
As stated earlier, it may be necessary to set up intermediate (supplemental) slope and
bearing points between a pair of cross-section transects if you do not have direct line-of-sight
along (and within) the channel between stations (see Figure 6.2-4). This can happen if brush is
too heavy, or if there are sharp slope breaks or tight meander bends. If you would have to sight
across land to measure slope or bearing between two transects, then you need to make one or
more supplemental measurements (i.e., do not "short-circuit" a meander bend). Mark these
supplemental locations with a different color of plastic flagging than used for the cross-section
transects to avoid confusion. Record these supplemental slope and bearing measurements,
along with the proportion of the stream segment between transects included in each
supplemental measurement, in the appropriate sections of the Slope and Bearing Form (Figure
6-5). Note that the main slope and bearing observations are always downstream of
supplemental observations (i.e., from or to the downstream transect). Similarly, first
supplemental observations are always downstream of second supplemental observations.
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Short pole with
clinometer at
height h
Surveyor rod
with flagging at
height h
Upstream
Transect
Both poles must be at water's
surface or at same depth
Downstream
Transect
Bearing Measurements Between Transects
Backsight with
compass and
record
main slope
and bearing
measurements
and % of reach
Supplemental slope Backsight with
and bearing point compass and record
supplemental slope
and bearing
Measurements and
% of reach
Backsight
with compass
and record
main slope
and bearing
measurements
and % of reach
Figure 6.2-4. Channel slope and bearing measurements.
Because of ease of use, portability, and cost, hand-held clinometers were previously
used to determine slope. In this instance, the field crews will have access to more sophisticated
instrumentation (e.g., surveyor's level), and have field personnel who are experienced in the use
of these instruments. The Slope and Bearing Form (Figure 6-5) is designed to allow for different
methods and/or different units of measuring slope. Mark the appropriate method circle (instead
of CL; method codes are identified in Tables 6.2-5 and 6.2-6), and mark the CM circle (instead of
the % circle) if the method or instrument measures the change in elevation rather than the
percent slope.
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Table 6.2-5. Procedure for obtaining slope and bearing data
1. Determine a location at transect K to hold a surveyor's rod that will be visible from a point between
transect J and transect K:
a) Set up the instrument at a point approximately halfway between points J and K and where a
clear line of sight is possible.
b) Position the staff at point K, holding the bottom of the staff at the water level and the staff as
vertical as possible and the numbers facing the instrument.
c) Site the staff and record the reading to the nearest centimeter.
d) Move the staff to point J and gently swivel the instrument to face the next reading. Hold the staff
as before, vertically, with the bottom at the water level and the numbers facing the instrument.
e) Site the staff and record the reading to the nearest centimeter.
f) Repeat measurements between each transect.
g) The difference in the readings is the height difference or gradient.
Note: In small streams with a clear line of site it may be possible to set the instrument up once and make
readings to several transects from a single set up. Simply record the readings for each transect and
do not skip transects.
• If you are backsighting from a supplemental point, record the bearing in the appropriate SUPPLE-
MENTAL section of the Slope and Bearing Form.
2. Proceed to the next cross-section transect (or supplementary point), and repeat Steps a - g above.
Instrument Setup:
a) Extend the tripod legs to approximately eye level and set the legs firmly into the ground; adjust
the legs so that they form a regular triangle and are firmly set with no wobble. Adjust the legs so
that the base plate is approximately level.
b) Hold the instrument on the tripod and start the centering screw. Ensure the adjustable feet are
roughly evenly adjusted. While the centering screw is still loose slide the instrument on the base
plate until the bubble is approximately centered in the circular level. Tighten the centering screw.
c) Adjust the leveling foot screws until the bubble is exactly level in the center circle.
d) Self Leveling instruments can now be swiveled gently on the base plate and maintain level as
long as the tripod remains steady.
e) Adjust focus, brightness and parallax according to manufactures specifications.
f) The instrument is ready to make measurements.
a Method codes are: CL=clinometer, 7f?=transit, /-/L=hand level, l/l/T=Watertube, L4=laser level, OrHER=method not
listed (describe in comments section of form).
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Table 6.2-6. Modified procedure for obtaining slope and bearing data
Use this procedure if you are starting at the upstream transect (K), after completing the thalweg
profile and other cross-section measurements at transects A through K.
1. Stand in the center of the channel at the upstream cross-section transect. Determine if you
can see the center of the channel at the next cross-section transect downstream without
sighting across land (i.e., do not "short-circuit" a meander bend). If not, you will have to take
supplementary slope and bearing measurements.
Mark a surveyor's rod and a calibrated rod (or meter ruler) at the same height. If a shorter pole
or ruler is used, measure the height from the ground to the opening of the clinometer when it is
resting on top.
2. Have one person take the marked surveyor's rod to the downstream transect. Hold the rod
vertical with the bottom at the same level as the water surface. If no suitable location is
available at the stream margin, position the rod in the water and note the depth.
• If you have determined in Step 1 that supplemental measurements are required for this
segment, walk downstream to the furthest point where you can stand in the center of the
channel and still see the center of the channel at the upstream cross-section transect.
Remember that your line of sight cannot "cross land." Mark this location with a different
color flagging than that marking the cross-section transects.
3. Place the base of the calibrated rod at the level as the surveyor's rod (either at the water
surface or at the same depth in the water).
4. Place the clinometer on the calibrated rod at the height determined in Step 2. With the
clinometer, sight back downstream to the flagged height on the surveyor's rod at the down-
stream transect (or at the supplementary point).
• If you are sighting to the next downstream transect, read and record the percent slope in
the MAIN section on the Slope and Bearing Form for the downstream transect (e.g., J <
K), which is at the bottom of the form (i.e., you are completing the form in reverse order).
Record the PROPORTION as 100%.
• If you are backsighting from a supplemental point, record the slope (%) and proportion
(%) of the stream segment that is included in the measurement in the appropriate
SUPPLEMENTAL section of the Slope and Bearing Form. The last sighting to a downstream
transect (from either the upstream transect or the nearest upstream supplemental point)
is always recorded as the MAIN reading.
5. Stand in the middle of the channel at upstream transect (or at a supplemental point), and sight
with your compass to the middle of the channel at the downstream transect (or at a
supplemental point). Record the bearing (degrees) in the same section of the Slope and
Bearing form (Supplemental or Main) as you recorded the slope in Step 6.
6. Proceed to the next cross-section transect (or to a supplementary point), and repeat Steps 3
through 7 above.
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Page 141
6.2.5.2 Substrate Size and Channel Dimensions
Substrate size and embeddedness are evaluated at 5 points at each of the 11 transects
(refer to Figure 6.2-6). Substrate size is also evaluated at 10 additional cross-sections located
midway between each of the 11 regular transects (A-K). In the process of measuring substrate
particle sizes at each channel cross-section, the wetted width of the channel and the water
depth at each substrate sample point are measured (at the 10 midway cross-sections, only
substrate size and wetted width are recorded). If the wetted channel is split by a mid-channel
bar (see Section 6.2.4.1), the five substrate points are centered between the wetted width
boundaries regardless of the mid-channel bar in between. Consequently, substrate particles
selected in some cross-sections may be "high and dry". For cross-sections that are entirely dry,
make measurements across the unvegetated portion of the channel.
Left
Bank
Figure 6.2-6. Substrate sampling cross-section.
The substrate sampling points along the cross-section are located at 0, 25, 50, 75, and
100 percent of the measured wetted width, with the first and last points located at the water's
edge just within the left and right banks. The procedure for obtaining substrate measurements is
described in Table 6.2-7 (including all particle size classifications). Record these measurements
on the Channel/Riparian Cross-section side of the field form, as shown in Figure 6.2-7. For the
supplemental cross-sections midway between regular transects,
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National Rivers and Streams Assessment
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Table 6.2-7. Substrate measurement procedure
1. Fill in the header information on page 1 of a Channel/Riparian Cross-section Form. Indicate the cross-
section transect. At the transect, extend the surveyor's rod or metric tape across the channel
perpendicular to the flow, with the "zero" end at the left bank (facing downstream).
NOTE: If a side channel is present, and contains 16- 49% of the total flow, establish a secondary cross-
section transect. Use a separate field data form to record data for the side channel, designating it
as a secondary transect by marking both the X-TRA SIDE CHANNEL circle and the associated
primary transect letter (e.g., XA, XB, etc.). Collect all channel and riparian cross-section
measurements from the side channel.
2. Divide the wetted channel width channel by 4 to locate substrate measurement points on the cross-
section. In the DISTLB fields of the form, record the distances corresponding to 0% (/.FT), 25% (LCTR),
50% (CfR), 75% (RCTR), and 100% (Ref) of the measured wetted width. Record these distances at
Transects A-K, but just the wetted width at midway cross-sections.
3. Place your sharp-ended meter stick or calibrated pole at the LFT location (0 m). Measure the depth
and record it on the field data form. (Cross-section depths are measured only at regular transects A-K,
not at the 10 midway cross-sections).
• Depth entries at the left and right banks may be 0 (zero) if the banks are gradual.
• If the bank is nearly vertical, let the base of the measuring stick fall to the bottom (i.e., the depth
at the bank will be > 0 cm), rather than holding it suspended at the water surface.
4. Pick up the substrate particle that is at the base of the meter stick (unless it is bedrock or boulder), and
visually estimate its particle size, according to the following table. Classify the particle according to its
median diameter (the middle dimension of its length, width, and depth). Record the size class code
on the field data form. (Cross-section side of form for transects A-K; special entry boxes on Thalweg
Profile side of form for midway cross-sections.)
Code
RS
RR
HP
LB
SB
CB
GC
GF
SA
FN
WD
RC
OT
Size Class
Bedrock (Smooth)
Bedrock (Rough)
Hard pan
Boulders (large)
Boulders (small)
Cobbles
Gravel (Coarse)
Gravel (Fine)
Sand
Fines
Wood
Concrete
Other
Size Range (mm)
>4000
>4000
>4000
>1 000 to 4000
>250to1000
>64 to 250
>16to64
> 2 to 16
>0.06to2
<0.06
Regardless of Size
Regardless of size
Regardless of Size
Description
Smooth surface rock bigger than a car
Rough surface rock bigger than a car
Firm, consolidated fine substrate
Yard/meter stick to car size
Basketball to yard/meter stick size
Tennis ball to basketball size
Marble to tennis ball size
Ladybug to marble size
Smaller than ladybug size - gritty between fingers
Silt Clay Muck (not gritty between fingers)
Wood & other organic particles
Record size class in comment field
Metal, tires, car bodies etc. (describe in comments)
5. Evaluate substrate embeddedness as follows at each transects. For particles larger than sand, examine
the surface for stains, markings, and algae. Estimate the average % embeddedness of particles in the
10 cm circle around the measuring rod. Record this value on the field data form. For sand and smaller
particles, you will not be able to pick up an individual particle, but a "pinch" of fine particles between your
fingers. Determine and record the dominant size of particles in the "pinch." By definition, sand and fines
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are embedded 100%; bedrock and hardpan are embedded 0%.
6. Move to the next location on the transect, and repeat Steps 4 - 6 at each location. Repeat Steps 1 - 6
at each transect, including any additional side channel transects established if islands are present.
record substrate size and wetted width data on the thalweg profile side of the field form. To
minimize bias in selecting a substrate particle for size classification, it is important to
concentrate on correct placement of the measuring stick along the cross-section, and to select
the particle right at the bottom of the stick (not, for example, a more noticeable large particle that
is just to the side of the stick). Classify the particle into one of the size classes listed on the field
data form (Figure 6.2-7) based on the middle dimension of its length, width, and depth. This
median dimension determines the sieve size through which the particle can pass. When you
record the size class as Other, assign an Fn flag on the field data form and describe the
substrate type in the comments section of the field form, as shown in Figure 6.2-7.
At substrate sampling locations on the 11 regular transects (A-K), examine particles
larger than sand for surface stains, markings, and algal coatings to estimate embeddedness of
all particles in the 10 cm diameter circle around the substrate sampling point. Embeddedness is
the fraction of a particle's volume that is surrounded by (embedded in) sand or finer sediments
on the stream bottom. By definition, record the embeddedness of sand and fines (silt, clay, and
muck) as 100 percent, and record the embeddedness of hardpan and bedrock as 0 percent.
6.2.5.3 Bank Characteristics
The procedure for obtaining bank and channel dimension measurements is presented in
Table 6.2-8. Data are recorded in the BANK MEASUREMENTS section of the Channel/Riparian
Cross-section Form as shown in Figure 6.2-7. Bank angle and bank undercut distance are
determined on the left and right banks at each cross-section transect. Figure 6.2-8 illustrates how
bank angle is determined for several different situations. The scale at which bank angle is
characterized is approximately 0.5 m. A short (approx. 1-m long) pole is used to determine bank
angle. The angle is determined based on the pole resting on the ground for about 0.5 m. Other
features include the wetted width of the channel (as determined in Section 6.2.5.2), the width of
exposed mid-channel bars of gravel or sand, estimated incision height, and the estimated height
and width of the channel at bankfull stage as described in Table 6-8. Bankfull height and incised
height are both measured relative to the present water surface (i.e. the level of the wetted edge
of the stream). This is done by placing the base of the small measuring rod at the bankfull
elevation and sighting back to the survey rod placed at the water's edge using the clinometer as
a level (i.e., positioned so the slope reading is 0%.). The height of the clinometer above the base
of the smaller rod is subtracted from the elevation sighted on the surveyor's rod.
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National Rivers and Streams Assessment Final Manual
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Table 6.2-8. Procedure for measuring bank characteristics
1. To measure bank angle, lay a meter ruler or a short (approx. 1-m long) rod down against the left bank
(determined as you face downstream), with one end at the water's edge. At least 0.5 m of the ruler or
rod should be resting comfortably on the ground to determine bank angle. Lay the clinometer on the
rod, and read the bank angle in degrees from the external scale on the clinometer. Record the angle in
the field for the left bank in the BANK MEASUREMENT section of the Channel/Riparian Cross-section
Form.
• A vertical bank is 90°, overhanging banks have angles >90° approaching 180°, and more
gradually sloped banks have angles <90°. To measure bank angles >90°, turn the clinometer
(which only reads 0 to 90°) over and subtract the angle reading from 180°.
• If there is a large boulder or log present at the transect, measure bank angle at a nearby point
where conditions are more representative.
2. If the bank is undercut, measure the horizontal distance of the undercutting to the nearest 0.01 m. The
undercut distance is the distance from the water's edge out to the point where a vertical plumb line
from the bank would hit the water's surface. Record the distance on the field data form. Measure
submerged undercuts by thrusting the rod into the undercut and reading the length of the rod that is
hidden by the undercutting.
3. Repeat Steps 1 and 2 on the right bank.
4. Hold the surveyor's rod vertical, with its base planted at the water's edge. Examine both banks, then
determine the channel incision as the height up from the water surface to elevation of the first terrace
of the valley floodplain (Note this is at or above the bankfull channel height). Whenever possible, use
the clinometer as a level (positioned so it reads 0% slope) to measure this height by transferring
(backsighting) it onto the surveyor's rod. Record this value in the INCISED HEIGHT field of the bank
measurement section on the field data form.
5. While still holding the surveyor's rod as a guide, and sighting with the clinometer as a level, examine
both banks to measure and record the height of bankfull flow above the present water level. Look for
evidence on one or both banks such as:
• An obvious slope break that differentiates the channel from a relatively flat floodplain terrace
higher than the channel.
• A transition from exposed stream sediments to terrestrial vegetation.
• Moss growth on rocks along the banks.
• Presence of drift material caught on overhanging vegetation.
• A transition from flood- and scour-tolerant vegetation to that which is relatively intolerant of these
conditions.
6. Record the wetted width value determined when locating substrate sampling points in the WETTED
WIDTH field in the bank measurement section of the field data form. Also determine the bankfull
channel width and the width of exposed mid-channel bars (if present). Record these values in the
BANK MEASUREMENT section of the field data form.
7. Repeat Steps 1 through 6 at each cross-section transect, (including any additional side channel
transects established when islands are present). Record data for each transect on a separate field
data form.
Bankfull flows are large enough to erode the stream bottom and banks, but frequent enough
(every 1 to 2 years) to not allow substantial growth of upland terrestrial vegetation.
Consequently, in many regions, it is these flows that have determined the width and depth of the
channel. Estimates of the bankfull dimensions of stream channels are extremely important in
EMAP surveys. They are used to calculate shear stress and bed stability (see Kaufmann et al.,
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1999). Unfortunately, we have to depend upon evidence visible during the low-flow sampling
season. If available, consult published rating curves relating expected bankfull channel
dimensions to stream drainage area within the region of interest. Graphs of these rating curves
can help you get a rough idea of where to look for field evidence to determine the level of
bankfull flows. Curves such as these are available from the USGS for streams in most regions
of the U.S. (e.g., Dunne and Leopold 1978; Harrelson et al. 1994, Leopold 1994). To use them,
you need to know the contributing drainage area to your sample site. Interpret the expected
bankfull levels from these curves as a height above the streambed in a riffle, but remember that
your field measurement will be a height above the present water surface of the stream. Useful
resources to aid your determination of bankfull flow levels in streams in the United States are
video presentations produced by the USDA Forest Service for western streams (USDA Forest
Service 1995) and eastern streams (USDA Forest Service 2002).
Bank Angle= Qnometer rearing
(A)
Pole is reslhg "comfortably'
from wetted edge
Pole lestsmost
'comfortably" -
here
(B)
Anole^ Cfriometer leadng
Too much space under
pole I measured from
water's edge
Shelf is not wide enough to
use for deteimnng baric angle
Bank AKje=Qnanieter rearing
Pdejs'comtoffctter
from waters edge
Not enough undercut
exposed to define
oveihangng bank
Bar* Angle=180* - Cf norneter leading
Figure 6.2-8. Determining bank angle under different types of bank conditions. (A) typical, (B)
incised channel, (C) undercut bank, and (D) overhanging bank.
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After consulting rating curves that show where to expect bankfull levels in a given size of
stream, estimate the bankfull flow level by looking at the following indicators:
• First look at the stream and its valley to determine the active floodplain. This is a
depositional surface that frequently is flooded and experiences sediment deposition
under the current climate and hydrological regime.
• Then look specifically for:
• An obvious break in the slope of the banks.
• A change from water-loving and scour-tolerant vegetation to more drought-tolerant
vegetation.
• A change from well-sorted stream sediments to unsorted soil materials.
In the absence of clear bankfull indications, consider the previous season's flooding as the best
evidence available (note: you could be wrong if very large floods or prolonged droughts have
occurred in recent years.). Look for:
• Drift debris ("sticky wickets" left by the previous seasons flooding).
• The level where deciduous leaf-fall is absent on the ground (carried away by
previous winter flooding).
• Unvegetated sand, gravel or mud deposits from previous year's flooding.
In years that have experienced large floods, drift material and other recent high flow
markers may be much higher than other bankfull indicators. In such cases, base your
determination on less-transient indicators such as channel form, perennial vegetation, and
depositional features. In these cases, flag your data entry and also record the height of drift
material in the comments section of the field data form.
We use the vertical distance (height) from the observed water surface up to the level of
the first major valley depositional surface (Figure 6.2-9) as a measure of the degree of incision
or downcutting of the stream below the general level of its valley. This value is recorded in the
INCISED HEIGHT field. It may not be evident at the time of sampling whether the channel is
downcutting, stable, or aggrading (raising its bed by depositing sediment). However, by
recording incision heights measured in this way and monitoring them over time, we will be able
to tell if streams are incising or aggrading.
If the channel is not greatly incised, bankfull channel height and incision height will be
the same (i.e., the first valley depositional surface is the active floodplain). However, if the
channel is incised greatly, the bankfull level will be below the level of the first terrace of the
valley floodplain, making bankfull channel height less than incision height (Figure 6.2-10).
Bankfull height is never greater than incision height. You may need to look for evidence of
recent flows (within about one year) to distinguish bankfull and incision heights. In cases where
the channel is cutting a valley sideslope and has oversteepened and destabilized that slope, the
bare "cutbank" against the steep hillside at the edge of the valley is not necessarily an indication
of recent incision. In such a case, the opposite bank may be lower, with a more obvious terrace
above bankfull height; choose that bank for your measurement of incised height. Examine both
banks to more accurately determine incision height and bankfull height. Remember that incision
height is measured as the vertical distance to the first major depositional surface above bankfull
(whether or not it is an active floodplain or a terrace. If terrace heights differ on left and right
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banks (both are above bankfull), choose the lower of the two terraces. In many cases your
sample reach may be in a "V" shaped valley or gorge formed over eons, and the slope of the
channel banks simply extends uphill indefinitely, not reaching a terrace before reaching the top
of a ridge (Figure 6.2-10). In such cases, record incision height values equal to bankfull values
and make appropriate comment that no terrace is evident. Similarly, when the stream has
extremely incised into an ancient terrace, (e.g., the Colorado River in the Grand Canyon), you
may crudely estimate the terrace height if it is the first one above bankfull level. If you cannot
estimate the terrace height, make appropriate comments describing the situation.
A. Channel not Incised
Downcutting over
geologic time
Active
floodplain at or near
valley bottom elevation
(Record this height)
First terrace on
valley bottom
above bankfull
level
Second
terrace
No recent incision- bankfull
level at valley bottom
Valley Fill
B. Incised Channel
Downcutting over
geologic time
Former second
terrace becomes
Former active floodplain Former first third terrace
no longer connected— terrace becomes
becomes new first terrace second terrace
above bankfuil level
(Record this height)
Recent incision—
bankfull level below
first terrace of valley
bottom
Valley Fill
Figure 6.2-9. Schematic showing relationship between bankfull channel and incision. (A) not
recently incised, and (B) recently incised into valley bottom. Note level of bankfull stage relative to
elevation of first terrace (abandoned floodplain)on valley bottom. (Stick figure included for scale).
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A) Deeply Incised Channel
Hill Slope
Incision Height (Always
equal to or greater than
bankfuii height)
Second Terrace
First Terrace
1
f
T
-
_ _ _ _
vO
1 T' '
Jk
i— Bankfuii
/ Height
/ (When
/ channel form
' is not a good
indicator, use
evidence of
recent
floodinq)
B) Small stream constrained in V-shaped valley
Rood-
ntolerant
vegetation
Bankfuii Height
(when channel form is
not a good indicator,
use evidence of recent [~|
flooding, lack of
permanent flood-
intolerant vegetation
No incision:
No evidence of
downcutting,
vertical bank
angle, etc.)
Incision Hekjlrt=
Bankhil Height
Figure 6.2-10. Determining bankfuii and incision heights for (A) deeply incised channels, and (B)
streams in deep V-shaped valleys. (Stick figure included for scale).
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6.2.5.4 Canopy Cover Measurements
Canopy cover over the stream is determined at each of the 11 cross-section transects. A
spherical densiometer (model A- convex type) is used (Lemmon 1957). Mark the densiometer
with a permanent marker or tape exactly as shown in Figure 6.2-11 to limit the number of
square grid intersections read to 17. Densiometer readings can range from 0 (no canopy cover)
to 17 (maximum canopy cover). Six measurements are obtained at each cross-section transect
(four measurements in each of four directions at mid-channel and one at each bank).
TAPE
BUBBLE LEVELED'
Figure 6.2-11. Schematic of modified convex spherical canopy densiometer. From Mulvey et al.
(1992). Note proper positioning with the bubble leveled and face reflected at the apex of the "V". In this
example, 10 of the 17 intersections show canopy cover, giving a densiometer reading of 10.
The procedure for obtaining canopy cover data is presented in Table 6.2-9. Hold the
densiometer level (using the bubble level) 0.3 m above the water surface with your face
reflected just below the apex of the taped "V", as shown in Figure 6.2-11. Concentrate on the 17
points of grid intersection on the densiometer that lie within the taped "V". If the reflection of a
tree or high branch or leaf overlies any of the intersection points, that particular intersection is
counted as having cover. For each of the six measurement points, record the number of
intersection points (maximum=17) that have vegetation covering them in the CANOPY COVER
MEASUREMENT section of the Channel/Riparian Cross-section Form as shown in Figure 6.2-7.
Table 6.2-9. Procedure for canopy cover measurements
1. At each cross-section transect, stand in the stream at mid-channel and face upstream.
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2. Hold the densiometer 0.3 m (1 ft) above the surface of the stream. Level the densiometer using the
bubble level. Move the densiometer in front of you so your face is just below the apex of the taped
"V".
3. Count the number of grid intersection points within the "V" that are covered by either a tree, a leaf, or
a high branch. Record the value (0 to 17) in the CENUP field of the canopy cover measurement
section of the Channel/Riparian Cross-section and Thalweg Profile Form.
4. Face toward the left bank (left as you face downstream). Repeat Steps 2 and 3, recording the value
in the CEA/L field of the field data form.
5. Repeat Steps 2 and 3 facing downstream, and again while facing the right bank (right as you look
downstream). Record the values in the CENDWN and CENR fields of the field data form.
6. Move to the water's edge (either the left or right bank). Repeat Steps 2 and 3 again, this time facing
the bank. Record the value in the LFT or RGT fields of the field data form. Move to the opposite bank
and repeat.
7. Repeat Steps 1 through 6 at each cross-section transect (including any additional side channel
transects established when islands are present). Record data for each transect on a separate field
data form.
6.2.5.5 Riparian Vegetation Structure
The previous section (6.2.5.4) described methods for quantifying the cover of canopy
over the stream channel. The following visual estimation procedures supplement those
measurements with a semi-quantitative evaluation of the type and amount of various types of
riparian vegetation. Additional measures within the riparian zone (legacy trees and invasive
riparian plants) are described in Section 6.2.5.9.
Riparian vegetation observations apply to the riparian area upstream 5 meters and
downstream 5 meters from each of the 11 cross-section transects (refer to Figure 6.2-1). They
include the visible area from the stream back a distance of 10m (-30 ft) shoreward from both
the left and right banks, creating a10mx10m riparian plot on each side of the stream (Figure
6.2-12). The riparian plot dimensions are estimated, not measured. On steeply sloping channel
margins, the 10 m x 10m plot boundaries are defined as if they were projected down from an
aerial view.
Table 6.2-10 presents the procedure for characterizing riparian vegetation structure and
composition. Figure 6.2-7 illustrates how measurement data are recorded on the
Channel/Riparian Cross-section Form. Conceptually divide the riparian vegetation into 3 layers:
the Canopy layer (> 5 m high), the Understory layer (0.5 to 5 m high), and the Ground cover
layer (< 0.5 m high). Note that several vegetation types (e.g., grasses or woody shrubs) can
potentially occur in more than one layer. Similarly note that some things other than vegetation
are possible entries for the Ground cover layer (e.g., barren ground).
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10m
10m
10m
RIPARIAN
PLOT
(Left Bank)
Cross-sectibn Transect
Instream Fish
Cover Plot
RIPARIAN
PLOT
(Right Bank)
10m
Figure 6.2-12. Riparian zone and instream fish cover plots for a stream cross-section transect.
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Table 6.2-10. Procedure for characterizing riparian vegetation structure
1. Standing in mid-channel at a cross-section transect, estimate a 5 m distance upstream and
downstream (10m total length).
2. Facing the left bank (left as you face downstream), estimate a distance of 10 m back into the riparian
vegetation.
On steeply-sloping channel margins, estimate the distance into the riparian zone as if it were
projected down from an aerial view.
3. Within this 10 m x 10 m area, conceptually divide the riparian vegetation into 3 layers: a Canopy
Layer (>5 m high), an Understory (0.5 to 5 m high), and a Ground Cover layer (<0.5 m high).
4. Within this 10 m x 10 m area, determine the dominant vegetation type for the CANOPY LAYER
(vegetation >5 m high) as either Deciduous, Coniferous, broadleaf Evergreen, Mixed, or None.
Consider the layer Mixed if more than 10% of the areal coverage is made up of the alternate
vegetation type. Indicate the appropriate vegetation type in the VISUAL RIPARIAN ESTIMATES section of
the Channel/Riparian Cross-section Form.
5. Determine separately the areal cover class of large trees (>0.3 m [1 ft] diameter at breast height
[dbh]) and small trees (<0.3 m dbh) within the canopy layer. Estimate areal cover as the amount of
shadow that would be cast by a particular layer alone if the sun were directly overhead. Record the
appropriate cover class on the field data form (0=absent zero cover, 1=sparse: <10%, 2=moderate:
10-40%, 3=heavy: 40-75%, or 4=very heavy: >75%).
6. Look at the UNDERSTORY layer (vegetation between 0.5 and 5 m high). Determine the dominant
woody vegetation type for the understory layer as described in Step 4 for the canopy layer. If there is
no woody vegetation in the understory layer, record the type as None.
7. Determine the areal cover class for woody shrubs and saplings separately from non-woody
vegetation within the understory, as described in Step 5 for the canopy layer.
8. Look at the GROUND COVER layer (vegetation <0.5 m high). Determine the areal cover class for
woody shrubs and seedlings, non-woody vegetation, and the amount of bare ground present as
described in Step 5 for large canopy trees.
9. Repeat Steps 1 through 8 for the right bank.
10. Repeat Steps 1 through 9 for all cross-section transects (including any additional side channel
transects established when islands are present). Use a separate field data form for each transect.
Before estimating the areal coverage of the vegetation layers, record the type of woody
vegetation (broadleaf Deciduous, Coniferous, broadleaf Evergreen, Mixed, or None) in each of
the two taller layers (Canopy and Understory). Consider the layer Mixed if more than 10% of the
areal coverage is made up of the alternate vegetation type. If there is no woody vegetation in
the understory layer, record the type as None.
Estimate the areal cover separately in each of the three vegetation layers. Note that the
areal cover can be thought of as the amount of shadow cast by a particular layer alone when
the sun is directly overhead. The maximum cover in each layer is 100%, so the sum of the areal
covers for the combined three layers could add up to 300%. The four areal cover classes are
Absent, Sparse (<10%), Moderate (10 to 40%), Heavy (40 to 75%), and Very Heavy (>75%).
These cover classes and their corresponding codes are shown on the field data form (Figure
6.2-7). When rating vegetation cover types for a single vegetation layer, mixtures of two or
more subdominant classes might all be given Sparse (1), Moderate (2), or Heavy (3) ratings.
One Very Heavy cover class with no clear subdominant class might be rated 4 with all the
remaining classes rated as either Moderate (2), Sparse (1) or Absent (0). Note that within a
given vegetation layer, two cover types with 40-75% cover can both be rated 3, but no more
than one cover type could receive a rating of 4.
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page 154
6.2.5.6 In stream Fish Cover, Algae, and Aquatic Macrophytes
The procedure to estimate the types and amounts of instream fish cover is outlined in
Table 6.2-11. Data are recorded on the Channel/Riparian Cross-section Form as shown in
Figure 6.2-7. Estimate the areal cover of all of the fish cover and other listed features that are in
the water and on the banks 5 m upstream and downstream of the cross-section (see Figure 6.2-
12). The areal cover classes of fish concealment and other features are the same as those
described for riparian vegetation (Section 6.2.5.5).
The entry FILAMENTOUS ALGAE refers to long streaming algae that often occur in slow
moving waters. AQUATIC MACROPHYTES are water-loving plants, including mosses, in the stream
that could provide cover for fish or macroinvertebrates. If the stream channel contains live
wetland grasses, include these as aquatic macrophytes. WOODY DEBRIS are the larger pieces of
wood that can influence cover and stream morphology (i.e., those pieces that would be included
in the large woody debris tally [Section 6.2.4]). BRUSH/WOODY DEBRIS refers to smaller wood
pieces that primarily affect cover but not morphology. LIVE TREES OR ROOTS are living trees that
are within the channel - estimate the areal cover provided by the parts of these trees or roots
that are inundated. OVERHANGING VEGETATION includes tree branches, brush, twigs, or other
small debris that is not in the water but is close to the stream (within 1 m of the surface) and
provides potential cover. BOULDERS are typically basketball- to car-sized particles. ARTIFICIAL
STRUCTURES include those designed for fish habitat enhancement, as well as in-channel
structures that have been discarded (e.g., concrete, asphalt, cars, or tires) or deliberately placed
for diversion, impoundment, channel stabilization, or other purposes.
Table 6.2-11. Procedure for estimating instream fish cover
1. Standing mid-channel at a cross-section transect, estimate a 5m distance upstream and downstream
(10 m total length).
2. Examine the water and both banks within the 10-m segment of stream for the following features and
types of fish cover: filamentous algae, aquatic macrophytes, large woody debris, brush and small
woody debris, in-channel live trees or roots, overhanging vegetation, undercut banks, boulders, and
artificial structures.
3. For each cover type, estimate the areal cover. Record the appropriate cover class in the FISH
COVER/OTHER section of the Channel/Riparian Cross-section Form:
0=absent zero cover,
1=sparse: <10%,
2=moderate: 10-40%,
3=heavy: >40-75%, or
4=very heavy: >75%).
4. Repeat Steps 1 through 3 at each cross-section transect (including any additional side channel
transects established when islands are present). Record data from each transect on a separate field
data form.
6.2.5.7 Human Influence
For the left and right banks at each of the 11 detailed Channel and Riparian
Cross-sections, evaluate the presence/absence and the proximity of 11 categories of human
influences with the procedure outlined in Table 6.2-12. Relate your observations and proximity
evaluations to the stream and riparian area within 5 m upstream and 5 m downstream from the
station (Figure 6.2-12). Four proximity classes are used: In the stream or on the bank within 5
m upstream or downstream of the cross-section transect, present within the 10 m x 10 m
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
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riparian plot but not in the stream or on the bank, present outside of the riparian plot, and
absent. Record data on the Channel/Riparian Cross-section Form as shown in Figure 6.2-7. If a
disturbance is within more than one proximity class, record the one that is closest to the stream
(e.g., C takes precedence over P).
A particular influence may be observed outside of more than one riparian observation
plot (e.g., at both transects D and £). Record it as present at every transect where you can see
it without having to sight through another transect or its 10 m x 10 m riparian plot.
Table 6.2-12. Procedure for estimating human influence
1. Standing mid-channel at a cross-section transect, look toward the left bank (left when facing
downstream), and estimate a 5 m distance upstream and downstream (10m total length). Also,
estimate a distance of 10 m back into the riparian zone to define a riparian plot area.
2. Examine the channel, bank and riparian plot area adjacent to the defined stream segment for the
following human influences: (1) walls, dikes, revetments, riprap, and dams; (2) buildings; (3)
pavement/cleared lots (e.g., paved, gravelled, dirt parking lot, foundation); (4) roads or railroads, (5)
inlet or outlet pipes; (6) landfills or trash (e.g., cans, bottles, trash heaps); (7) parks or maintained
lawns; (8) row crops; (9) pastures, range/and, hay fields, or evidence of livestock; (10) logging; and
(11) mining (including gravel mining).
3. For each type of influence, determine if it is present and what its proximity is to the stream and riparian
plot area. Consider human disturbance items as present if you can see them from the cross-section
transect. Do not include them if you have to sight through another transect or its 10 m x10 m riparian
plot.
4. For each type of influence, record the appropriate proximity class in the HUMAN INFLUENCE part of the
VISUAL RIPARIAN ESTIMATES section of the Channel/Riparian Cross-section Form. Proximity classes
are:
B (Bank) Present within the defined 10 m stream segment and located in the stream or on
the stream bank.
C (Close) Present within the 10 x 10 m riparian plot area, but away from the bank.
P (Present) Present, but outside the riparian plot area.
0 (Absent) Not present within or adjacent to thel 0 m stream segment or the riparian plot
area at the transect
5. Repeat Steps 1 through 4 for the right bank.
6. Repeat Steps 1 through 5 for each cross-section transect, (including any additional side channel
transects established when islands are present). Record data for each transect on a separate field
form.
6.2.5.8 Cross-section Transects on Side Channels
If the wetted channel is split by an island, and the estimated flow in the side channel is
less than or equal to 15% of the total flow, the bank and riparian measurements are made at
each side of the main channel (the minor side channel is ignored other than to note its presence
on the thalweg profile form), so one riparian plot is established on the island as shown in Figure
6.2-13. If an island is present that creates a major side channel containing more than 15% of
the total flow (Section 6.2.4.1), an additional cross-section transect is established for the side
channel as shown in Figure 6.2-13. Separate substrate, bank and riparian measurements are
made for side channel transects. Data from the additional side channel transect are recorded on
a separate Channel/Riparian Cross-section Form as shown in Figure 6.2-14. Riparian plots
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National Rivers and Streams Assessment
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Final Manual
Date: April 2009
Page 156
established on the island for each transect may overlap (and be < 10m shoreward) if the island
is less than 10m wide at the transect.
A) Island and minor side channel
No side channel cross-section transect,
Note presence on field form
Riparian plot established on Island
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Figure 6.2-13. Riparian and instream fish cover plots for a stream with minor and major side
channels.
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National Rivers and Streams Assessment
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Final Manual
Date: April 2009
Page 157
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page 158
6.2.5.9 Riparian "Legacy" Trees and Invasive Alien Species
Follow the procedures in Table 6.2-13 to locate the largest tree associated with each
transect. The tree you choose may not truly be an old legacy tree -just choose the largest you
see. We use these data to determine if there are true legacy trees somewhere within the
support reach. Note that only one tree is identified for each transect between that transect and
the next one upstream; at transect K, look upstream a distance of 4 channel widths. Record the
type of tree, and, if possible, the taxonomic group (using the list provided in Table 6.2-13) on the
left-hand column of the Riparian "Legacy" Trees and Invasive Alien Plants form (Figure 6.2-15).
Estimate the height of the tree and the diameter at breast height (dbh), and mark the
appropriate height and dbh classes on the form. Estimate and record the distance of the legacy
tree from the wetted margin of the stream.
Search in the 10 m x 10 m riparian and littoral plots on both banks for the presence of
any invasive alien species listed in the NRSA Invasive Species Guide provided to each field
crew. Document the species observed on the Riparian "Legacy" Trees and Invasive Alien Plants
form (Figure 5.2-8), answering the question of whether each of the target species is present in
the plot. If you have a camera, document the species with a photograph. If you observe no alien
taxa within the riparian and littoral plots, but can confidently identify them outside of the plots,
include your observations in the comments portion of the form. If the river is too wide to
effectively observe the far bank at a transect, record what you observe for the plot on the near
bank, record a "U" flag, and explain in the comments section of the form.
Table 6.2-13. Procedure for identifying riparian legacy trees
Legacy Trees:
• Beginning at Transect A, look upstream and downstream as far as you can see confidently.
Search both sides of the stream downstream to the next transect. Locate the largest tree
visible within 100m (or as far as you can see, if less) from the wetted bank.
• Classify this tree as broadleaf deciduous, coniferous, or broadleaf evergreen (classify
western larch as coniferous). Identify, if possible, the species or the taxonomic group of this
tree from the list below.
1. Acacia/Mesquite 10. Poplar/Cottonwood
2. Alder/Birch 11. Snag (Dead Tree of Any Species)
3. Ash 12. Spruce
4. Cedar/Cypress/Sequoia 13. Sycamore
5. Fir (including Douglas Fir, 14. Willow
Hemlock)
6. Juniper 15. Unknown, other Broadleaf Evergreen
7. Maple/Boxelder 16. Unknown or Other Conifer
8. Oak 17. Unknown or Other Deciduous
9. Pine
NOTE: If the largest tree is a dead "snag", enter "Snag" as the taxonomic group.
Estimate the height of the potential legacy tree, its diameter at breast height (dbh ) and its
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page 159
distance from the wetted margin of the stream. Enter this information on the left hand column of
the Riparian "Legacy" Trees and Invasive Alien Plants field form.
Alien Invasive Plants:
Examine the 10m x 10m riparian and littoral plots on both banks for the presence of alien
species. (Species lists will be provided)
Record the presence of any species listed within the plots on either the left or right bank on the
Riparian "Legacy" Trees and Invasive Alien Species field form. If none of the species listed is
present in the plots at a given transect, fill in the circle indicating "None" for this transect.
Repeat for each remaining transect (B through K). At transect "K", look upstream a distance of 4
channel widths when locating the legacy tree.
6.2.6 Channel Constraint, Debris Torrents, Recent Floods, and Discharge
6.2.6.1 Channel Constraint
After completing the thalweg profile and riparian/channel cross-section measurements
and observations, envision the stream at bankfull flow and evaluate the degree, extent and type
of channel constraint, using the procedures presented in Table 6.2-14. Record data on the
Channel Constraint Assessment Form (Figure 6.2-16). First, classify the stream reach channel
pattern as predominantly a single channel, an anastomosing channel, or a braided channel
(Figure 6.2-17):
1. Single channels may have occasional in-channel bars or islands with side channels,
but feature a predominant single channel, or a dominant main channel with a
subordinate side channel.
2. Anastomosing channels have relatively long major and minor channels (but no
predominant channel) in a complex network, diverging and converging around many
vegetated islands. Complex channel pattern remains even during major floods.
3. Braided channels also have multiple branching and rejoining channels, (but no
predominant channel) separated by unvegetated bars. Channels are generally
smaller, shorter, and more numerous, often with no obvious dominant channel.
During major floods, a single continuous channel may develop
After classifying the channel pattern, determine whether the channel is constrained
within a narrow valley, constrained by local features within a broad valley, unconstrained and
free to move about within a broad floodplain, or free to move about, but within a relatively
narrow valley floor. Then examine the channel to ascertain the bank and valley features that
constrain the stream. Entry choices for the type of constraining features are bedrock, hillslopes,
terraces/alluvial fans, and human land use (e.g., a road, a dike, landfill, rip-rap, etc.). Estimate
the percent of the channel margin in contact with constraining features (for unconstrained
channels, this is 0%). To aid in this estimate, you may wish to refer to the individual transect
assessments of incision and constraint. Finally, estimate the "typical" bankfull channel width and
estimate the average width of the valley floor either with a topographic map or visually. If you
cannot directly estimate the valley width (e.g., it is further than you can see, or if your view is
blocked by vegetation), record the distance you can see and mark the appropriate circle on the
field form.
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National Rivers and Streams Assessment
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Final Manual
Date: April 2009
Page 160
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page 161
Table 6.2-14. Procedures for assessing channel constraint
NOTE: These activities are conducted after completing the thalweg profile and littoral-riparian
measurements and observations, and represent an evaluation of the entire stream reach.
CHANNEL CONSTRAINT: Determine the degree, extent, and type of channel constraint based on
envisioning the stream at bankfull flow.
Classify the stream reach channel pattern as predominantly a single channel, an anasto-
mosing channel, or a braided channel.
Single channels may have occasional in-channel bars or islands with side channels,
but feature a predominant single channel, or a dominant main channel with a
subordinate side channel.
Anastomosing channels have relatively long major and minor channels branching and
rejoining in a complex network separated by vegetated islands, with no obvious
dominant channel.
Braided channels also have multiple branching and rejoining channels, separated by
unvegetated bars. Subchannels are generally small, short, and numerous, often with no
obvious dominant channel.
After classifying the channel pattern, determine whether the channel is constrained within a
narrow valley, constrained by local features within a broad valley, unconstrained and free to
move about within a broad floodplain, or free to move about, but within a relatively narrow valley
floor.
Then examine the channel to ascertain the bank and valley features that constrain the stream.
Entry choices for the type of constraining features are bedrock, hillslopes, terraces/alluvial fans,
and human land use (e.g., a road, a dike, landfill, rip-rap, etc.).
Based on your determinations from Steps 1 through 3, select and record one of the constraint
classes shown on the Channel Constraint Form.
Estimate the percent of the channel margin in contact with constraining features (for uncon-
strained channels, this is 0%). Record this value on the Channel Constraint Form.
Finally, estimate the "typical" bankfull channel width, and visually estimate the average width of
the valley floor. Record these values on the Channel Constraint Form.
NOTE: To aid in this estimate, you may wish to refer to the individual transect assessments of
incision and constraint that were recorded on the Channel/Riparian Cross-Section Forms.
NOTE: If the valley is wider than you can directly estimate, record the distance you can see
and mark the circle on the field form.
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National Rivers and Streams Assessment
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Final Manual
Date: April 2009
Page 162
CHANNEL CONSTRAINT FORM - WADEABLE/BOATABLE
SITE FWD8XXO00
DATE:
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Percent of Channel Margin Examples
Comments
03/OSKOM 2Mi Chan
Figure 6.2-16. Channel Constraint Form, showing data for channel constraint.
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National Rivers and Streams Assessment
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Date: April 2009
Page 163
A) Anastomosing channel pattern
2 Vegetated islands above bankfull flow. Multiple
channels remain during major flood events.
B) Braided channel pattern
Unvegetated bars below bankfull flow. Multiple
channel pattern disappears during major flood events.
Figure 6.2-17. Types of multiple channel patterns.
6.2.6.2 Debris Torrents and Recent Major Floods
Debris torrents, or lahars, differ from conventional floods in that they are flood waves of
higher magnitude and shorter duration, and their flow consists of a dense mixture of water and
debris. Their high flows of dense material exert tremendous scouring forces on streambeds. For
example, in the Pacific Northwest, flood waves from debris torrents can exceed 5 meters deep
in small streams normally 3 m wide and 15 cm deep. These torrents move boulders in excess of
1 m diameter and logs >1 m diameter and >10 m long. In temperate regions, debris torrents
occur primarily in steep drainages and are relatively infrequent, occurring typically less than
once in several centuries. They are usually set into motion by the sudden release of large
volumes of water upon the breaching of a natural or human-constructed impoundment, a
process often initiated by mass hillslope failures (landslides) during high intensity rainfall or
snowmelt. Debris torrents course downstream until the slope of the stream channel can no
longer keep their viscous sediment suspension in motion (typically <3% for small streams); at
this point, they "set up", depositing large amounts of sediment, boulders, logs, and whatever
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else they were transporting. Upstream, the torrent track is severely scoured, often reduced in
channel complexity and devoid of near-bank riparian vegetation. As with floods, the massive
disruption of the stream channel and its biota are transient, and these intense, infrequent events
will often lead to a high-quality complex habitat within years or decades, as long as natural
delivery of large wood and sediment from riparian and upland areas remains intact.
In arid areas with high runoff potential, debris torrents can occur in conjunction with flash
flooding from extremely high-intensity rainfall. They may be nearly annual events in some steep
ephemeral channels where drainage area is sufficient to guarantee isolated thunderstorms
somewhere within their boundaries, but small enough that the effect of such storms is not
dampened out by the portion of the watershed not receiving rainfall during a given storm.
Because they may alter habitat and biota substantially, infrequent major floods and
torrents can confuse the interpretation of measurements of stream biota and habitat in regional
surveys and monitoring programs. Therefore, it is important to determine if a debris torrent or
major flood has occurred within the recent past. After completing the thalweg profile and
channel/riparian measurements and observations, examine the stream channel along the entire
sample reach, including its substrate, banks, and riparian corridor, checking the presence of
features described on the Torrent Evidence Assessment Form (Figure 6.2-18). It may be
advantageous to look at the channel upstream and downstream of the actual sample reach to
look for areas of torrent scour and massive deposition to answer some of the questions on the
field form. For example, you may more clearly recognize the sample reach as a torrent
deposition area if you find extensive channel scouring upstream. Conversely, you may more
clearly recognize the sample reach as a torrent scour reach if you see massive deposits of
sediment, logs, and other debris downstream.
6.2.6.3 Stream Discharge
Stream discharge is equal to the product of the mean current velocity and vertical cross-
sectional area of flowing water. Discharge measurements are critical for assessing trends in
streamwater acidity and other characteristics that are very sensitive to streamflow differences.
Discharge should be measured at a suitable location within the sample reach that is as close as
possible to the location where chemical samples are collected, so that these data correspond.
Discharge is usually determined after collecting water chemistry samples.
No single method for measuring discharge is applicable to all types of stream channels.
The preferred procedure for obtaining discharge data is based on "velocity-area" methods (e.g.,
Rantz and others, 1982; Linsley et al., 1982). For streams that are too small or too shallow to
use the equipment required for the velocity-area procedure, two alternative procedures are
presented. One procedure is based on timing the filling of a volume of water in a calibrated
bucket. The second procedure is based on timing the movement of a neutrally buoyant object
(e.g., an orange or a small rubber ball) through a measured length of the channel, after
measuring one or more cross-sectional depth profiles within that length.
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TORRENT EVIDENCE ASSESSMENT FORM
r
SITE ID: FW08XXOO0
DATE: Q ^ / o f I 20
?,
•"TORRQNnr.EVIDENCE
Please fill in any of the following that are evident.
EVIDENCE OF TORRENT SCOURING:
01 • Stream channel has a recently devegetated corridor two or more times (he width of the tow flow channel. This
O corridor lacks riparian vegetation with possible exception of firewood even-aged alder or cottonwood seedlings,
j grasses, or other herbaceous plants..
O
02 - Stream substrate cobbles or large grave! particles are NOT IMBRICATED. (Imbricated means that they lie with flat
sides horizontal and that they are stacked like roof shingles - imagine the upstream direction as the top of the "roof.") In
a torrent scour or deposition channel, the stones are laying in unorganized patterns, lying "every which way." In addition
many of trie substrate partlctes are angular (not "water-worn.")
O
O
03 • Channel has little evidence of pool-riffle structure. (Fot example, could yoy ride a mountain bike down the channel?)
04 - The stream channel Is scoured down to bedrock for substantial portion of reach.
O
05 - There are gravel or cobble berrre {little tevoes) above taankfull level.
O
O
06 - Downstream of the scoured roach (possibly several mites), there are massive deposits of sediment, logs and other
debris.
07 - Riparian trees havo fresh bark teats at many points along the stream at seemingly unbelievable heights above the
channel bed.
O
08 - Riparian trws have fallen into the channel as a resuR of scouring near their roots
EVIDENCE OF TORRENT DEPOSITS:
O
O
09 - There are massivo daposits of sediment, logs, and other debris in the reach They may contain wood and boulders
that, in your judgement, could not have been moved by the stream at even extreme flood st^ge.
10 - If ttw stream has begun to erode newly laid deposits, it is evident that these deposits am "MATRIX SUPPORTED."
This means that the large particles, like boulders and cobbles, are often not touching each other, but havo silt, sand, and
other fine partlctes between them (their weight is supported by these fine particles - in contrast to a normal stream
deposit, where fines, if present, normally "fill-In" the interstices between coarser particles.)
NO EVIDENCE:
11 - f-k> evidence of torrent scouring or torrent deposits.
COMMENTS
NRSA Twrtiml Eviderjce
Figure 6.2-18. Torrent Evidence Assessment Form.
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6.2.6.4 Velocity-Area Procedure
Because velocity and depth typically vary greatly across a stream, accuracy in field
measurements is achieved by measuring the mean velocity and flow cross-sectional area of
many increments across a channel (Figure 6.2-19). Each increment gives a subtotal of the
stream discharge, and the whole is calculated as the sum of these parts. Discharge
measurements are made at only one carefully chosen channel cross-section within the
sampling reach. It is important to choose a channel cross-section that is as much like a canal
as possible. A glide area with a "U" shaped channel cross-section that is free of obstructions
provides the best conditions for measuring discharge by the velocity-area method. You may
remove rocks and other obstructions to improve the cross-section before any measurements
are made. However, because removing obstacles from one part of a cross-section affects
adjacent water velocities, you must not change the cross-section once you commence collecting
the set of velocity and depth measurements.
The procedure for obtaining depth and velocity measurements is outlined in Table 6.2-
15. Record the data from each measurement on the Stream Discharge Form as shown in Figure
6.2-20. In the field, data will be recorded using only one of the available procedures.
15 to 20 equally spaced
intervals across stream.
beginning at left margin
Measure stream depth at the midpoint
of each interval, and obtain velocity
measurements at 0.6 depth
Extended surveyor's
rod or tape measure
\
Record distance
and depth of
right margin
Figure 6.2-19. Layout of channel cross-section for obtaining discharge data by the velocity-area
procedure.
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Table 6.2-15. Velocity-Area procedure for determining stream discharge
1. Locate a cross-section of the stream channel for discharge determination that has most of the
following qualities (based on Rantz and others, 1982):
• Segment of stream above and below cross-section is straight
• Depths mostly greater than 15 centimeters, and velocities mostly greater than 0.15
meters/second. Do not measure discharge in a pool.
• "U" shaped, with a uniform streambed free of large boulders, woody debris or brush, and dense
aquatic vegetation.
• Flow is relatively uniform, with no eddies, backwaters, or excessive turbulence.
2. Lay the surveyor's rod (or stretch a measuring tape) across the stream perpendicular to its flow, with
the "zero" end of the rod or tape on the left bank, as viewed when looking downstream. Leave the tape
tightly suspended across the stream, approximately one foot above water level.
3. Attach the velocity meter probe to the calibrated wading rod. Check to ensure the meter is functioning
properly and the correct calibration value is displayed. Calibrate (or check the calibration) the velocity
meter and probe as directed in the meter's operating manual. Fill in the "VELOCITY AREA" circle on
the Stream Discharge Form.
4. Divide the total wetted stream width into 15 to 20 equal-sized intervals. To determine interval width,
divide the width by 20 and round up to a convenient number. Intervals should not be less than 10 cm
wide, even if this results in less than 15 intervals. The first interval is located at the left margin of the
stream (left when looking downstream), and the last interval is located at the right margin of the stream
(right when looking downstream).
5. Stand downstream of the rod or tape and to the side of the first interval point (closest to the left bank if
looking downstream).
6. Place the wading rod in the stream at the interval point and adjust the probe or propeller so that it is at
the water surface. Fill in the appropriate "Distance Units" and "Depth Units" circles on the Stream
Discharge Form. Record the distance from the left bank and the depth indicated on the wading rod on
the Stream Discharge Form.
Note for the first interval, distance equals 0 cm, and in many cases depth may also equal 0 cm. For
the last interval, distance will equal the wetted width (in cm) and depth may again equal 0 cm.
7. Stand downstream of the probe or propeller to avoid disrupting the stream flow. Adjust the position of
the probe on the wading rod so it is at 0.6 of the measured depth below the surface of the water. Face
the probe upstream at a right angle to the cross-section, even if local flow eddies hit at oblique angles
to the cross-section.
8. Wait 20 seconds to allow the meter to equilibrate, then measure the velocity. Fill in the appropriate
"Velocity Units" circle on the Stream Discharge Form. Record the value on the Stream Discharge
Form. Note for the first interval, velocity may equal 0 because depth will equal 0.
• For the electromagnetic current meter (e.g., Marsh-McBirney), use the lowest time constant
scale setting on the meter that provides stable readings.
• For the impeller-type meter (e.g., Swoffer2100), set the control knob at the mid-position of
"DISPLAY AVERAGING". Press "RESET" then "START" and proceed with the measurements.
9. Move to the next interval point and repeat Steps 6 through 8. Continue until depth and velocity
measurements have been recorded for all intervals. Note for the last interval (right margin), depth and
velocity values may equal 0.
10. At the last interval (right margin), record a "Z" flag on the field form to denote the last interval sampled.
11. If using a meter that computes discharge directly, check the "Q" circle on the discharge form, and
record calculated discharge value. In this case, you do not have to record the depth and velocity data
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for each interval.
DISCHARGE FORM - WADEABLE
SITE ID: FW08 XXOOC?
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Cwnments
- _P^TA jF^*. itt-t- F«c/A MtTftePS A»e~ SH»t^fJ, \
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Ffag Codes: K ^ No m»a««t^m©-fi!t or e^servat^on made; U » Susp^cl jfi«f.asurement osr Qte«rviHioti; Q * Unace«ptgEbl© CC
ciwck aasecsated wMh rrKasuremen!, 2 « Lssi ^tatSesri ttwasured ftf not Suwtiwi 20^ Fl, F2, etc. = Miscellaneous Wags Ofafi
as§ign*d by each field crew. Explain all flags in eamnwntiS section 1^^ !
03^18/2DOi NRSA Stream Discharge L V J
Figure 6.2-20. Discharge Form, showing data recorded for all discharge measurement
procedures.
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6.2.6.5 Timed Filling Procedure
In channels too "small" for the velocity-area method, discharge can sometimes be
measured by filling a container of known volume and timing the duration to fill the container.
"Small" is defined as a channel so shallow that the current velocity probe cannot be
placed in the water, or where the channel is broken up and irregular due to rocks and debris,
and a suitable cross-section for using the velocity area procedure is not available. This can be
an extremely precise and accurate method, but requires a natural or constructed spillway of
freefalling water. If obtaining data by this procedure will result in a lot of channel disturbance or
stir up a lot of sediment, wait until after all biological and chemical measurements and sampling
activities have been completed.
Choose a cross-section of the stream that contains one or more natural spillways or
plunges that collectively include the entire stream flow. A temporary spillway can also be
constructed using a portable V-notch weir, plastic sheeting, or other materials that are available
onsite. Choose a location within the sampling reach that is narrow and easy to block when using
a portable weir. Position the weir in the channel so that the entire flow of the stream is
completely rerouted through its notch (Figure 6-3). Impound the flow with the weir, making sure
that water is not flowing beneath or around the side of the weir. Use mud or stones and plastic
sheeting to get a good waterproof seal. The notch must be high enough to create a small
spillway as water flows over its sharp crest.
The timed filling procedure is presented in Table 6.2-16. Make sure that the entire flow of
the spillway is going into the bucket. Record the time it takes to fill a measured volume on the
Discharge Measurement Form as shown in Figure 6-2. Repeat the procedure 5 times. If the
cross-section contains multiple spillways, you will need to do separate determinations for each
spillway. If so, clearly indicate which time and volume data replicates should be averaged
together for each spillway; use additional Stream Discharge Form if necessary.
Table 6.2-16. Timed filling procedure for determining stream discharge
NOTE: If measuring discharge by this procedure will result in significant channel disturbance or will
stir up sediment, delay determining discharge until all biological and chemical measurement and
sampling activities have been completed.
1. Choose a cross-section that contains one or more natural spillways or plunges, or construct a
temporary one using on-site materials, or install a portable weir using a plastic sheet and on-site
materials.
2. Fill in the "TIMED FILLING" circle in the stream discharge section of the Stream Discharge Form.
3. Position a calibrated bucket or other container beneath the spillway to capture the entire flow. Use
a stopwatch to determine the time required to collect a known volume of water. Record the
volume collected (in liters) and the time required (in seconds) on the Stream Discharge Form.
4. Repeat Step 3 a total of 5 times for each spillway that occurs in the cross-section. If there is more
than one spillway in a cross-section, you must use the timed-filling approach on all of them.
Additional spillways may require additional data forms
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6.2.6.6 Neutrally-Buoyant Object Procedure
In very small, shallow streams with no waterfalls, where the standard velocity-area or
timed-filling methods cannot be applied, the neutrally buoyant object method may be the only
way to obtain an estimate of discharge. The required pieces of information are the mean flow
velocity in the channel and the cross-sectional area of the flow. The mean velocity is estimated
by measuring the time it takes for a neutrally buoyant object to flow through a measured length
of the channel. The channel cross-sectional area is determined from a series of depth
measurements along one or more channel cross-sections. Since the discharge is the product of
mean velocity and channel cross-sectional area, this method is conceptually very similar to the
standard velocity-area method.
The neutrally buoyant object procedure is described in Table 6.2-17. Examples of
suitable objects include plastic golf balls (with holes), small sponge rubber balls, or small sticks.
The object must float, but very low in the water. It should also be small enough that it does not
"run aground" or drag bottom. Choose a stream segment that is roughly uniform in cross-
section, and that is long enough to require 10 to 30 seconds for an object to float through it.
Select one to three cross-sections to represent the channel dimensions within the segment,
depending on the variability of width and/or depth. Determine the stream depth at 5 equally
spaced points at each cross-section. Three separate times, measure the time required for the
object to pass through the segment that includes all of the selected cross-sections. Record data
on the Stream Discharge Form as shown in Figure 6.2-20.
Table 6.2-17. Neutrally buoyant object procedure for determining stream discharge
1. Fill in the "NEUTRALLY BUOYANT OBJECT" circle on the Stream Discharge Form.
2. Select a segment of the sampling reach that is deep enough to float the object freely, and long
enough that it will take between 10 and 30 seconds for the object to travel. Mark the units used
and record the length of the segment in the "FLOAT DIST." field of the Stream Discharge Form.
3. If the channel width and/or depth change substantially within the segment, measure widths and
depths at three cross-sections, one near the upstream end of the segment, a second near the
middle of the segment, and a third near the downstream end of the segment.
If there is little change in channel width and/or depth, obtain depths from a single "typical" cross-
section within the segment.
4. At each cross-section, measure the wetted width using a surveyor's rod or tape measure, and
record both the units and the measured width on the Stream Discharge Form. Measure the stream
depth using a wading rod or meter stick at points approximately equal to the following proportions
of the total width: 0.1, 0.3, 0.5, 0.7, and 0.9. Record the units and the depth values (not the
distances) on the Stream Discharge Form.
5. Repeat Step 4 for the remaining cross-sections.
6. Use a stopwatch to determine the time required for the object to travel through the segment.
Record the time in the "FLOAT TIME" field of the Stream Discharge Form.
7. Repeat Step 6 two more times. The float time may differ somewhat for the three trials.
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Water Level
Bucket
Figure 6.2-21. Use of a portable weir in conjunction with a calibrated bucket to obtain an estimate
of stream discharge.
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6.2.7 Equipment and Supplies
Table 6.2-18 lists the equipment and supplies required to conduct all the activities
described for characterizing physical habitat. This checklist is similar to the checklist presented
in Appendix A, which is used at the base location (Section 3) to ensure that all of the required
equipment is brought to the stream.
Table 6.2-18. Checklist of equipment and supplies for physical habitat
For taking
measurements
Surveyor's telescoping leveling rod (round profile, metric scale, 7.5 m extended)
50 m or 100 m measuring tape & reel
Laser rangefinder (400 ft. distance range) and clear waterproof bag
Digital camera with extra memory card & battery
Two 1/2-inch diameter PVC pipe, 2-3 m long: Two of these, each marked at the same
height (for use in slope determinations involving two persons)
Meter stick, or a short rod or pole (e.g., a ski pole) with cm markings for thalweg
measurements, or the PVC pipe for slope determinations can be marked in cm
1 roll each colored surveyor's flagging tape (2 colors)
Convex spherical canopy densiometer (Lemmon Model A), modified with taped "V"
Clinometer
Bearing compass (Backpacking type)
Binoculars
1 or 2 fisherman's vest with lots of pockets and snap fittings. Used to hold the various
measurement equipment (densiometer, clinometer, compass, etc.).
2 pair chest waders (hip waders can be used in shallower streams).
Current velocity meter, probe, and operating manual
Top-set wading rod for use with current velocity meter
Portable Weir with 60° "V" notch (optional) and plastic sheeting to use with weir
Plastic bucket (or similar container) with volume graduations
Stopwatch
Neutrally buoyant object (e.g., plastic golf ball with holes, small rubber ball, stick)
Field Methods Manual and/or laminated quick reference guide
For recording
data
Covered clipboards (lightweight, with strap or lanyard)
Soft (#2) lead pencils (mechanical are acceptable)
11 plus extras Channel/Riparian Cross-section Forms
11 plus extras Thalweg Profile and Woody Debris Forms
1+ extras field Form: Stream Verification Form
1+ extras field Form: Field Measurement Form
1+ extras field Form: Discharge Form
1+ extras field Form: Sample Collection Form
1+ extras field Form: Riparian "Legacy" Trees and Invasive Alien Plants
1+ extras field Form: Channel Constraint
1+ extras field Form: Torrent Evidence Form
1+ extras field Form: Fish Gear and Voucher/Tissue Information Form
1+ extras field Form: Fish Collection Form
1+ extras field Form: Slope and Bearing Form
1+ extras field Form: Visual Assessment Form
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6.3 Periphyton
6.3.1 Summary of Method
Collect periphyton from the 11 cross-section transects ("A" through "K") established
within the sampling reach. Collect periphyton samples at the same time as sediment enzyme
samples (Section 6.1.3) and benthic macroinvertebrate samples (Sections 6.4.1). Prepare one
composite "index" sample of periphyton for each site. At the completion of the day's sampling
activities, but before leaving the site, prepare four types of laboratory samples (an
ID/enumeration sample to determine taxonomic composition and relative abundances, a
chlorophyll sample, a biomass sample (for ash-free dry mass [AFDM]), and a acid/alkaline
phosphatase activity [APA] sample) from the composite periphyton sample.
6.3.2 Equipment and Supplies
Table 6.3-1 is a checklist of equipment and supplies required to conduct periphyton
sample collection and processing activities. This checklist is similar to the checklist presented in
Appendix A, which is used at the base location (Section 3) to ensure that all of the required
equipment is brought to the river.
Table 6.3-1. Equipment and supplies list for periphyton at wadeable sites
For collecting samples
Large Funnel (15-20 cm diameter)
12-cm2 area delimiter (3.8 cm diameter pipe, 3 cm tall)
Stiff-bristle toothbrush with handle bent at 90° angle
1-L wash bottle for stream water
500-mL plastic bottle for the composite sample
60-mL plastic syringe with 3/8" hole bored into the end
Field Operations Manual or laminated Quick Reference Guide
For recording measurements
Sample Collection Form
Soft (#2) lead pencils for recording data on field forms
Fine-tipped indelible markers for filling out sample labels
Sample labels (4 per set) with the same Sample ID Number
Clear tape strips for covering labels
6.3.3 Sampling Procedure
At each of the 11 transects, collect samples from the sampling station assigned during
the layout of the reach. Collect the substrate selected for sampling from a depth no deeper than
0.5 m. If a sample cannot be collected because the location is too deep, skip the transect. The
procedure for collecting samples and preparing a composite sample is presented in Table 6.3-2.
Collect one sample from each of the transects and composite in one bottle to produce one
composite sample for each site. Record the volume of the sample on the Sample Collection
Form as shown in Figure 6.1-3.
Table 6.3-2. Procedure for collecting composite index samples of periphyton at wadeable sites
1. Starting with Transect "A", collect a single sample from the assigned sampling station using the
procedure below.
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a) Collect a sample of substrate (rock or wood) that is small enough (< 15 cm diameter) and can be
easily removed from the river. Place the substrate in a plastic funnel which drains into a 500-mL
plastic bottle with volume graduations marked on it.
b) Use the area delimiter to define a 12-cm2 area on the upper surface of the substrate. Dislodge
attached periphyton from the substrate within the delimiter into the funnel by brushing with a stiff-
bristled toothbrush for 30 seconds. Take care to ensure that the upper surface of the substrate is
the surface that is being scrubbed, and that the entire surface within the delimiter is scrubbed.
c) Fill a wash bottle with river water. Using a minimal volume of water from this bottle, wash the
dislodged periphyton from the funnel into the 500-mL bottle. If no coarse sediment (cobbles or
larger) are present:
• Use the area delimiter to confine a 12-cm2 area of soft sediments.
• Vacuum the top 1 cm of sediments from within the delimited area into a de-tipped 60-mL
syringe.
• Empty the syringe into the same 500-mL plastic bottle as above.
d) Put the bottle in a cooler on ice while you travel between transects and collect the
subsequent samples. (The samples need to be kept cool and dark because a chlorophyll
sample will be filtered from the composite.)
2. Repeat Step 1 for transects "B" through "K". Place the sample collected at each sampling site into the
single 500-mL bottle to produce the composite index sample.
3. If all 11 samples are not collected, record the number of transects collected and reason for any missed
collection on the field forms.
4. After samples have been collected from all 11 transects, thoroughly mix the 500-mL bottle regardless
of substrate type. Record the total estimated volume of the composite sample in the periphyton section
of the Sample Collection Form.
6.3.4 Sample Processing in the Field
You will prepare four different types of laboratory samples from the composite index
samples: an ID/enumeration sample (to determine taxonomic composition and relative
abundances), a chlorophyll sample, a biomass sample (for ash-free dry mass [AFDM]), and
an acid/alkaline phosphatase activity (APA) sample. All the sample containers required for an
individual site should be sealed in plastic bags until use to avoid external sources of
contamination (e.g., dust, dirt, or mud) that are present at site shorelines. Please refer to
Sections 7.2.5 and 7.2.6 processing the periphyton samples.
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6.4 Benthic Macroinvertebrates
6.4.1 Summary of Method
Collect benthic macroinvertebrate composite samples using a D-frame net with 500 urn
mesh openings. Take the samples from the sampling stations at the 11 transects equally
distributed along the targeted reach. You will proportionally sample multiple habitats at sampling
stations randomly assigned on each transect. Multiple habitats will include bottom substrate as
well as woody debris, macrophytes, and leaf packs. Composite all sample material and field-
preserve with -95% ethanol.
High gradient streams
• Primary samples are taken at each transect at either 25%, 50%, or 75% transect
distance (according to the initial randomized pattern). Primary samples will be
collected from a 1 square foot quadrat.
Low gradient streams
• Primary samples are taken at each transect at either 25%, 50%, or 75% transect
distance (according to the initial randomized pattern). Primary samples will be
collected from a 1 square foot quadrat.
• additional, separate samples taken at either 0%, 50%, or 100% transect distance to
include edge samples (snags, undercut banks, root wads, macrophyte beds, etc.).
Low gradient samples will be collected from a 1 linear meter sweep.
6.4.2 Equipment and Supplies
Table 6.4-1 shows the checklist of equipment and supplies required to complete the
collection of benthic macroinvertebrates. This checklist is similar to the checklist presented in
Appendix A, which is used at the base location to ensure that all of the required equipment is
brought to the site. Record collection data on the Sample Collection Form (Fig. 6.1-2).
Table 6.4-1. Equipment and supplies list for benthic macroinvertebrate collection at wadeable
sites
For collecting
samples
Modified kick net (D-frame with 500
|jm mesh) and 4-ft handle
Watch with timer or stopwatch
Buckets, plastic, 8- to 10-qt
Sieve bucket with 500 urn mesh
openings (U.S. std No. 35)
Watchmakers' forceps
Wash bottle, 1-L capacity labeled
"STREAM WATER"
Funnel, with large bore spout
Small spatula, spoon, or scoop to
transfer sample
Sample jars, 1-L HOPE plastic
suitable for use with ethanol
95% ethanol, in a proper container
Cooler (with absorbent material) for
transporting ethanol & samples
Plastic electrical tape
Scissors
Field Operations Manual or
laminated Quick Reference Guide
For recording
measurements
Composite benthic sample labels
with & without preprinted ID
numbers
Blank labels on waterproof paper for
inside of jars
Soft (#2) lead pencils
Fine-tip indelible markers
Clear tape strips
Sample Collection Form
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6.4.3 Sampling Procedure
Figure 6.4-1 summarizes how samples will be collected from wadeable sites. The
transect sample design for collecting benthic macroinvertebrates is shown in Figure 6.4-2. This
design was used in the EPA's Wadeable Streams Assessment, which provides continuity for a
nationwide assessment. Collect a sample from 1-m downstream of each of the 11 cross-
section transects at the assigned sampling station. The process for selecting the sample
stations is described in the Initial Site Procedures Section (Section 4). At transects assigned a
"Center" sampling point where the stream width is between one and two net widths wide, pick
either the "Left" or "Right" sampling point instead. If the stream is only one net wide at a
transect, place the net across the entire stream width and consider the sampling point to be
"Center". If a sampling point is located in water that is too deep or unsafe to wade, select an
alternate sampling point on the transect at random.
The procedure for collecting a sample at each transect is described in Table 6.4-2. At
each sampling point, determine if the habitat is a "riffle/run" or a "pool/glide" (any area where
there is not sufficient current to extend the net is operationally defined as a pool/glide habitat).
Record the dominant substrate type (fine/sand, gravel, coarse substrate (coarse gravel or
larger) or other (e.g., bedrock, hardpan, wood, aquatic vegetation, etc.) and the habitat type
(pool, glide, riffle, or rapid) for each sample collected on the Sample Collection Form as shown
in Figure 6.1-2. As you proceed upstream from transect to transect, combine all samples into a
bucket. An additional separate sample will be taken at low gradient streams to include
edge habitat (leaf litter, organic deposits, undercut banks, root wads, macrophyte beds, etc.)
6.4.4 Sample Processing in Field
Use a 500 |o,m mesh sieve bucket placed inside a larger bucket full of site water while
sampling to carry the composite sample as you travel around the site. It is recommended that
teams carry a sample bottle containing a small amount of ethanol with them to enable them to
immediately preserve larger predaceous invertebrates such as helgramites and water beetles.
Doing so will help reduce the chance that other specimens will be consumed or damaged prior
to the end of the field day. Once the composite sample from all stations is sieved and reduced in
volume, store in a 1-liter jar and preserve with 95% ethanol. Do not fill jars more than 1/3 full of
material. Multiple jars may be required if detritus is heavy (Table 6.4-3). Try to use no more than
5 jars per site. If more than one jar is used for a composite sample, use the "extra jar" label
provided; record the SAME sample ID number on this "extra jar" label. DO NOT use two
different sample numbers on two jars containing one single sample. Cover the labels with
clear tape. The sample ID number is also recorded with a No. 2 lead pencil on a waterproof
label that is placed inside each jar. Be sure the inside label and outside label describe the same
sample.
NON-WADEABLE
LOW GRADIENT
STREAMS ONLY
At Transect "A", randomly locate the first
sampling station (left, center or right facing
downstream)
r>nd
Collect 2 separate sample from 0%, 50%
or 100% of the stream width.
(Collect at the station immediately to the
right of the primary station)
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Collect sample using riffle/run or pool/glide
procedure from 1 ft2 quadrat at 25%, 50% or
75% width of the channel
Transfer sample into sieve bucket.
Thoroughly rinse net into the sieve bucket.
Immediately preserve large predaceous
invertebrates in ethanol.
Mark the substrate and channel habitat type
on the sample collection form
Proceed to sampling station on Transect "B"
and collect next sample; continue collecting
samples through Transect "K".
The samples from all stations are
composited to create a single sample for the
site.
Collect sample from a 1 linear meter sweep
Transfer sample into sieve bucket.
Thoroughly rinse net into the sieve bucket.
Immediately preserve large predaceous
invertebrates in ethanol.
Mark the substrate and channel habitat type
on the sample collection form
For edge samples, mark the dominant edge
type present
Proceed to sampling station on Transect "B"
and collect next sample; continue collecting
samples through Transect "K".
The samples from all stations are
composited to create a single sample for the
site. Be sure to keep the primary and low
gradient samples separate.
Figure 6.4-1. Benthic macroinvertebrate collection at wadeable sites.
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FLOW
K
\
R
i — _j
C
\
i__
_j
/
R
Combine ALL kick net samples collected from ALL transects
TRANSECT SAMPLES (1 per transect)
Sampling point of each transect selected systematically after random start
(separate samples for wadeable low-gradient streams at 0%, 50% or 100%)
Modified D-frame kick-net
1 ft2 quadrat sampled for 30 seconds
(1 linear meter sweep for additional low-gradient sample)
Composite Reachwide Sampling
V
7
Sieving
• 500 urn mesh
• Remove as much debris &
fine sediment as possible
Composite Index Sample
• 1-Ljars
• Fill < 1/3 with sample
• Preserve with 95% ethanol to
final concentration of ~ 70%
• Try to use < 5 1-L jars
Figure 6.4-2. Transect sample design for collecting benthic macroinvertebrates at wadeable
sites.
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Table 6.4-2. Procedure for benthic macroinvertebrate sampling at wadeable sites
1. At 1 m downstream of each transect, beginning with Transect "A", randomly locate the first sampling
station (Left, Center, or Right as you face downstream) as 25%, 50%, and 75% of the wetted width,
respectively. If you cannot collect a sample at the designated point because of deep water or unsafe
conditions, relocate to another random point on the same transect.
2. Determine if there is sufficient current in the area at the sampling station to fully extend the net. If so,
classify the habitat as "riffle/run" and proceed to Step 3. If not, use the sampling procedure described
for "pool/glide" habitats starting at Step 9.
NOTE: If the net cannot be used, hand pick a sample for 30 seconds from about 1 ff of substrate
at the sampling point. For vegetation-choked sampling points, sweep the net through the
vegetation within a 1 ft2 quadrat for 30 seconds. Place this hand-picked sample directly into the
sample container. Assign a "U" flag (non-standard sample) to the sample and indicate which
transect(s) required the modified collection procedure in the comments section. Go to Step 13.
Riffle/Run Habitats:
3. With the net opening facing upstream, quickly position the net securely on the stream bottom to
eliminate gaps under the frame. Avoid large rocks that prevent the net from seating properly on the
stream bottom.
NOTE: If there is too little water to collect the sample with the D-net, randomly pick up 10 rocks
from the riffle and pick and wash the organisms off them into a bucket which is half full of water.
4. Holding the net in position on the substrate, visually define a quadrat that is one net width wide and
long upstream of the net opening. The area within this quadrat is 1 ft2
5. Check the quadrat for heavy organisms, such as mussels and snails. Remove these organisms by
hand and place them into the net. Pick up loose rocks or other larger substrate particles in the
quadrat. Use your hands or a scrub brush to dislodge organisms and wash them into the net. Scrub
all rocks that are golf ball-sized or larger and which are halfway into the quadrat. After scrubbing,
place the substrate particles outside of the quadrat.
6. Hold the D-net securely in position. Starting at the upstream end of the quadrat, vigorously kick the
remaining finer substrate within the quadrat for 30 seconds (use a stopwatch).
NOTE: For samples located within dense beds of long, filamentous aquatic vegetation (e.g.,
algae or moss), kicking within the quadrat may not be sufficient to dislodge organisms in the
vegetation. Usually these types of vegetation are lying flat against the substrate due to current.
Use a knife or scissors to remove only the vegetation that lies within the quadrat (i.e., not
entire strands that are rooted within the quadrat) and place it into the net.
7. Pull the net up out of the water. Immerse the net in the stream several times to remove fine
sediments and to concentrate organisms at the end of the net. Avoid having any water or material
enter the mouth of the net during this operation.
8. Go to Step 13.
Pool/Glide Habitats:
9. Visually define a quadrat that is one net width wide and long at the sampling point. The area within
this quadrat is 1 ft .
10. Check the quadrat for heavy organisms, such as mussels and snails. Remove these organisms by
hand and place them into the net. Pick up loose rocks or other larger substrate particles in the
quadrat. Use your hands or a scrub brush to dislodge organisms and wash them into the net. Scrub
all rocks that are golf ball-sized or larger and which are halfway into the quadrat. After scrubbing,
place the substrate particles outside of the quadrat.
11. Vigorously kick the remaining finer substrate within the quadrat with your feet while dragging the net
repeatedly through the disturbed area just above the bottom. Keep moving the net all the time so that
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the organisms trapped in the net will not escape. Continue kicking the substrate and moving the net
for 30 seconds.
NOTE: If there is too little water to use the kick net, stir up the substrate with your gloved hands
and use a sieve with 500 yin mesh size to collect the organisms from the water in the same way
the net is used in larger pools.
12. After 30 seconds, remove the net from the water with a quick upstream motion to wash the organisms
to the bottom of the net.
All samples:
13. Invert the net into a sieve bucket and transfer the sample. Remove as much gravel as possible so
that the organisms do not get damaged. Inspect the net for any residual organisms clinging to the net
and deposit them into the bucket. Use forceps if necessary to remove organisms from the net.
Carefully inspect any large objects (such as rocks, sticks, and leaves) in the bucket and wash any
organisms found off of the objects and into the bucket before discarding the object. Remove as much
detritus as possible without losing organisms.
14. Determine the predominant substrate size/type you within the sampling quadrat. Fill in the
appropriate circle for the dominant substrate type for the transect on the Sample Collection Form.
NOTE: If there are co-dominant substrate types, you may fill in more than one circle; note the co-
dominants in the comments section of the form.
• Fine/sand: not gritty (silt/clay/muck <0.06 mm diam.) to gritty, up to ladybug sized (2 mm)
• Gravel: fine to coarse gravel (ladybug to tennis ball sized; 2 mm to 64 mm)
• Coarse: Cobble to boulder (tennis ball to car sized; 64 mm to 4000 mm)
• Other: bedrock (larger than car sized; > 4000 mm), hardpan (firm, consolidated fine
substrate), wood of any size, aquatic vegetation, etc.). Note type of "other" substrate in
comments on field form.
15. Identify the habitat type where the sampling quadrat was located. Fill in the appropriate circle for
channel habitat type for the transect on the Sample collection Form.
• Pool; Still water; low velocity; smooth, glassy surface; usually deep compared to other parts
of the channel
• GLide: Water moving slowly, with smooth, unbroken surface; low turbulence
• Riffle: Water moving, with small ripples, waves, and eddies; waves not breaking, and
surface tension is not broken; "babbling" or "gurgling" sound.
• RApid: Water movement is rapid and turbulent; surface with intermittent "white water" with
breaking waves; continuous rushing sound.
16. Thoroughly rinse the net before proceeding to the next sampling station. Proceed upstream to the
next transect (through Transect K, the upstream end of the reach) and repeat steps 1-16. Combine
all kick net samples from riffle/run and pool/glide habitats into the bucket.
Additional Sample for low gradient streams:
17. At low gradient stream sites, an additional separate composite sample will be taken. The sample will
be collected with the same methods above, with the following modifications:
18. Collect the samples at 0, 50, or 100% transect distance to include edge samples (collected from leaf
litter, snags, organic deposits, undercut banks, root wads, macrophyte beds, etc.).
19. If the primary sample was collected at the Left at Transect A, collect the additional sample at the
Center of Transect A, then continue with Right at Transect B, Left at Transect C, until you collect at
every transect rotating through Left, Center, and Right.
20. Collect the samples over 1 linear meter. Vigorously disturb the bank or bottom habitat and quickly
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sweep the net to collect the loosened material.
21. Composite and label this sample separately from the first sample collected. This will be identified in
the lab as two separate samples.
22. Write in the appropriate abbreviation for substrate & channel habitat type on the Sample Collection
Form. For samples taken at the left or right edge of the transect, write in the appropriate abbreviation
for the dominant edge type present.
Record information for each composite sample on the Sample Collection Form as shown
in Figure 6.1-2(a). If a sample requires more than one jar, make sure the correct number of jars
for the sample is recorded on the Sample Collection Form. Do not fill out the collection form
until you have collected (or confirmed at the site that you will collect) samples. If forms
are filled out before you arrive at the site, and then no samples are collected, a lot of time is
wasted by others later trying to find samples that do not exist. Place the samples in a cooler or
other secure container for transporting and/or shipping to the laboratory (see Appendix C).
Table 6.4-3. Procedure for preparing composite samples for benthic macroinvertebrates at
wadeable sites
1. Pour the entire contents of the bucket into a sieve bucket with 500 urn mesh size. Remove any large
objects and wash off any clinging organisms back into the sieve before discarding. Remove any
inorganic material, such as cobble or rocks.
2. Using a wash bottle filled with river water, rinse all the organisms from the bucket into the sieve. This
is the composite sample for the reach.
3. Estimate the total volume of the sample in the sieve and determine how large a jar will be needed for
the sample (500-mL or 1-L) and how many jars will be required. Try to use no more than 5 jars per
site.
4. Fill in a sample label with the Sample ID and date of collection. Attach the completed label to the jar
and cover it with a strip of clear tape. Record the sample ID number for the composite sample on the
Sample Collection Form. For each composite sample, make sure the number on the form matches the
number on the label.
5. Wash the contents of the sieve to one side by gently agitating the sieve in the water. Wash the sample
into a jar using as little water from the wash bottle as possible. Use a large-bore funnel if necessary. If
the jar is too full pour off some water through the sieve until the jar is not more than 1/3 full, or use a
second jar if a larger one is not available. Carefully examine the sieve for any remaining organisms
and use watchmakers' forceps to place them into the sample jar.
• If a second jar is needed, fill in a sample label that does not have a pre-printed ID number on it.
Record the ID number from the pre-printed label prepared in Step 4 in the "SAMPLE ID" field of
the label. Attach the label to the second jar and cover it with a strip of clear tape. Record the
number of jars required for the sample on the Sample Collection Form. Make sure the number
you record matches the actual number of jars used. Write "Jar N ofX" on each sample label
using a waterproof marker ("N" is the individual jar number, and "X" is the total number of jars for
the sample).
6. Place a waterproof label inside each jar with the following information written with a number 2 lead
pencil:
Site ID • Collectors initials
Type of sampler and mesh size used • Number of stations sampled
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Table 6.4-3. Procedure for preparing composite samples for benthic macroinvertebrates at
wadeable sites
Name of site
Date of collection • Jar "N" of "X"
7. Completely fill the jar with 95% ethanol (no headspace). It is very important that sufficient ethanol be
used, or the organisms will not be properly preserved. Existing water in the jar should not dilute the
concentration of ethanol below 70%.
NOTE: Composite samples can be transported back to the vehicle before adding ethanol if
necessary. In this case, fill the jar with stream water, which is then drained using the net (or
sieve) across the opening to prevent loss of organisms, and replace with ethanol.
8. Replace the cap on each jar. Slowly tip the jar to a horizontal position, then gently rotate the jar to mix
the preservative. Do not invert or shake the jar. After mixing, seal each jar with plastic tape.
9. Store labeled composite samples in a container with absorbent material that is suitable for use with
70% ethanol until transport or shipment to the laboratory.
6.5 Fish
6.5.1 Summary of Method
The fish sampling method is designed to provide a representative sample of the fish
community, collecting all but the rarest fish taxa inhabiting the site. It is assumed to accurately
represent species richness, species guilds, relative abundance, and anomalies. The goal is to
collect fish community data that will allow the calculation of an Index of Biotic Integrity (IBI) and
Observed/Expected (O/E) models. Backpack or barge electrofishing is the preferred method. If
electrofishing is not possible due to safety concerns, high turbidity, or extremes in conductivity,
complete the "Not Fished" section of the field form and comment why.
Streams with mean wetted widths less than 12.5 m will be electrofished in their entirety,
covering all available habitats. However, the time and effort necessary to sample reaches
greater than or equal to 12.5 m wide is prohibitive in the context of the survey, thus sub-
sampling is required. Sub-sampling is defined by 5-10 sampling zones, each starting at a
transect. In all instances electrofishing in wadeable systems should proceed in an upstream
direction using a single anode. Identification and processing offish should occur at the
completion of each subreach.
6.5.2 Equipment and Supplies
Table 6.5-1 shows the checklist of equipment and supplies required to complete the fish
assessment. This checklist is similar to the one presented in Appendix A, which is used at the
base location to ensure that all of the required equipment is brought to the site. Record fish
collection data on the Fish Collection Form, Side 1 (Fig. 6.5-1).
Table 6.5-1. Equipment and supplies — fish assessment at wadeable sites.
For collecting
samples
Electrofishing equipment (including • 1 Scalpel for slitting open large fish
variable voltage pulsator unit, wiring before preservation.
cables, generator, electrodes, dip . 1 container of 10% buffered formalin
nets, protective linesman gloves, . Severa, Leak-proof HOPE jars for fish
boots, and necessary safety voucher specimens (various sizes from
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For recording
measurements
equipment)
• Extra electrofishing unit batteries
• Scientific collection permit
• Digital camera with extra memory
card & battery
• 1 Laser rangefinder (optional)
• Linesman gloves
• Sample labels
• Sample Collection Form
• Clear tape strips
250 mL - 4 L)
• 2 non-conducting dip nets with 1/4" mesh
1 Minnow net for dipping small fish from
live well
• 2 measuring boards (3 cm size classes)
• 1 set Fish ID keys
• Field Operations Manual and/or
laminated Quick Reference Guide
• Soft (#2) lead pencils
• Fine-tip indelible markers
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s S
Figure 6.5-1. Fish Collection Form for Small Wadeable Streams, Side 1.
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!c
'o
O
a S
EH «
> -2
O
O
6
I
fe
O o
R
U
Figure 6.5-2. Fish Collection Form for Large Wadeable Streams (Subreach A-B).
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6.5.3 Sampling Procedure
At sites with a total reach length <500m, fishing will occur continuously for all habitats
along the entire sample reach (40 times the average stream width), regardless of catch. At sites
with a total reach length >500 m, sampling is accomplished using subreaches so that effort is
distributed along the entire reach. In these streams, electrofishing will occur in sample zones
beginning the zero mark at each transect on alternating banks (Figure 6.5-3). Determination of
the initial stream bank sampling location at transect A (i.e., right or left bank) is determined at
random. The crew should consist of one electrofishing operator, and one dip netter and an
optional bucket carrier (who may also have a net to aid in transferring fish to the livewell).
Sampling will proceed in an upstream direction from transect to transect.
The total reach extent fished in large wadeable streams (>12.5 m) is a minimum reach
length of 20 times the average stream width (20X) and a maximum reach length of 40 times the
average stream width (40X). The subsampling routine is similar to boatables. Fish each
subreach for a maximum of 700 seconds or until the next transect is reached. Begin sampling at
a randomly determined bank at the beginning of the subreach and fish an area approximately
8m wide in an upstream direction. Fish the subreach thoroughly, covering bank habitat as well
as midstream habitat for a maximum of 700 seconds. When 700 seconds are reached, stop
electrofishing unless you are "pushing" a large school of fish, in which case continue fishing until
you capture them (typically at some form of structure or physical barrier). At a minimum, 5
subreaches or 20 times the mean channel width is sampled. If 500 individuals are caught within
this 20X, you may stop sampling. If not, continue sampling subreaches on alternating banks
until 500 individuals are captured. Crews must complete each of the additional subreaches as
described above, do not stop in the middle of any subreach, even if the 500 fish minimum is
attained before the end of the subreach. To reduce stress and mortality, immobilized fish should
be netted immediately and deposited into a live-well for processing. For safety, all crew
members are required to wear non-breathable waders and insulated gloves. Polarized
sunglasses and caps to aid vision are also required. Table 6.5-2 presents the procedure for
electrofishing in wadeable streams.
Figure 6.5-3. Transect sample design for fish sampling at wadeable sites >500 m (>12.5m width).
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Table 6.5-2. Procedure for electrofishing at wadeable sites <500 m
1. Review all collecting permits to determine if any sampling restrictions are in effect for the site. In
some cases, you may have to cease sampling if you encounter certain listed species.
2. Search for fish even if the stream is extremely small, and it appears that sampling may produce no
specimens. If none are collected, check the "NONE COLLECTED" circle on the Fish Collection
Form. Explain why in comments section. Although not required, you may note amphibians and
reptiles captured in the Comments.
3. Backpack and barge tote electrofishing will be used in wadeable streams, and direction of fishing
will be in an upstream manner. If you do not sample, complete the "NOT FISHED" field on the Fish
Collection Form and comment why.
4. At sites with a total reach length <500 m, fishing will occur continuously for all habitats along the
entire sample reach. No subsampling.
5. Set unit to pulsed DC. Select initial voltage setting (150-400 V for high conductivity [>300 S/cm];
500-800 V for medium conductivity [100 to 300 S/cm]; 900-1100 V for low conductivity [< 100 S/cm]
waters). In waters with strong-swimming fish (length >200 mm), use a pulse rate of 30 Hz with a
pulse width of 2 m/sec. If mostly small fish are expected, use a pulse rate of 60-70 Hz. Start the
electrofisher, set the timer, and depress the switch to begin fishing. If fishing success is poor,
increase the pulse width first and then the voltage. Increase the pulse rate last to minimize mortality
or injury to large fish. If mortalities occur, first decrease pulse rate, then voltage, then pulse width.
Start cleared clocks. Note, some electrofishers do not meter all the requested header data; provide
what you can. If button time is not metered, estimate it with a stop watch and flag the data.
6. Once the settings on the electrofisher are adjusted properly to sample effectively and minimize
injury and mortality, begin sampling at the downstream end of the reach (Transect A) and fish in an
upstream direction. Depress the switch and slowly sweep the electrode from side to side. Sample
all habitats and available cut-bank and snag habitat as well. Move the anode wand into cover with
the current off, turn the anode on when in the cover, and then remove the wand quickly to draw fish
out. In fast, shallow water, sweep the anode and fish downstream into a net. Be sure that deep,
shallow, fast, slow, complex, and simple habitats are all sampled. In stretches with deep pools, fish
the margins of the pool as much as possible, being extremely careful not to step or slide into deep
water. Keep the cathode near the anode if fish catch is low.
7. Depending upon crew size, there may be from 2 to 3 people fishing small wadeable sites. Crews
may choose to have more than one person holding a net, but no more than one person should
be netting at any one time. For example, in a wide stream there may be a netter on both sides of
an operator. As the operator moves the probe from the left bank to the right bank the netters will
remain on one side or the other and only one netter will be actively netting at any one time. The
same fishing effort can be accomplished with 1 netter moving from side to side with the probe.
8. The netter, with the net 1 to 2 ft from the anode, follows the operator, nets stunned individuals, and
places them in a bucket.
9. Continue upstream until the next transect is reached. Process fish and/or change water after each
subreach to reduce mortality and track sampling effort.
10. Complete header information on the Fish Collection Form Small Wadeable.
11. Repeat Steps 6 through 9 until the last subreach is finished.
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Table 6.5-3. Procedure for electrofishing at wadeable sites >500 m
1. Review all collecting permits to determine if any sampling restrictions are in effect for the site. In
some cases, you may have to cease sampling if you encounter certain listed species.
2. Search for fish even if the stream is extremely small, and it appears that sampling may produce no
specimens. If none are collected, check the "NONE COLLECTED" circle on the Fish Collection
Form. Explain why in comments section. Although not required, you may note amphibians and
reptiles captured in the Comments.
3. Backpack and barge tote electrofishing will be used in wadeable streams, and direction of fishing
will be in an upstream manner. If you do not sample, complete the "NOT FISHED" field on the Fish
Collection Form and comment why.
4. Fishing will occur in sample zones of approximately 8M in width with the zero mark at each transect
on alternating banks.
5. Set unit to pulsed DC. Select initial voltage setting (150-400 V for high conductivity [>300 S/cm];
500-800 V for medium conductivity [100 to 300 S/cm]; 900-1100 V for low conductivity [< 100 S/cm]
waters). In waters with strong-swimming fish (length >200 mm), use a pulse rate of 30 Hz with a
pulse width of 2 m/sec. If mostly small fish are expected, use a pulse rate of 60-70 Hz. Start the
electrofisher, set the timer, and depress the switch to begin fishing. If fishing success is poor,
increase the pulse width first and then the voltage. Increase the pulse rate last to minimize mortality
or injury to large fish. If mortalities occur, first decrease pulse rate, then voltage, then pulse width.
Start cleared clocks. Note, some electrofishers do not meter all the requested header data; provide
what you can. If button time is not metered, estimate it with a stop watch and flag the data.
6. Once the settings on the electrofisher are adjusted properly to sample effectively and minimize
injury and mortality, begin sampling at the downstream end of the reach (Transect A). Randomly
choose a bank on which to start and fish in an upstream direction within 8 M of the chosen bank.
Depress the switch and slowly sweep the electrode from side to side sampling all habitats
thoroughly and available cut-bank and snag habitat as well. Move the anode wand into cover with
the current off, turn the anode on when in the cover, and then remove the wand quickly to draw fish
out. In fast, shallow water, sweep the anode and fish downstream into a net. Be sure that deep,
shallow, fast, slow, complex, and simple habitats are all sampled. In stretches with deep pools, fish
the margins of the pool as much as possible, being extremely careful not to step or slide into deep
water. Keep the cathode near the anode if fish catch is low.
7. When using a barge or pram, the minimum crew size for electrofishing is three. The barge
operator must remain actively at the control box and navigate the barge. The probe operator will
use one probe. Depending upon crew size, there may be from 1 to 2 people additional crew
members. Crews may choose to have more than one person holding a net, but no more than one
person should be netting at any one time. For example, in a wide stream there may be a netter
on both sides of an operator. As the operator moves the probe from the left bank to the right bank
the netters will remain on one side or the other and only one netter will be actively netting at any
one time. The idle netter can assist the active netter by depositing fish into the live well. The same
fishing effort can be accomplished with one netter moving from side to side with the probe.
8. Continue upstream for a maximum of 700 seconds. Process fish after each transect to reduce
mortality and track sampling effort by transect.
9. Continue sampling subreaches at alternating banks until Transect F is reached. If less than 500
fish have been collected from the first five subreaches, continue sampling additional subreaches
along alternating banks until 500 individuals are captured, or at a maximum, subreach J-K is
finished. Crews must complete each of the additional subreaches as described above, do not stop
in the middle of any subreach, even if the 500 fish minimum is attained before the end of the
subreach.
10. Complete header information on the Fish Collection Form Large Wadeable/Boatable/Raftable.
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6.5.4 Processing Fish
Processing of fish must be completed at the end of each transect; however, if fish show
signs of stress (e.g., loss of righting response, gaping, gulping air, excessive mucus), change
water or stop fishing and initiate processing. Similarly, State- and Federally-listed threatened or
endangered species or large game fish should be processed and released as they are captured.
If periodic processing is required, fish should be released in a location that prevents the
likelihood of their recapture.
For streams <12.5 m wide, use the Fish Collection Form Small Wadeable. For streams
>12.5 m wide, use the Fish Collection Form - Large Wadeable/Boatable/Raftable. Taxonomic
identification and processing should only be completed on specimens greater than 25 mm total
length and by crew members designated as "fish taxonomic specialists" by EPA regional
coordinators. Fish are tallied by species, evaluated for maximum and minimum length, and
examined for the presence of DELT (Deformities, Eroded Fins, Lesions and Tumors) anomalies.
Common names of species should follow those established under the American Fisheries
Society's publication, "Common and Scientific Names of Fishes from the United States, Canada
and Mexico" (Nelson, et al. 2004). A list of species common to freshwater systems of the United
States is presented in Appendix D.
Species not positively identified in the field should be separately retained (up to 20
individuals per species) for laboratory identification. Common names for retained species should
be assigned as "unknown", followed by its common family name and sequential lettering to
designate separate species (e.g., UNKNOWN SCULPIN A). For large wadeable streams, each
transect has its own form. Following positive laboratory identification, field form information
should be updated to reflect the actual species count and number in the Final Count field. For
example, if a sample of 20 specimens of species A is later identified as 15 individuals of species
A and 5 of species B, the Final Count of species A should be corrected by assigning 25% to
species B and 75% to species A. Table 6.5-4 presents the procedure for processing fish.
Table 6.5-4. Procedure for processing fish at wadeable sites
1. Complete all header information accurately and completely. If no fish were collected, complete the
"NONE COLLECTED" field on the Fish Collection Form.
2. Complete the information on the Fish Gear and Voucher/Tissue Sample Information Form.
3. For small wadeable streams (<12.5 m) use the Fish Collection Form - Small Wadeable. For large
wadeable streams (>12.5 m) use the Fish Collection Form - Large Wadeable/Boatable/ Raftable.
4. For small wadeables, use one form for the entire reach.
5. For large wadeables, use one form persubreach and indicate Subreach on form in "SUBREACH"
Field.
6. Only identify and process individuals > 25mm in total length, ideally handling specimens only once.
Record the common name on the first blank line in the "COMMON NAME" Field of the Fish
Collection Form.
7. Fill in the Tag Number. The tag number is a number starting with 01 and continuing sequentially to
a number equal to the total number of species collected within the entire sample reach. Each
reoccurrence of a species within the entire reach should be assigned the same tag number as it was
assigned initially. For example, if a bluegill is assigned tag number 01 when processing fish from
the first subreach, all bluegills from the other subreaches will also be assigned tag number 01. The
purpose of the tag number is to connect species identifications with subsequent verification and
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voucher collections.
8. If a species cannot be positively identified, assign it a sequential tag number in the Tag Number
Field and leave the "COMMON NAME" Field Blank. Flag this line and indicate in the "COMMENT"
field its common family name (e.g., UNKNOWN SCULPIN A). Retain a maximum subsample of 20
individuals for in-house laboratory identification of Unknowns. Do not include the number of each
species retained solely for in-house lab verification in the Voucher Count column of the fish
collection form. This column is reserved only for those fish that are to be sent in for independent re-
identification as part of a complete voucher collection.
9. Process species listed as threatened and endangered first and return individuals immediately to the
stream. Photograph specimens for verification purposes if conditions permit and stress to individuals
will be minimal. Indicate if photographed on Fish Collection Form. If individuals are killed, prepare
them as verification specimens and preserve noting them in the "MORTALITY COUNT" field.
10. Tally the number of individuals of each species collected in the "TALLY" box on the Fish Collection
Form and record the total number in the "COUNT" field on the form.
11. Measure the total length of the largest and smallest individual to provide a size range for the
species. Record these values in the "LENGTH" area of the Fish Collection Form. For small
wadeables, this is done for the entire reach. For large wadeables, this is recorded by transect.
12. Examine each individual for external anomalies and tally those observed. Identify external
anomalies including missing organs (eye, fin), skeletal deformities, shortened operculum, eroded
fins, irregular fin rays or scales, tumors, lesions, ulcerous sores, blisters, cysts, blackening, white
spots, bleeding or reddening, excessive mucus, and fungus. After all of the individuals of a species
have been processed, record the total number of individuals affected in the "ANOMALIES" area of
the Fish Collection Form. For small wadeables, this is done for the entire reach. For large
wadeables, this is recorded by transect
13. Record total number of mortalities in the "MORTALITY COUNT" field due to electrofishing or
handling on the Fish Collection Form.
14. Follow the appropriate procedure to prepare voucher specimens and/or to select specimens for
tissue samples. Release all remaining individuals so as to avoid their recapture.
15. For any line with a fish name on the Fish Collection Form, ensure that all spaces on that line are
filled in with a number, even if it is zero.
16. Repeat Steps 1 through 10 for all other species and subreaches.
6.5.5 Taxonomic Quality Assurance/Quality Control
6.5.5.1 Sample Preservation
Fish retained for laboratory identification/verification or voucher purposes should be
placed in a large sample jar containing a 10% buffered formalin solution in a volume equal to or
greater than the total volume of specimens. Individuals larger than 200 mm in total length
should be slit along the right side of the fish in the lower abdominal cavity to allow penetration of
the solution.
Fish retained for laboratory identification or as vouchers should be preserved in the field
following the precautions outlined in the MSDS. All personnel handling 10% buffered formalin
must read the MSDS (Appendix D). Formalin is a potential carcinogen and should be used
with extreme caution, as vapors and solution are highly caustic and may cause severe
irritation on contact with skin, eyes, or mucus membranes. Wear vinyl or nitrile gloves
and safety glasses, and always work in a well-ventilated area.
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6.5.5.2 Laboratory Identification
Fish that are difficult to identify in the field should be kept for laboratory identification or
to verify difficult field identifications. Table 6.5-5 outlines the laboratory identification process
and completing the Fish Collection Form. Field crews may use a supplemental Fish
Identification Lab sheet such as that shown in Figure 6.5-4 for internal laboratory use only.
Crews should retain the Fish verification sample - contact your regional EPA coordinator if you
cannot store the samples at your facility.
Do not include the number of each species retained solely for in-house lab verification in
the Voucher Count column of the fish collection form. This column is reserved only for those
fish that are to be sent in for independent re-identification as part of a complete voucher
collection.
Field crews should not retain the Fish Collection Form(s) if the laboratory identification
process cannot be completed within a short period of time. If the time needed to complete the
identification/verification is expected to exceed two weeks, make copies of the Fish Collection
Form(s) and send the entire pack of original data forms to the Information Management
Coordinator. When the identification/verification process is complete, make the necessary
changes to the copied Fish Collection Form(s) and send them as soon as possible to the
Information Management Coordinator as well.
Table 6.5-5. Procedure for laboratory identification offish samples.
1. Fish may be retained for routine laboratory identification and verification purposes. Fish tags are
provided with each site kit. Crews may use these tags at their discretion in order to identify fish at
their laboratory.
2. Retained fish should be placed in a large sample jar containing a 10% buffered formalin solution in a
volume equal to or greater than the total volume of specimens. Individuals larger than 200mm in
total length should be slit along the right side of the fish in the lower abdominal cavity to allow
penetration of the solution.
3. Following fixation for 5 to 7 days, the volume of formalin should be properly discarded and replaced
with tap water for soaking specimens over a 4-5 day period. Soaking may require periodic water
changes and should continue until the odor of formalin is barely detectable. Final storage of
specimens is done in 45%-50% isopropyl alcohol or 70% ethanol. Formalin is a potential carcinogen
and should be used with extreme caution, as vapors and solution are highly caustic and may cause
severe irritation on contact with skin, eyes, or mucus membranes. Wear vinyl or nitrile gloves and
safety glasses, and always work in a well-ventilated area.
4. Formalin must be disposed of properly. Contact your regional EPA coordinator if your laboratory
does not have the capability of handling waste formalin.
5. Unknown fish are identified to species in the laboratory. You may use a Fish Identification Lab
Sheet such as the one presented in Figure 6.5-4.
6. Fill in the Unknown species name in the "COMMON NAME" field of the Fish Collection Form and
make certain the "FINAL COUNT" field is correct.
7. If species field identifications were incorrect, correct the "COMMON NAME" Field by completely
erasing the Common Name and replacing the correct name. Add an additional Common Name if
needed. Make certain the "FINAL COUNT" field is correct. If the "COMMON NAME" Field was
incorrect or cannot be cleanly erased, cross out the line of data and fill out a new line with the
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I correct "COMMON NAME" and "FINAL COUNT". I
6.5.5.3 Voucher Specimens
Approximately 10% of each field crews' sites will be randomly pre-selected for re-
identification by an independent taxonomist. A minimum of one complete voucher is required for
each person performing field taxonomy and will consist of either preserved specimen(s) or
digital images representative of all species in the sample, including common species. Multiple
specimens per species can be used as vouchers, if necessary (i.e., to document different life or
growth stages, or sexes). Note that a complete sample voucher does not mean that all
individuals of each species will be vouchered, only enough so that independent verification can
be achieved.
Digital images should be taken as voucher documentation for species that are
recognized as Rare, Threatened, or Endangered - they should not be killed. Digital images
should also be taken of fish specimens too large for preservation.
Certain states or regions may require that more fish vouchers are taken. Check with
your state/regional coordinators to determine if your team will be required to collect complete
vouchers at more than 10% or your sites.
For the sample voucher, specimen containers should be labeled with the sample
number, site ID number, site name, and collection date. There should be no taxonomic
identification labels in or on the container, or in any of the digital photos.
Choose individual specimens that are intact and in good condition, such that re-
identification will be possible. Fish that are damaged, have significant scale loss or those that
have been dead for a significant amount of time prior to preservation should be avoided if
possible. Fish in pristine condition and those possessing clear identification characteristics are
preferred. Additionally, fish that are preserved while still live will typically flare their fins and gills
and will allow for easier re-identification in the laboratory.
Place one or more representative specimens of each species in plastic mesh sleeves
along with one of the corresponding tag number labels provided in your site kit. (Several fish
may be placed in a single mesh sleeve, as long as they are of the same species). Ensure that
the tag numbers in the voucher collection match the tag numbers on the fish collection data
forms. Seal both ends of the mesh sleeve with zip ties and place it inside the voucher collection
jar with the appropriate preservative. Unknown fish may be identified in the laboratory as
described in section 5.5.5.2 and subsequently included in the voucher collection.
Record the total number of each fish species retained for voucher purposes in each
subreach on the fish collection form. Record the voucher sample ID number on the fish gear /
voucher / fish tissue collection form. If no voucher is prepared for the site, fill in the "no
vouchers preserved" circle on the fish gear form.
Table 6.5-6. Procedure for vouchering fish samples.
1. Approximately 10% of each field crews' sites will be randomly pre-selected for re-identification by
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an independent taxonomist. A minimum of one complete voucher is required for each person
performing field taxonomy and will consist of either preserved specimen(s) and/or digital images
representative of all species in the sample, even common species.
2. Take digital images as voucher documentation for species that are recognized as Rare,
Threatened, or Endangered; or when fish specimens are too large for preservation.
3. For the sample voucher, label the specimen containers with the sample number, site ID number,
site name, and collection date. Do not put taxonomic identification labels in or on the container.
4. Place one or more representative specimens of each species in plastic mesh sleeves along with
one of the corresponding tag number labels provided in your site kit. (Several fish may be placed in
a single mesh sleeve, as long as they are of the same species).
5. Ensure that the tag numbers in the voucher collection match the tag numbers on the fish collection
data forms.
6. Seal both ends of the mesh sleeve with zip ties and place it inside the voucher collection jar with
the appropriate preservative.
7. Unknown fish may be identified in the laboratory as described in section 5.5.5.2 and subsequently
included in the voucher collection.
8. Record the total number of each fish species retained for voucher purposes in each subreach on
the fish collection form.
9. Record the voucher sample ID number on the fish gear / voucher / fish tissue collection form.
10. If no voucher is prepared for the site, fill in the "no vouchers preserved" circle on the fish gear form.
6.5.5.4 Photovouchering
Digital imagery should be used for fish species that cannot be retained as preserved
specimens (e.g., RTE species; very large bodied; or very common). Views appropriate and
necessary for an independent taxonomist to accurately identify the specimen should be the
primary goal of the photography. Additional detail for these guidelines is provided in Stauffer et
al. (2001), and is provided to all field crews as a handout.
The recommended specifications for digital images to be used for photovouchering
include: 16-bit color at a minimum resolution of 1024x768 pixels; macro lens capability allowing
for images to be recorded at a distance of less than 4 cm; and built-in or external flash for use in
low-light conditions. Specimens should occupy as much of the field of view as possible, and the
use of a fish board is recommended to provide a reference to scale (i.e., ruler or some
calibrated device) and an adequate background color for photographs. Information on Station
ID, Site Name, Date and a unique species ID (i.e., A, B, C, etc.) should also be captured in the
photograph, so that photos can be identified if file names become corrupted. All photovouchered
species should have at least a full-body photo (preferably of the left side of the fish) and other
zoom images as necessary for individual species, such as lateral line, ocular/oral orientation, fin
rays, gill arches, or others. It may also be necessary to photograph males, females, or juveniles.
Images should be saved in medium- to high-quality jpeg format, with the resulting file
name of each picture noted one the Fish Collection Form. It is important that time and date
stamps are accurate as this information can also be useful in tracking the origin of photographs.
It is recommended that images stored in the camera be transferred to a PC or storage device at
the first available opportunity. At this time the original file should be renamed to follow the logic
presented below:
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F01_CT003_20080326_A.jpg
Where:
F = fish
01 = tag number
CT003 = state (Connecticut) and site number
20080326 = date (yyyymmdd)
A = first of several pictures of same fish (e.g., A, B, C)
Field crews should maintain files for the duration of the sampling season. Notification
regarding the transfer of all images to the existing database will be provided at the conclusion of
the sampling.
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Fish Identification Lab Sheet
Site ID
Identified / /
Preservative(Field/Lab)
Used
Date Collected_
~ ID'd by_
Date Data Corrected on Field Sheet
_Date(s)
Keys
Initials
Tag
no.
Photo (P)
or
Specimen
(S)
Common
Name
(Field)
Common
Name
(Lab)
Count
Transect
(if known)
PhotoFile
(Field)
PhotoFile
(Final)
Figure 6.5-4. Fish Identification Lab Sheet.
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6.6 Fecal Indicator (Enterococci)
6.6.1 Summary of Method
You will collect a fecal indicator sample at the last transect (Transect K) after all other
sampling is completed. Use a pre-sterilized, 250 ml bottle and collect the sample approximately
1 m off the bank at about 0.3 meter (12 inches) below the water. Following collection, place the
sample in a cooler, chill for at least 15 minutes, and maintain on ice prior to filtration of four 50
ml_ volumes. (Samples must be filtered and frozen on dry ice within 6 hours of collection). In
addition to collecting the sample, look for signs of disturbance throughout the reach that would
contribute to the presence of fecal contamination to the waterbody. Record these disturbances
on the Site Assessment Form (Figure 7-2).
6.6.2 Equipment and Supplies
Table 6.6-1 provides the equipment and supplies needed for field crews to collect the
fecal indicator sample. Record the fecal indicator sample data on the Sample Collection Form
(Figure 6.1-3).
Table 6.6-1. Equipment and supplies list for fecal indicator sampling at wadeable sites
For collecting samples
nitrile gloves
pre-sterilized, 250 ml sample bottle
sodium thiosulfate tablet
Wet ice
cooler
For recording
measurements
Sample Collection Form
Site Assessment Form
Fecal Indicator sample labels (4 vial labels and 1 bag label)
Pencils (for data forms)
Fine-tipped indelible markers (for labels)
Clear tape strips
6.6.3 Sampling Procedure
Table 6.6-2 provides the procedure for collecting fecal indicator (i.e., Enterococci)
samples at wadeable sites.
Table 6.6-2. Procedure for fecal indicator (Enterococci) sample collection at wadeable sites
Collect the Enterococci Sample
1. Put on nitrile gloves.
2. Select a sampling location at transect K that is approximately 1 m from the bank and approximately
1 m deep. Approach the sampling location slowly from downstream or downwind.
3. Lower the un-capped, inverted 250 ml sample bottle to a depth of 1 foot below the water surface,
avoiding surface scum, vegetation, and substrates. Point the mouth of the container away from the
body or boat. Right the bottle and raise it through the water column, allowing bottle to fill completely.
If the depth does not reach 1 foot along the transect at 1 m from the bank, take the sample and flag
it on the field form.
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4. After removing the container from the water, discard a small portion of the sample to allow for proper
mixing before analyses.
5. Add the sodium thiosulfate tablet, cap, and shake bottle 25 times.
6. Store the sample in a cooler on ice to chill (not freeze). Chill for at least 15 minutes and do not hold
samples longer than 6 hours before filtration and freezing.
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7.0 FINAL SITE ACTIVITIES
The activities described in this section apply to both wadeable and non-wadeable sites.
Prior to leaving the site, make a general visual assessment of the site and its surrounding
catchment. The objective of the site assessment is to record observations of catchment and site
characteristics that are useful for future data interpretation, ecological value assessment,
development of associations, and verification of stressor data. Your observations and
impressions are extremely valuable.
You will filter and process the fecal indicator, chlorophyll a, and periphyton samples.
Conduct a final check of the data forms, labels and samples. The purpose of the second check
of data forms, labels and samples is to assure completeness of all sampling activities. Finally,
clean and pack all equipment and supplies, and clean the launch site and staging areas. After
you leave the site, report the sampling event to the Information Management Coordinator, and
ship or store the samples. Activities described in this section are summarized in Figure 7-1.
COMPLETE SITE
ASSESSMENT
(4 People)
REVIEW DATA FORMS
(Crew Leader)
• Completeness
• Accuracy
• Legibility
• Flags/Comments
FILTER, PRESERVE, &
INSPECT SAMPLES
(3 People)
• Complete
• Sealed
• Ice packs
• Packed for transport
REVIEW SAMPLE LABELS
(Crew Leader)
• Completeness
• Accuracy
• Legibility
• Cross-check with forms
INSPECT BOAT, MOTOR,
TRAILER, AND NETS FOR
PRESENCE OF PLANT AND
ANIMAL MATERIAL, AND
CLEAN THOROUGHLY
(3 People)
PACK EQUIPMENT AND
SUPPLIES FOR TRANSPORT
(2 People)
LOAD BOAT ONTO TRAILER;
CLEAN UP LAUNCH SITE
AND STAGING AREA
(2 People)
LEAVE SITE
COMMUNICATIONS
SHIP SAMPLES
Figure 7.1. Final site activities summary.
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7.1 General Site Assessment
Complete the Site Assessment Form (Figure 7-2) after sampling, recording all
observations from the site that were noted during the course of the visit. This Site Assessment
Form is designed as a template for recording pertinent field observations. It is by no means
comprehensive, and any additional observations should be recorded in the General Assessment
section.
7.1.1 Watershed Activities and Disturbances Observed
Record any of the sources of potential stressors listed in the "Watershed Activities and
Disturbances Observed" section on the Site Assessment Form (Figure 7-2). Include those that
were observed while on the site, while driving or walking through the site and catchment, or
while flying over the site and catchment. For activities and stressors that you observe, rate their
abundance or influence as low (L), moderate (M), or heavy (H) on the line next to the listed
disturbance. Leave the line blank for any disturbance not observed. The distinction between
low, moderate, and heavy will be subjective. For example, if there are two to three houses on a
site, circle "L" for low next to "Houses." If the site is ringed with houses, rate it as heavy (H).
Similarly, a small patch of clear-cut logging on a hill overlooking the site would rate a low
ranking. Logging activity right on the site shore, however, would get a heavy disturbance
ranking. This section includes residential, recreational, agricultural, industrial, and stream
management categories.
7.1.2 Site Characteristics
Record observations regarding the general characteristics of the site on the Site
Assessment Form (Figure 7-2). When assessing these characteristics, look at a 200 m riparian
distance on both banks. Rank the site between "pristine" and "highly disturbed", and between
"appealing" and "unappealing." Document any signs of beaver activity and flow modifications.
Record the dominant land use and forest age class. Document the weather conditions on the
day of sampling, and any extreme weather conditions just prior to sampling.
7.1.3 General Assessment
Record any additional information and observations in this narrative section. Information
to include could be observations on biotic integrity, vegetation diversity, presence of wildlife,
local anecdotal information, or any other pertinent information about the site or its catchment.
Record any observations that may be useful for future data interpretation.
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VISUAL ASSESSMENT FORM - WADEABLE/BOATABLE (Front) *'
SITE ID: FW08 XX £>OO
DATE:
WATERSHED ACTIVITIES AND DISTURBANCES OBSERVED dutrcaiiy Bl»nN=Nat ab erved L=lo* M=Mod«ate, H=Heavy)
Residential | RecreationalI Agricultural f Industrial
M H «.«i*-».l
M H t*H>'mm*a
L M H cwi.irycia.
L M H Psc«. C^at
L M H e
L M H P.
L M H Pnnnlta. p.
L M H T,mu»
L M M &*te* »H
Agricultural
L M H Cray i'O
L M (^P_U**
L M H
L M H
L M H
L M H
L M H
L M H
L M H
L M M
L M H
Stream Management
L M H I ™,»
L M M u-wsxna
L M H i*im
L M H DnfKlgi
L H H c>i«i«
L M H «M>r
L M H r*»
L M H u»i»
Waterbody
Character
Beaver
Dominant
Lan d U se
SITE CHARACTERiSTICS {200 m radius)
Pristine O 5
Appealing O 5
O 4
O 4
O 3
• 3
* 2
O2
O 1
O1
HigMy Disturbed
Unappealing
Beaver Flow Modifications:
O Rare
O Minor
O Common
O Major
Aroamrr O For&st O Agocuttuw • Rarsge
Sf Forest, DoMiinanl Ag« ^ ^^ ^
Cl««» O ° • 2S >"• O 2S - 75 yr». () > 75 yrs.
O yrtsin
O SubyrbaruTawn
WEATHER
GENERAL ASSESSMENT (BioHe Integrity, VegetaUon diversity, Lcx:al anecdotal Intormrtonj
Locum Juir
Of=
Rev: i: M4. ;;W3 Vttusll Assmsinml ^ tJRSA
B^B ^
Figure 7.2. Site Assessment Form.
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7.2 Processing the Fecal Indicator, Chlorophyll a, and Periphyton Samples
7.2.1 Equipment and Supplies (Fecal Indicator)
Table 7-1 provides the equipment and supplies needed for field crews to collect the fecal
indicator sample.
Table 7.1. Equipment and supplies list for fecal indicator sample
For processing samples
Nitrile gloves
sterile screw-cap 50-mL centrifuge tube
Sterile filter holder, Nalgene 145/147
Vacuum pump (electric pump may be used if available)
Sterile phosphate buffered saline (PBS)
Osmotics 47 mm polycarbonate 0.4 urn sterile filters
Sterile disposable forceps
4 sterile microcentrifuge tubes containing sterile glass beads
Dry ice
Cooler
Field Operations Manual or laminated Quick Reference Guide
For recording
measurements
Sample Collection Form
Soft (#2) lead pencils for recording data on field forms
Fine-tipped indelible markers for filling out sample labels
Fecal Indicator sample labels (4 vial labels and 1 bag label)
Clear tape strips for covering labels
7.2.2 Procedures for Processing the Fecal Indicator Sample
The fecal indicator sample must be filtered before the chlorophyll a and periphyton
samples, since the filtering apparatus needs to be sterile for this sample. The procedures for
processing the fecal indicator sample are presented in Table 7-2. The sample must be filtered
and frozen within 6 hours of collection.
Table 7.2. Processing procedure—fecal indicator sample
Processing procedure—fecal indicator filter blank (to be done at Revisit sites only)
Enterococci filter blanks will be prepared at all revisit sites during the first visit (see Fig. 8-1). Prepare the
filter blanks before filtering the river sample.
1. Set up sample filtration apparatus using same procedure as used for the river sample. Chill Filter
Extraction tubes with beads on dry ice.
2. Aseptically transfer 4 polycarbonate filters from filter box to base of opened Petri dish. Close filter box
and set aside.
3. Remove cellulose nitrate (CN) filter (the filter with grid design on it) from funnel and discard. Be sure
to leave the support pad in the filter funnel.
4. Load filtration funnel with sterile polycarbonate filter on support pad (shiny side up).
5. Measure 10-mL of the chilled phosphate buffered saline (PBS) in the sterile graduated centrifuge
tube and pour into filter funnel.
6. Replace cover on filter funnel and pump to generate a vacuum (do not generate more than 7 inches
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Table 7.2. Processing procedure—fecal indicator sample
of Hg of pressure). Keep pumping until all liquid is in filtrate collection flask.
7. Remove filter funnel from base without disturbing filter. Using sterile disposable forceps remove the
filter (touching only the filter edges) and fold it in half, in quarters, in eighths, and then in sixteenths
(filter will be folded 4 times).
8. Insert filter into chilled filter extraction tube (with beads) open end down. Replace and tighten the
screw cap, insert tube(s) into bubble wrap bag on dry ice for preservation during transport and
shipping.
9. Label the samples as "blank" on the label and field form, and package and submit these samples to
the lab with the standard samples.
10. Repeat steps 4 to 9 for the remaining three 10-mL volumes of PBS to be filtered.
Processing procedure—fecal indicator samples (All sites)
1. Put on nitrile gloves.
2. Set up sample filtration apparatus on flat surface and attach vacuum pump. Set-out 50-mL sterile
centrifuge tube, sterile 60-mm Petri dish, 2 bottles of chilled phosphate buffered saline (PBS),
Osmotics 47 mm polycarbonate sterile filter box, and 2 filter forceps.
3. Chill Filter Extraction tubes with beads on dry ice.
4. Aseptically transfer 4 polycarbonate filters from filter box to base of opened Petri dish. Close filter box
and set aside.
5. Remove cellulose nitrate (CN) filter (the filter with grid design on it) from funnel and discard. Be sure
to leave the support pad in the filter funnel.
6. Load filtration funnel with sterile polycarbonate filter on support pad (shiny side up).
7. Shake sample bottle(s) 25 times to mix well.
8. Measure 25-mL of the mixed water sample in the sterile graduated centrifuge tube and pour into filter
funnel.
9. Replace cover on filter funnel and pump to generate a vacuum (do not generate more than 7 inches
of Hg of pressure). Keep pumping until all liquid is in filtrate collection flask.
10. If the first 25 mL volume passes readily through the filter, add another 25 mL and continue filtration. If
the filter clogs before completely filtering the first or second 25 mL volume, discard the filter and
repeat the filtration using a lesser volume.
11. Pour approx. 10-mL of the chilled phosphate buffered saline (PBS) into the graduated PP tube used
for the sample. Cap the tube and shake 5 times. Remove the cap and pour rinsate into filter funnel to
rinse filter.
12. Filter the rinsate and repeat with another 10 mL of phosphate buffered saline (PBS).
13. Remove filter funnel from base without disturbing filter. Using sterile disposable forceps remove the
filter (touching only the filter edges) and fold it in half, in quarters, in eighths, and then in sixteenths
(filter will be folded 4 times).
14. Insert filter into chilled filter extraction tube (with beads) open end down. Replace and tighten the
screw cap, insert tube(s) into bubble wrap bag on dry ice for preservation during transport and
shipping.
15. Record the volume of water sample filtered through each filter and the volume of buffer rinsate each
filter was rinsed with on the Sample Collection Form, Side 2. Record the filtration start time and finish
time for each sample.
16. Repeat steps 6 to 15 for the remaining three 50-mL sub-sample volumes to be filtered.
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7.2.3 Equipment and Supplies (Chlorophyll a from Water Sample)
Table 7-3 provides the equipment and supplies needed to process the chlorophyll a
water sample.
Table 7.3. Equipment and supplies list for chlorophyll a processing
For filtering chlorophyll a sample
Whatman GF/F 0.7 |jm glass fiber filter
Filtration apparatus with graduated filter holder
Vacuum pump (electric pump may be used if available)
50-mL screw-top centrifuge tube
Aluminum foil square
Dl water
Nitrile gloves
Forceps
For recording measurements
Sample Collection Form
Sample labels
#2 pencils
Fine-tipped indelible markers
Clear tape strips
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7.2.4 Procedures for Processing the Chlorophyll a Water Sample
The procedures for processing chlorophyll a water samples are presented in Table 7-4.
Whenever possible, sample processing should be done in subdued light, out of direct sunlight.
Table 7.4. Processing procedure—chlorophyll a sample
1. Put on nitrile gloves.
2. Use clean forceps to place a Whatman GF/F 0.7 |jm glass fiber filter in the graduated filter holder
apparatus with the gridded side of the filter facing down.
3. Pour 250 ml of water into the filter holder, replace the cap, and use the vacuum pump to draw the
sample through the filter. If 250 ml of site water will not pass through the filter, change the filter, rinse
the apparatus with Dl water, and repeat the procedures using 100-mL of site water. NOTE: IF the
water is green or turbid, use a smaller volume to start with.
4. Rinse the upper portion of the filtration apparatus thoroughly with Dl water to include any remaining
cells adhering to the sides and pump through the filter (do not exceed 7 inches of Hg). Monitor the
level of water in the lower chamber to ensure that it does not contact the filter or flow into the pump.
5. Observe the filter for visible color. If there is visible color, proceed; if not, repeat steps 3 & 4 until color
is visible on the filter or until a maximum of 2,000 ml have been filtered. Record the actual sample
volume filtered on the Sample Collection Form.
6. Remove the bottom portion of the apparatus and pour off the water from the bottom.
7. Remove the filter from the holder with clean forceps. Avoid touching the colored portion of the filter.
Fold the filter in half, with the colored side folded in on itself.
8. Place the folded filter into a 50-mL screw-top centrifuge tube and cap. Record the sample volume
filtered on a chlorophyll label and attach it to the centrifuge tube. Ensure that all written information is
complete and legible. Cover with a strip of clear tape. Wrap the tube in aluminum foil and place in a
self-sealing plastic bag. Place this bag between two small bags of ice in a cooler.
7.2.5 Equipment and Supplies (Periphyton Sample)
Table 7-5 lists the equipment and supplies needed to process the periphyton sample.
Table 7.5. Equipment and supplies list for periphyton sample processing
For filtering
periphyton
samples
Whatman 47 mm 0.7 micron GF/F glass fiber filter
Whatman 47 mm 1.2 micron GF/C glass fiber filter
Filtration apparatus with graduated filter holder
Vacuum pump (electric pump may be used)
25 or 50-mL graduated cylinder
4 50 ml screw-top centrifuge tubes
60-mL syringe
Aluminum foil squares
Forceps
deionized water in wash bottle
plastic electrical tape
dry ice
wet ice
coolers
For data
recording
Sample Collection Form
Sample labels
Pencils
Fine-tipped indelible markers
Clear tape strips
7.2.6 Procedures for Processing the Periphyton Samples
Four different types of laboratory samples are prepared from the composite index
samples: an ID/enumeration sample (to determine taxonomic composition and relative
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abundances), a chlorophyll sample, a biomass sample (for ash-free dry mass [AFDM]), and
an acid/alkaline phosphatase activity (APA) sample. All the sample containers required for an
individual site should be sealed in plastic bags until use to avoid external sources of
contamination (e.g., dust, dirt, or mud) that are present at site shorelines.
7.2.6.1 ID/Enumeration Sample
Prepare the ID/Enumeration sample as a 50-mL aliquot from the composite index
sample, following the procedure presented in Table 7-6. Preserve each sample with Lugol's.
Record the sample ID number from the container label and the total volume of the sample in the
appropriate fields on the Sample Collection Form as shown in Figure 5.1-2 and 6.1-2. Store the
preserved samples upright in a container containing absorbent material.
Table 7.6. Procedure for ID/enumeration samples of periphyton
1. Prepare a sample label (with a sample ID number) for the Periphyton ID sample. Record the volume of
the subsample (typically 50 ml) and the volume of the composite index sample on the label. Attach
completed label to a 50-mL centrifuge tube; avoid covering the volume graduations and markings.
Cover the label completely with a clear tape strip.
2. Record the sample ID number of the label and the total volume of the composite index sample on the
form.
3. Rinse a 60-mL syringe with deionized water.
4. Thoroughly mix the bottle containing the composite sample.
5. Withdraw 50 ml of the mixed sample into the syringe. Right after mixing, place the contents of syringe
sample into the labeled 50-mL centrifuge tube.
6. Use a syringe or bulb pipette to add 1 ml Lugol's to the tube. Cap the tube tightly and seal with plastic
electrical tape. Shake gently to distribute preservative.
7. Record the volume of the sample in the centrifuge tube (excluding the volume of preservative) in
"Assemblage ID Subsample Vol." field of the Sample Collection Form.
7.2.6.2 Chlorophyll Sample
Prepare the chlorophyll sample by filtering a 25-mL aliquot of the composite index
sample through a 47 mm 0.7 micron GF/F glass fiber filter. The procedure for preparing
chlorophyll samples is presented in Table 7-7. Chlorophyll can degrade rapidly when exposed to
bright light. If possible, prepare the samples in subdued light (or shade), filtering as quickly as
possible after collection to minimize degradation. Keep the glass fiber filters in a dispenser
inside a sealed plastic bag until use.
It is important to measure the volume of the sample being filtered accurately (±1 mL)
with a graduated cylinder. During filtration, do not exceed 7 inches of Hg to avoid rupturing cells.
If the vacuum pressure exceeds 7 inches of Hg, prepare a new sample. If the filter clogs
completely before all the sample in the chamber has been filtered, discard the sample and filter,
and prepare a new sample using a smaller volume of sample.
Table 7.7. Procedure for preparing chlorophyll samples of periphyton
1. Using clean forceps, place a Whatman GF/F 0.7 urn glass fiber filter on the filter holder gridded side
down. Use a small amount of deionized water from a wash bottle to help settle the filter properly.
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Attach the filter funnel to the filter holder and filter chamber, then attach the hand vacuum pump to the
chamber.
2. Rinse the sides of the filter funnel and the filter with a small volume of deionized water.
3. Rinse a 25-mL or 50-mL graduated cylinder three times with small volumes of deionized water.
4. Mix the composite sample bottle thoroughly.
5. Measure 25 ml (±1 ml) of sample into the graduated cylinder. • NOTE: Fora composite sample
containing fine sediment, allow grit to settle for 10 - 20 seconds before pouring the sample into the
graduated cylinder.
6. Pour the 25-mL aliquot into the filter funnel, replace the cap, and pull the sample through the filter
using the hand pump. Vacuum pressure from the pump should not exceed 7 inches of Hg to avoid
rupture of fragile algal cells. • NOTE: If 25 mL of sample will not pass through the filter, discard the
filter and rinse the chamber thoroughly with deionized water. Collect a new sample using a smaller
volume of sample, measured to±1 mL. Be sure to record the actual volume sampled on the sample
label and the Sample Collection Form.
7. Remove both plugs from the filtration chamber and pour out the filtered water in the chamber.
Remove the filter funnel from the filter holder. Remove the filter from the holder with clean forceps.
Avoid touching the colored portion of the filter. Fold the filter in half, with the colored sample (filtrate)
side folded in on itself. Place the folded filter in a 50 ml centrifuge tube. Discard filtered water.
8. Complete a periphyton sample label for chlorophyll, including the volume filtered, and attach it to the
centrifuge tube. Cover the label completely with a strip of clear tape. Place the centrifuge tube into a
self-sealing plastic bag.
9. Record the sample ID number of the label and the total volume of the composite index sample on the
form. Record the volume filtered in the "Chlorophyll" field on the Sample Collection Form. Double
check that the volume recorded on the collection form matches the total volume recorded on the
sample label.
10. Place the centrifuge tube containing the filter on dry ice.
7.2.6.3 Biomass Sample
Prepare the ash-free dry mass (AFDM) sample by filtering a 25-mL aliquot of the
composite index sample through a 47 mm 1.2 micron GF/C glass fiber filter. The procedure for
preparing AFDM samples is presented in Table 7-8. Keep the glass fiber filters in a dispenser
inside a sealed plastic bag until use.
It is important to measure the volume of the sample being filtered accurately (±1 mL)
with a graduated cylinder. During filtration, do not exceed 7 inches of Hg to avoid rupturing cells.
If the vacuum pressure exceeds 7 inches of Hg prepare a new sample. If the filter clogs
completely before all the sample in the chamber has been filtered, discard the sample and filter,
and prepare a new sample using a smaller volume of sample.
Table 7.8. Procedure for preparing ash-free dry mass (AFDM) samples of periphyton
1. Using clean forceps, place a Whatman 47 mm 1.2 micron GF/C glass fiber filters on the filter holder
gridded side down. Use a small amount of deionized water from a wash bottle to help settle the filter
properly. Attach the filter funnel to the filter holder and filter chamber, then attach the hand vacuum
pump to the chamber.
2. Rinse the sides of the filter funnel and the filter with a small volume of deionized water.
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3. Rinse a 25-mL or 50-mL graduated cylinder three times with small volumes of deionized water.
4. Mix the composite sample bottle thoroughly.
5. Measure 25 ml (±1 ml) of sample into the graduated cylinder. NOTE: Fora composite sample
containing fine sediment, allow grit to settle for 10 - 20 seconds before pouring the sample into the
graduated cylinder.
6. Pour the 25-mL aliquot into the filter funnel, replace the cap, and pull the sample through the filter
using the hand pump. Vacuum pressure from the pump should not exceed 7 inches of Hg to avoid
rupture of fragile algal cells.
NOTE: If 25 mL of sample will not pass through the filter, discard the filter and rinse the chamber
thoroughly with deionized water. Collect a new sample using a smaller volume of sample, measured
to±1 mL. Be sure to record the actual volume sampled on the sample label and the Sample
Collection Form.
1. Remove both plugs from the filtration chamber and pour out the filtered water in the chamber.
Remove the filter funnel from the filter holder. Remove the filter from the holder with clean forceps.
Avoid touching the colored portion of the filter. Fold the filter in half, with the colored sample (filtrate)
side folded in on itself. Place the folded filter in a 50 ml centrifuge tube. Discard filtered water.
8. Complete a periphyton sample label for biomass, including the volume filtered, and attach it to the
centrifuge tube. Cover the label completely with a strip of clear tape. Place the centrifuge tube into a
self-sealing plastic bag.
9. Record the sample ID number of the label and the total volume of the composite index sample on the
form. Record the volume filtered in the "Biomass" field on the Sample Collection Form. Double check
that the volume recorded on the collection form matches the total volume recorded on the sample
label.
10. Place the centrifuge tube containing the filter on dry ice.
7.2.6.4 Acid/Alkaline Phosphatase Activity Sample
Prepare the Acid/Alkaline phosphatase activity (APA) sample from a 50-mL subsample
of the composite index sample. Table 7-9 presents the procedure for preparing APA samples.
No field treatment (i.e., filtration, preservation) of the APA sample is necessary. Complete a
label for the sample and affix it to a 50-mL centrifuge tube. Record the sample ID number, and
the volume of the subsample on the Sample Collection Form (Figure 6.1-3). Check to ensure
that the information recorded on the Sample Collection Form matches the corresponding
information recorded on the sample label. Store APA samples frozen until shipment to the
laboratory.
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Table 7.9. Procedure for preparing acid alkaline phosphatase activity samples for periphyton
1. Prepare a sample label (with a sample number) for the APA sample. Record the volume of the sample
(typically 50 ml) and the volume of the composite index sample on the label. Attach the completed
label to a 50-mL centrifuge tube; avoid covering the volume graduations and markings. Cover the label
completely with a clear tape strip.
2. Rinse a 60-mL syringe with deionized water.
3. Thoroughly mix the bottle containing the composite sample.
4. Withdraw 50 ml of the mixed sample into the syringe. Place the contents of the syringe sample into
the labeled 50-mL centrifuge tube. Cap the tube tightly and seal with plastic electrical tape.
5. Record the sample ID number of the label and the total volume of the composite index sample on the
form.
6. Record the volume of the sample in the centrifuge tube in the "APA Sample" field of the Sample
Collection Form.
7. Freeze the sample immediately and keep frozen until shipping.
7.3 Data Forms and Sample Inspection
After the Site Assessment Form is completed, the Field Team Leader reviews all of the
data forms and sample labels for accuracy, completeness, and legibility. The other team
members inspect all sample containers and package them in preparation for transport, storage,
or shipment. Refer to Appendix C for details on preparing samples for shipping.
Ensure that all required data forms for the site have been completed. Confirm that the
SITE-ID, the visit number, and date of visit are correct on all forms. On each form, verify that all
information has been recorded accurately, the recorded information is legible, and any flags are
explained in the comments section. Ensure that written comments are legible, with no
"shorthand" or abbreviations. Make sure there are no marking s in the scan code boxes. Make
sure the header information is completed on all pages of each form. After reviewing each form
initial the upper right corner of each page of the form.
Ensure that all samples are labeled, all labels are completely filled in, and each label is
covered with clear plastic tape. Compare sample label information with the information recorded
on the corresponding field data forms (e.g., the Sample Collection Form) to ensure accuracy.
Make sure that all sample containers are properly sealed.
7.4 Launch Site Cleanup
Load the boat on the trailer and inspect the boat, motor, and trailer for evidence of
weeds and other macrophytes. Clean the boat, motor, and trailer as completely as possible
before leaving the launch site. Inspect all nets for pieces of macrophyte or other organisms and
remove as much as possible before packing the nets for transport. Pack all equipment and
supplies in the vehicle and trailer for transport. Keep equipment and supplies organized so they
can be inventoried using the equipment and supply checklists presented in Appendix A. Lastly,
be sure to clean up all waste material at the launch site and dispose of or transport it out of the
site if a trash can is not available.
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8.0 FIELD QUALITY CONTROL
Standardized training and data forms provide the foundation to help assure that data
quality standards for field sampling are met. These Standard Operating Procedures for field
sampling and data collection are the primary guidelines for all cooperators and field teams. In
addition, repeat sampling, duplicate sampling, and field evaluation and assistance visits will
address specific aspects of the data quality standards for the National Rivers and Streams
Assessment.
8.1 Repeat and Duplicate Sampling
Repeat and duplicate sampling will provide data to make variance estimates (for
measurement variation and index period variation) that can be used to evaluate the NRSA
design for its potential to estimate status and detect trends in the target population of sites. A
summary of the repeat and duplicate sampling design is provided in Figure 8-1.
Revisits and Field Duplicate Design
First 10% of sites on list
On either Visit 1 or Visit 2,
collect duplicate samples
1
Visit 1
Primary Sample
(P)
water chemistry
Secchi depth
In situ measures
chlorophyll-a
sediment enzymes
periphyton
benthos
enterococci
fish
fish tissue
physical habitat
Space revisits as fa
apart as practical
1
Filter Blank
(F)
Enterococci
Collect on visit where
duplicate samples are
NOT collected
1
Field Duplicate
(D)
fish tissue
1
1
Visit 2
1
1 1
Primary Sample
(P)
water chemistry
Secchi depth
In situ measures
chlorophyll-a
sediment enzymes
periphyton
benthos
enterococci
fish
physical habitat
Field Duplicate
(D)
Water chemistry
Chlorophyll-a
Sediment
enzymes
Periphyton
Benthos
Duplicates = "measurement" variation
Revisits = "measurement" variation + index period variation
Figure 8.1. Summary of the repeat and duplicate sampling design.
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8.1.1 Repeat Sampling
A total of 10% of the target sites visited will be revisited during the same field season by
the same field team that initially sampled the site. Repeated samples and measurements are
taken from the same reach as the first visit. Each state has four repeat sites; the first two
wadeable and the first two non-wadeable sites in their list. If a site selected for repeat sampling
is dropped, then the alternate assigned to replace it should be revisited. If a non-wadeable site
is sampled with wadeable methods, the next non-wadeable site should be selected as the
repeat site. The primary purpose of this "revisit" set of sites is to collect temporal replicate
samples to provide variance estimates for both measurement variation and index period
variation. The revisit will include the full set of indicators and associated parameters. The time
period between the initial and repeat visit to a site should be as long as possible, but not less
than 2 weeks. Fish tissue and PPCP water samples will only be collected on the first visit (see
Section 8.1.2).
8.1.2 Duplicate Sampling
Duplicate samples will be collected for certain indicators from the sites that are revisited.
They will be collected at one of the visits, not both. These duplicate samples will be collected for
water chemistry, chlorophyll a, sediment enzymes, periphyton, benthos, enterococci, and fish
tissue (not for fish community data or physical habitat). These samples and measurements are
taken from the same reach as the primary sample. The samples should be taken by the same
field crew and <2 days later. These spatial replicates will provide measurement variance and
spatial variance estimates. Label the samples as (primary site /D#)-D to indicate that they are
duplicate samples. Duplicates for fish tissue should be taken on the first visit, no fish tissue
needs to be collected during the second visit. Duplicate PPCP water samples should also be
collected during the first visit at the designated urban river sites.
In addition, a filter blank will be collected for enterococci. The teams will filter a small
amount (10 ml_) of sterile buffer through 4 filters, label them and write "blank" on the label and
field form, and package and submit these samples to the lab. The filter blanks should be run
before the sample is filtered. The filter blanks should be collected on the field visit that duplicate
samples are not collected (Figure 8-1). A detailed description of the filter blanks is found in table
7-2.
8.1.3 Taking Field Duplicates
On the visit crew are taking duplicates samples, ensure that there are two site kits for
supplies and materials. If you are taking duplicates on a subsequent field day follow standard
sample procedures for collecting the duplicate samples. If you are collecting duplicates on the
same day as the primary sample follow the modified protocols in this section. Fish tissue, both a
primary and duplicate, is collected on the first visit only.
After you take the first water chemistry sample, rinse the beaker three times with stream
water, replace any torn gloves, and collect a second sample with a new cubitainer following the
procedures in the water chemistry sections. The water chemistry chlorophyll a sample can be
filtered from the same container as the primary sample. If there is not sufficient water for both
filters, process the primary sample, then collect a second water sample from the index site for
the duplicate sample.
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For transect sample duplicates (sediment enzymes, benthic macroinvertebrates, and
periphyton) move 1 meter upstream of the primary sample location. At this new location
upstream of the transect, take a duplicate sample following the same procedures that are used
to collect the primary sample. You do not need to collect a duplicate for the low gradient
samples.
8.2 Field Evaluation and Assistance Visits
A rigorous program of field and laboratory evaluation and assistance visits has been
developed to support the National Rivers and Streams Assessment Program. These evaluation
and assistance visits are explained in detail in the Quality Assurance Project Plan (QAPP) for
the NRSA. The following sections will focus only on the field evaluation and assistance visits.
These visits provide a QA/QC check for the uniform evaluation of the data collection
methods, and an opportunity to conduct procedural reviews as required to minimize data loss
due to improper technique or interpretation of field procedures and guidance. Through uniform
training of field teams and review cycles conducted early in the data collection process,
sampling variability associated with specific implementation or interpretation of the protocols will
be significantly reduced. The field evaluations will be based on the Field Evaluation Plan and
Checklists. This evaluation will be conducted for each unique team collecting and contributing
data under this program (EPA will make a concerted effort to evaluate every team, but will rely
on the data review and validation process to identify unacceptable data that will not be included
in the final database).
8.2.1 Specifications for QC Assurance
Field evaluation and assistance personnel are trained in the specific data collection
methods detailed in this Field Operations Manual. A plan and checklist for field evaluation and
assistance visits have been developed to detail the methods and procedures. The plan and
checklist are included in the QAPP. Table 8-1 summarizes the plan, the checklist, and corrective
action procedures.
Table 8.1. General information noted during field evaluation
Field
Evaluation
Plan
Regional Coordinators will arrange the field evaluation visit with each Field Team,
ideally within the first two weeks of sampling.
The Evaluator will observe the performance of a team through one complete set of
sampling activities.
If the Team misses or incorrectly performs a procedure, the Evaluator will note it on
the checklist and immediately point it out so the mistake can be corrected on the spot.
The Evaluator will review the results of the evaluation with the Field Team before
leaving the site, noting positive practices and problems.
Field
Evaluation
Checklist
The Evaluator observes all pre-sampling activities and verifies that equipment is
properly calibrated and in good working order, and NRSA protocols are followed.
The Evaluator checks the sample containers to verify that they are the correct type
and size, and checks the labels to be sure they are correctly and completely filled out.
The Evaluator confirms that the Field Team has followed NRSA protocols for locating
the site.
The Evaluator observes the complete set of sampling activities, confirming that all
protocols are followed.
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Table 8.1. General information noted during field evaluation
The Evaluator will record responses or concerns, if any, on the Field Evaluation and
Assistance Check List.
If the Evaluator's findings indicate that the Field Team is not performing the
tive
Action
procedures correctly, safely, or thoroughly, the Evaluator must continue working with
corrective jeam untj| certajn of the Team's ability to conduct the sampling properly so
that data quality is not adversely affected.
Procedures
If the Evaluator finds major deficiencies in the Field Team operations the Evaluator
must contact a NRSA QA official.
It is anticipated that evaluation and assistance visits will be conducted with each Field
Team early in the sampling and data collection process, and that corrective actions will be
conducted in real time. If the Field Team misses or incorrectly performs a procedure, the
Evaluator will note this on the checklist and immediately point this out so the mistake can be
corrected on the spot. The role of the Evaluator is to provide additional training and guidance so
that the procedures are being performed consistent with the Field Operations Manual, all data
are recorded correctly, and paperwork is properly completed at the site.
8.2.2 Reporting
When the sampling operation has been completed, the Evaluator will review the results
of the evaluation with the Field Team before leaving the site (if practicable), noting positive
practices and problems (i.e., weaknesses [might affect data quality] or deficiencies [would
adversely affect data quality]). The Evaluator will ensure that the Team understands the findings
and will be able to perform the procedures properly in the future. The Evaluator will record
responses or concerns, if any, on the Field Evaluation and Assistance Check List. After the
Evaluator completes the Field Evaluation and Assistance Check List, including a brief summary
of findings, all Field Team members must read and sign off on the evaluation.
If the Evaluator's findings indicate that the Field Team is not performing the procedures
correctly, safely, or thoroughly, the Evaluator must continue working with this Field Team until
certain of the Team's ability to conduct the sampling properly so that data quality is not
adversely affected. If the Evaluator finds major deficiencies in the Field Team operations (e.g.,
major misinterpretation of protocols, equipment or performance problems) the Evaluator must
contact the following QA official:
• Sara/? Lehmann, EPA National Rivers and Streams Assessment Project QA Officer
The QA official will contact the Project Manager to determine the appropriate course of
action. Data records from sampling sites previously visited by this Field Team will be checked to
determine whether any sampling sites must be redone.
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9.0 LITERATURE CITED
Allen-Gil, S., M. Green, and D. H. Landers. 19XX. Fish abundance, instream habitat and the
effects of historical land use practices in two large alluvial rivers on the Olympic
Peninsula, Washington. U.S. EPA, WED. In review.
American Red Cross. 1979. Standard First Aid and Personal Safety. American National
Red Cross. 269 pp.
Arend, K.K. 1999. Macrohabitat identification. Pages 75-93 in M.B. Bain and N.J. Stevenson
(editors). Aquatic habitat assessment: common methods. American Fisheries Society,
Bethesda, Maryland.
Bain, M.B., J.T., Finn, and H.E. Brooke. 1985. Quantifying stream substrate for habitat analysis
studies. North American Journal of Fisheries Management 5:499-506.
Bain, M.B., and N.J. Stevenson (editors). 1999. Aquatic habitat assessment: common methods.
American Fisheries Society, Bethesda, Maryland.
Baker, J.R., D.V. Peck, and D.W. Sulton (editors). 1997. Environmental Monitoring and
Assessment Program Surface Waters Field Operations Manual for Lakes. EPA/620/R-
97/001. U.S. Environmental Protection Agency, Washington DC.
Barbour, M.T., J. Gerritsen, B.D. Snyder, and J.B. Stribling. 1999. Rapid Bioassessment
Protocols for Use in Streams and Wadeable Rivers: Periphyton, Benthic
Macroinvertebrates, and Fish, Second Edition. EPA 841-B-99-002. U.S. Environmental
Protection Agency, Office of Water, Washington D.C.
Bisson, P.A., J.L. Nielsen, R.A. Palmason, and L.E. Grove. 1982. A system of naming habitat
types in small streams, with examples of habitat utilization by salmonids during low
streamflow. In N.B. Armantrout [ed.] Acquisition and Utilization of Aquatic Habitat
Inventory Information: Proceedings of the Symposium. [Portland, OR, October 1981].
Dietrich, W.E., Kirchner, J.W., Ikeda, H., Iseya, F. 1989. Sediment supply and the development
of the coarse surface layer in gravel-bedded rivers. Nature 340: 215-217.
Dunne, T., and L.B. Leopold. 1978. Water in environmental planning. W.H. Freeman, New York.
818 p.
Flotemersch, J.E.1, B.C. Autrey2, and S.M. Cormierl (editors). 2000. Logistics of Ecological
Sampling on Large Rivers. U.S. Environmental Protection Agency, Cincinnati OH.
Flotemersch, J.E., J.B.Stribling, and M.J. Paul. 2006. Concepts and Approaches for the
Bioassessment of Non-wadeable Streams and Rivers. EPA 600-R-06-127. U.S.
Environmental Protection Agency, Cincinnati, OH.
Frissell, C.A.; W.J. Liss, W.J.; Warren, C.E.; Hurley, M.C. 1986. A hierarchical framework for
stream habitat classification: viewing streams in a watershed context. Environmental
Management 10: 199-214.
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National Rivers and Streams Assessment Final Manual
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Page 214
Harrelson, C.C., C.L. Rawlins, and J.P. Potyondy. 1994. Stream channel reference sites: an
illustrated guide to field technique. USDA Forest Service, General Technical Report RM-
245, Rocky Mountain Forest and Range Experiment Station, Fort Collins, Colorado. 61 p.
Hawkins, C.P., J.L Kershner, P.A. Bisson, M.D. Bryant, LM. Decker, S.V. Gregory, D.A.
McCullough, C.K. Overton, G.H. Reeves, R.J. Steedman, and M.K. Young. 1993. A
hierarchical approach to classifying stream habitat features. Fisheries 18:3-12.
Helm, W.T. 1985. Aquatic habitat inventory: standard methods and glossary. American
Fisheries Society, Western Division, Bethesda, Maryland.
Kaufmann, P.R. (ed.), 1993, Physical Habitat, pp. 59-69, in R.M. Hughes (ed), Stream Indicator
and Design Workshop, EPA/600/R-93/138, U.S. Environmental Protection Agency,
Office of Research and Development, Corvallis, Oregon.
Kaufmann, P.R. and T.R. Whittier. 1997. Habitat Assessment. Pages 5-1 to 5-26 in J.R. Baker,
D.V. Peck, and D.W. Sutton editors. Environmental Monitoring and Assessment
Program -Surface Waters: Field Operations Manual for Lakes. EPA/620/R-97/001. U.S.
Environmental Protection Agency, Washington, D.C.
Kaufmann, P.R., P. Levine, E.G. Robinson, C. Seeliger, and D. Peck. 1999. Quantifying
Physical Habitat in Wadeable Streams. EPA/620/R-99/003. U.S. Environmental
Protection Agency, Washington, D.C.
Klemm, D. J., P. A. Lewis, F. Fulk, and J. M. Lazorchak. 1990. Macroinvertebrate Field and
Laboratory Methods for Evaluating the Biological Integrity of Surface Waters. EPA
600/4-90/030. U.S. Environmental Protection Agency, Cincinnati, Ohio.
Lemmon, P.E. 1957. A new instrument for measuring forest overstory density. Journal of
Forestry 55:667-669.
Leopold, L.B. 1994. A view of the river. Harvard University Press, Cambridge, Massachusetts.
298 p.
Linsley, R.K., M.A. Kohler, and J.L.H. Paulhus. 1982. Hydrology for engineers. McGraw-Hill
Book Co. New York, NY. 508 p.
Moore, K.M., K.K. Jones, and J.M. Dambacher. 1993. Methods for stream habitat surveys:
Oregon Department of Fish and Wildlife, Aquatic Inventory Project. Version 3.1. Oregon
Department of Fish and Wildlife, Corvallis, OR 34 pp.
Mulvey, M., L. Caton, and R. Hafele. 1992. Oregon nonpoint source monitoring protocols:
stream bioassessment field manual for macroinvertebrates and habitat assessment.
Oregon Department of Environmental Quality, Laboratory Biomonitoring Section.
Portland, Oregon.
National Institute for Occupational Safety and Health. 1981. Occupational Health Guidelines
for Chemical Hazards (Two Volumes). NIOSH/OSHA Publication No. 81-123.
U.S. Government Printing Office, Washington, D.C.
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Page 215
Nelson, J.S., E.J. Grossman, H. Espinosa-Perez, L.T. Findley, C.R. Gilbert, R.N. Lea, and J.D.
Williams. 2004. Common and Scientific Names of Fishes from the United States,
Canada, and Mexico. American Fisheries Society, Special Publication 29, Bethesda,
Maryland.
Ohio EPA. 1990. Ohio EPA Fish Evaluation Group Safety Manual. Ohio Environmental
Protection Agency, Ecological Assessment Section, Division of Water Quality Planning
and Assessment, Columbus, Ohio.
Occupational Safety & Health Administration (OSHA). 2006. Regulations (Standards - 29 CFR).
Substance technical guidelines for formalin - 1910.1048 App A. Occupational Safety &
Health Administration. Washington, DC 20210.
Peck, D. V., Herlihy, AT., Hill, B.H., Hughes, R.M., Kaufmann, P.R., Klemm, D.J., Lazorchak,
J.M., McCormick, F.H., Peterson, S.A., Ringold, P.L, Magee, T., Cappaert, M., 2006.
Environmental Monitoring and Assessment Program-Surface Waters Western Pilot
Study: Field Operations Manual for Wadeable Streams. EPA/620/R-06/003. U.S.
Environmental Protection Agency, Office of Research and Development, Washington,
DC.
Plafkin, J.L., Barbour, M.T., Porter, K.D., Gross, S.K., Hughes, R.M. (1989). "Rapid
bioassessment protocols for use in streams and rivers: Benthic macroinvertebrates and
fish," EPA/440/489/001, U.S. Environmental Protection Agency, Assessment and
Watershed Protection Division, Washington, DC.
Platts, W.S., Megahan, W.F., and Minshall W.G. (1983). "Methods for evaluating stream,
riparian, and biotic conditions," General Technical Report INT-138, USDA Forest
Service, Rocky Mountain Research Station, Ogden, UT.
Rankin, E.T. (1995) The qualitative habitat evaluation index (QHEI) in W.S. Davis and T.
Simons (eds.). Biological Assessment Criteria; Tools for Risk-based Planning and
Decision Making. CRC Press/Lewis Publishers, Ann Arbor Ml.
Robison, E.G. and R.L. Beschta. 1990. Characteristics of coarse woody debris for several
coastal streams of southeast Alaska, USA. 47(9): 1684-1693.
Robison, E.G. and P.R. Kaufmann. 1994. Evaluating two objective techniques to define pools in
small streams, pgs 659-668, |ri R.A. Marston and V.A. Hasfurther (eds.) Effects of
Human Induced changes on hydrologic systems. Summer Symposium proceedings,
American Water Resources Association,. June 26-29, 1994, Jackson Hole, Wyoming.
1182pp.
Stack, B.R. 1989. Factors influencing pool morphology in Oregon coastal streams. M.S. Thesis,
Oregon State University. 109 p.
Standard Methods for the Examination of Water and Wastewater, Method 10300 C,D. 20th Ed.
1998. American Public Health Association, Washington, D.C.
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National Rivers and Streams Assessment Final Manual
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Page 216
Stauffer, Dr. Jay R., J. Karish and T.D. Stecko. 2001. Guidelines for Using Digital Photos as
Fish Vouchers for Pennsylvania Fishes. The Pennsylvania State University and National
Park Service.
Steinman, A.D. and G. A. Lamberti. 1996. Biomass and pigments of benthic algae. P. 297. In
"Methods in Stream Ecology". Hauer, F.R. and G.A. Lamberti (eds). Academic Press,
San Diego, CA.
Stevens, D. L, Jr., and A. R. Olsen. 2004. Spatially-balanced sampling of natural resources in
the presence of frame imperfections. Journal of American Statistical Association:99:262-
278.
Strahler, A.M. 1957. Quantitative Analysis of Watershed Geomorphology. Trans. Am. Geophys.
Un. 38,913-920.
U.S. Coast Guard. 1987. Federal Requirements for Recreational Boats. U.S. Department
of Transportation, United States Coast Guard, Washington, D.C.
USDA Forest Service, 1995. A guide to field identification ofbankfull stage in the western United
States. Rocky Mountain Forest and Range Experiment Station, Stream Systems
Technology Center, Fort Collins, Colorado (31 minute video, closed captioned).
USDA Forest Service, 2002. Identifying bankfull stage in forested streams in the eastern United
States. Rocky Mountain Forest and Range Experiment Station, Stream Systems
Technology Center, Fort Collins, Colorado (46 minute video, closed captioned).
USEPA. 2000a. EPA Quality Manual for Environmental Programs 5360A1. May 2000.
http://www.epa.gov/quality/qs-docs/5360.pdf
USEPA. 2000b. EPA Order 5360.1 A2 CHG2, Policy and Program Requirements for Mandatory
Agency-wide Quality System, May 5, 2000. http://www.epa.gov/qualitv/qs-docs/5360-
l.pdf
U.S. EPA (Environmental Protection Agency). 2004. Wadeable Streams Assessment: Field
Operations Manual. EPA 841-B-04-004. U.S. Environmental Protection Agency, Office of
Water and Office of Research and Development, Washington, D.C.
USEPA. 1986. Occupational Health and Safety Manual. Office of Planning and Management
U.S. Environmental Protection Agency, Washington, D.C.
Walsh, S. J. and M. R. Meador. 1998. Guidelines for quality assurance and quality control offish
taxonomic data collected as part of the National Water-Quality Assessment Program.
U.S. Geological Survey, Water-Resources Investigations Report 98-4239, Raleigh, North
Carolina.
Wilcock, P.R. 1988. Sediment maintenance flows: Feasibility and basis for prescription, in
Gravel-Bed Rivers in the Environment, edited by P.C. Klingeman et al., pp. 609-632,
Water Resour. Publ., Highlands Ranch, Colo., 1998.
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Wilcock, P.R. 1988. Two-fraction model of initial sediment motion in gravel-bed rivers. Science
280:410-412.
Wolman, M.G. (1954). "A method of sampling coarse riverbed materials," Transactions of the
American Geophysical Union 35(6), 951-956.
Web Pages:
US EPA Aquatic Monitoring Research: http://www.epa.gov/nheerl/arm
NHD Plus: http://www.horizon-systems.com/nhdplus
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APPENDIX A
List of Equipment and
Supplies
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EQUIPMENT & SUPPLY LISTS
General
Field Operations Manual and/or laminated
Quick Reference Guide
Laminated invasive species guide
Covered clipboards
Filed forms and sample labels
Clear tape strips for covering labels
Pencils (#2)
Fine-tipped indelible markers
Digital camera with extra memory card &
battery
Maps and access instructions
Sampling permits and/or permission letters
GPS unit with manual and reference card
50 m or 100 m measuring tape with reel
Surveyor's flagging tape
Equipment
• Laser rangefinder (400 ft. distance range)
and clear waterproof bag
• Batteries
• 1%- 10% Bleach
• Barometer or elevation chart to use for
calibration
• Calibration cups and standards for multi-
probe unit
• Electrical tape
• Scissors
• Plastic storage tub
• Cell phone, 2-way radios, and/or walkie-
talkies
• 2 pair chest waders
• 1 or 2 fisherman's vest with lots of pockets
and snap fittings.
Boat Equipment List
Motor
Gas Can
Lifejackets (1/person)
Type IV PFD (Throwable Life Saving device)
Bow/Stern lights
Anchor with 75m line or sufficient to anchor in 50m
depth
Float to attach to anchor
Sonar Unit
Oars or Paddles
First Aid Kit
Extra Boat Plug
Spare Prop Shear Pin
Emergency Tool kit
Hand Bilge pump
Fire Extinguisher
Boat horn
Spare prop
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Sample/Data Collection
Multi-parameter water quality meter with pH,
DO, temperature, and conductivity probes
20 cm diameter Secchi disk and calibrated
sounding line, marked in 0.5 m intervals
3 L Nalgene beaker
1-2L Amber Nalgene bottle
Tape measure (in centimeters)
Nitrile gloves
Calibrated PVC sounding rod, 3-m length,
marked in 0.1 m increments
Convex spherical canopy densiometer
(Lemmon Model B), modified with taped "V"
Clinometer
Bearing compass (Backpacking type)
Binoculars
Surveyor's telescoping leveling rod (round
profile, metric scale, 7.5m extended)
Meter stick for bank angle measurements
Current velocity meter, probe, and operating
manual
Top-set wading rod for use with current
velocity meter
Neutrally buoyant object (e.g., plastic golf ball
with holes, small rubber ball, stick)
Portable Weir with 60° "V" notch (optional) and
plastic sheeting to use with weir
Plastic bucket (or similar container) with
volume graduations
Petite Ponar sampler with plastic tub, drop
line, and spare pinch pin. (Standard Ponar
may substitute)
60-mL plastic syringe with 3/8" hole bored into
the end
Large stainless steel spoon for mixing
sediment composite
Large Funnel (15-20 cm diameter)
12-cm2 area delimiter (3.8 cm diameter
pipe, 3 cm tall)
Stiff-bristle toothbrush with handle bent
at 90° angle
Modified kick net (D-frame, 500 urn
mesh, 4-ft handle)
Sieve-bucket, 500 urn mesh (U.S. std
No. 35)
Watch with timer or stopwatch
Watchmakers' forceps
Buckets, plastic, 8- to 10-qt capacity
Plastic electrical tape
Electrofishing equipment (boat, barge,
and/or backpack units, including
variable voltage pulsator unit, wiring
cables, generator, electrodes, dip
nets, and all safety equipment)
Linesman gloves
Livewell and/or buckets
2 Non-conducting dip nets with 1/4"
mesh
1 Minnow net for dipping small fish from
live well
Measuring board (millimeter scale)
Pre-sterilized, 250 ml sample bottle
Sodium thiosulfate tablet
500-mL plastic bottles for the periphyton
composite sample
25-mL or 50-mL graduated cylinder
1-L wash bottle for stream water
1-L wash bottle containing deionized
water
Coolers
Sample Processing/Preservation
• Whatman 47 mm 1.2 micron GF/C glass
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Wet ice
Dry ice
95% ethanol
10% buffered formalin
Lugol's solution
Sterile filtration unit (Nalgene 145/147),
including filter funnel, cap, filter holder,
and receiving chamber
Vacuum hand pump and clear plastic tubing
Sterile disposable forceps
Whatman 47 mm polycarbonate 0.4 micron
filters
Whatman 47 mm 0.7 micron GF/F glass
fiber filters
fiber filters
60 x 15 disposable Petri dishes
Phosphate buffered saline solution
Aluminum foil squares (3" x 6")
Dl water
Small spatula, spoon, or scoop to transfer
sample
Aluminum foil (solvent-rinsed and baked)
Heavy-duty food grade polyethylene tubing
Large plastic (composite) bags
Knife or scissors
Plastic cable ties
Scalpel for slitting open large fish before
preservation
Sample Storage
One 4-L cube container
Three 1-L Nalgene bottles
Several Leak-proof HOPE jars for fish
voucher specimens (various sizes from
250 mL - 4L)
500-mL plastic bottle for sediment sample
Sample jars, 1-L HOPE plastic suitable for
use with ethanol (benthic samples)
50-mL screw-top centrifuge tube
sterile microcentrifuge tubes containing
sterile glass beads
Coolers
Packaging/Shipping
Coolers
Cooler liners (30-gal garbage bags)
Dry ice (-60 Ibs per site)
Wet ice (-50 Ibs per site; additional for
shipping)
1-gallon self-sealing bags
Packing/strapping tape
FedEx airbills
Class 9 Dangerous Goods label (for dry ice
shipments)
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A site kit will be provided to the field crews for each sampling site. Site kits will be shipped out
based on the schedule that each field crew provides prior to the start of the sampling season.
Field crew leaders MUST provide a schedule in order to receive the site kits. If your
schedule changes, please report the change as soon as possible to the Field Logistics
Coordinator (Jennifer Pitt; 410-356-8993). Prior to sampling, inspect each site kit to ensure all
supplies are included.
Supplies provided in each Site Kit:
• Field Data Forms
• Sample Labels
• National Rivers and Streams Assessment Fact Sheets
• 1 4-L cube container
• 1 1-L Nalgene bottle
• 500-mL plastic bottle for sediment sample
• 1 sterile 250 mL fecal indicator bottle
• 1 Zip tie
• 2 1-L HOPE plastic sample jars suitable for use with ethanol (benthic samples)
• 5 50-mL screw-top centrifuge tubes (4 for periphyton, 1 for measuring enterococci
sample for filtering and then for storing the chlorophyll a filter)
• 4 sterile microcentrifuge tubes containing sterile glass beads
• Funnel analytical test filter 250 mL
• Sterile disposable forceps (2)
• Sterile phosphate buffered saline (PBS)
• Large Plastic Bags
• Foam envelope
• FedEx airbills for all labs
• Dry ice box will be included in approximately every 4th site kit
• Dry ice shipping label
Supplies Provided in Each Fish Tissue Sampling Kit:
• Aluminum foil (solvent-rinsed and baked)
• Heavy-duty food grade polyethylene tubing
• Large plastic (composite) bags
• Plastic cable ties
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Supplies Provided in Each Base Kit:
• Nitrile Gloves
• Clinometer
• Spherical Densiometer
• Bottle of 50 Sodium Thiosulfate Tablets
• Aluminum foil 3x6"
• 15" stainless steel spoon
• (2) D-frame Kick Net - 500 urn mesh, 52" handle
• (2) Sieve bucket - 500 urn
• Weighted Secchi disk
• Rectangular fiberglass surveying rod - metric
• CST Berger SAL 20 Automatic Level
• Level tripod
• (2) 1 Liter Nalgene wash bottles
• 3 gallon Rubbermaid Roughneck tote
• Graduated cylinder 250 mL
• 2 Liter amber Nalgene rectangular bottle
• 500-mL plastic bottle for periphyton sample collection
• Nalgene filtering flask
• #8 silicone stopper
• Filter funnel adapter
• Whatman 47 mm polycarbonate 0.4 u filters
• Whatman 47 mm glass fiber GF/F 0.7 u filters
• Whatman 47 mm glass fiber GF/C 1.2 u filters
• Disposable petri dishes 60x15
• 3 Liter Nalgene beaker
• Utility funnel 15cm diameter
• Centrifuge tube stand
• Hand vacuum pump
• 500 mL Lugol's solution
• 4 Liters of QC check solution
• Tape dispenser
• Tape strips
• 1A gallon bucket
• 60 cc syringe with 3/8" hole and tubing
• 12 cm2 area delimiter
• (2) 2 mL pipet and pipet bulb
• Toothbrush bent to 90°
• 24 ct of 1 Liter Nalgene bottles
Note: Lugol's solution, calibration QC check solution, filters, 1 Liter Nalgene bottles, aluminum
foil squares, and disposable nitrile gloves will be provided in the base kit; you may order more
throughout the field season if needed.
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APPENDIX B
Field Forms
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BOATABLE
FORMS
PACKET
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STREAM VERIFICATION FORM - WADEABLE/BOATABLE (Front)
SITE NAME:
SITE ID: FW08
DATE: / / 2 0
VISIT: O1 O2 O 3
_ , , -. Don't forget to record
State of Site Location. Reacn Le*qth on back TEAM:
STREAM/RIVER VERIFICATION INFORMATION
Stream/River Verified by (fill in all that apply) Q GPS O Local Contact O Signs O Roads O Topo. Map
O other (Describe Here): O Not Verified (Explain in Comments)
Coordinates Latitude North
Degrees, Minutes.
and Seconds
MAP OR " " J
Decimal Degrees
Degrees, Minutes,
and Seconds
GPS i • ' • • •
OR
Decimal Degrees
4t
Longitude West Sg(e
O
O
of Are GPS Coordinates
lites w/j 10 Sec. of map?
<3 O Yes
O No
>4
GPS Datum Used
(e.g. NAD27):
DID YOU SAMPLE THIS SITE?
QYES If YES, check one below
SAMPLEABLE (Choose method used)
O Wadeable - Continuous water, greater than 50% wadea
O Boatable
O Partial - Sampled by wading (>50% of reach sampled).
O Partial - Sampled by boat (>50% of reach sampled). Ex
O Wadeable Interrupted - Not continuous water along re.
O Boatable Interrupted - Not continuous water along re-'
O Altered - Stream/River Channel Present but differ froi
V
GENERAL COMMENTS:
O ^' -> If N°> check one below
NON-,-A> .PLEABLE-PERMANENT
ble C Iry- Vis,,ed
C Di, ,-lot visited
O Wetland (No Definable Channel)
Explain belov. O Map Error - No evidence channel/waterbody ever present
Dlain below. % O Impounded (Underneath Lake or Pond)
ft k O Other (explain in comments)
h NON-SAMPLEABLE-TEMPORARY
O Not boatable - Need a different crew - Reschedule for this year
i M. o O Not wadeable - Need a different crew - Reschedule for this year
O Other (Explain in comments)
NO ACCESS
O Access Permission Denied
O Permanently Inaccessible (Unable/Unsafe to Reach Site)
O Temporarily Inaccessible-Fire, etc. - Reschedule for next year
DIRECTIONS TO STREAM/RIVER SITE:
Record information used to define length of reach, and sketch general features of reach on reverse side.
04/07/2009 NRSA Stream Verification 2009
36530
I tall
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STREAM VERIFICATION FORM - WADEABLE/BOATABLE (Back) Re^
Reviewed by
SITE NAME:
DATE:
2 0
VISIT: O1 O2 O 3
SITE ID:
FW08
TEAM:
STREAM/RIVER REACH DETERMINATION
Channel Width
Used to Define
Reach (m)
DISTANCE (m) FROM X-SITE
Upstream
Length
Downstream
Length
Total Reach
Length Intended
(m)
Comment
SKETCH MAP - Arrow Indicates North; Mark site L=Launch X=lndex T= Take Out
NOTE: If an outline map is attached here, use a continuous strip of clear tape across the top edge.
You can also attach a separate sheet with the outline map on it.
For boatable sites you can attach topo map with reach, X-site and transect locations marked.
PERSONNEL
NAME
Bio/Chem
Sampling
O
O
O
O
O
Habitat
O
O
O
O
O
Forms
Review
O
O
O
O
O
a; 530
04/07/2009 NRSA Stream Verification 2009
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FIELD MEASUREMENT FORM - BOATABLE
Reviewed
by (initial):.
SITE ID: FW08
DATE:
/ / 2 o
CALIBRATION INFORMATION
Instrument manufacturer and model:
Instrument ID number:
Operator:
TEMPERATURE
Thermometer Sensor Reading Ft,n
Reading CO (t) ' '
Comments
Elevation
OR
DO
Barometric
Pressure
(mm Hg)
On
_,O m
Calibration
Value
O mg/L
Displayed
Value
O mg/L
Flag
Cal. STD 1 Description
Cal. STD 1 Value
Cal. STD 2 Description
Cal. STD 2 Value
pH
Calibration Verified with Quality Control Sample (QCS)
QCS Description QCS True QCS Measured Flag
T
Cal. STD 1 Description
Cal. STD 1 Value
r .i. STD 2 Description
Cal. STD 2 Value
A
CONDUCTIVITY
Calibration Verif! H w ' i Quality Control Sample (QCS)
QCS Description QCS True <.< QCS Measured g'™ Fiag
Flag
Comments
FIELD MEASURMENTS
TRANSECT:
Time of Day
(HH:MM)
DO(mg/L)XX.X
Temp. (
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SAMPLE COLLECTION FORM - BOATABLE (Front)
Reviewed by
(Initials):
SITE ID: FW08
DATE: / / 2 0
WATER CHEMISTRY (4-L CUBITAINER) No Sample Collected O
Sample ID
Sample
Category *
OP
OD
OP
OD
Chilled
O
O
Comments
WATER COLUMN CHLOROPHYLL (Target Volume = 1000 mL; maxvol =2000 mL) No Sample Collected Q
Sample ID
Sample
Category*
OP
OP
OD
Volume
Filtered
(mL)
Frozen
O
O
WATER CHEMISTRY
Sample ID
Sample
Category
OP
OD
OP
OD
Chilled
0
0
Comments
PPCP (AMBER GLf-,5 CUBITAINER) No Sample Collected Q
^ Cor nents
TRANSECT
Location (L/R):
Dominant
Habitat:
Secondary
Habitat:
IONEPERTR4NSECTI
Substrate:
LONE PERTRSMSECTl
Channel:
IC'ME PER TRANSECT.
T,
A
OL OR
Oc OL
OF O M
O OT
O c O L
OF OM
OOT
OF Oc
OG OOT
OP ORA
0 Rl 0 GL
O OT
B
O L OR
O c OL
OF OM
O OT
O c OL
OF O M
OOT
O F Oc
OG OOT
O P O RA
O Rl O GL
O OT
c
O L OR O
D
' R
O c OL J c C L
O F O • O CM
O OT O o-
Oc OL 0
O F OM O F
OOT Oo
O F O c O
O G O OT O
OL
OM
T
F O C
G O OT
OP O RA O P O RA
O Rl O GL O Rl O GL
O OT O OT
Habitat:
C = Coarse Substrate / LWD L = Leaf Pack
F = Organic Fine Muds / Sand M = Macrophyte beds
OT = Other (Explain in comment section below)
Sample ID
Sample
Category *
OP
OD
OP
OD
No. Jars
1 1
^'SE-T BENTHOS No Sample Collected O
E F G H 1 J K
OL OROL OROL OROL OROL OROL OR OL OR
Oc OLQC OLQC OLQC OLQC OL Oc OL Oc OL
OF O M O F O M O F O '•' O F O M O F O M O F OM OF O M
O OT O OT O OT O OT O OT O OT O OT
Oc OLOC OLQC OLQC O L O c OL Oc OL Oc OL
OF O M O F O M O F O '•' O F O M O F O M O F OM OF OM
O OT O OT O OT O OT O OT O OT O OT
OF Oc OF Oc OF Oc OF Oc OF Oc OF Oc OF Oc
OG OOTQG OOTQG OOTOG OOTQG O OT O G OOT QG OOT
OP ORAQP ORAQP ORAOP o RA o p ORA OP ORA OP ORA
O Rl O GL O Rl O GL O Rl O GL O Rl O GL O Rl O GL O Rl O GL Q Rl O GL
O OT O OT O OT O OT O OT Q OT Q OT
Substrate: Channel:
F = Fine / Sand G = Gravel P = Pool GL = Glide
C = Coarse substrate OT = Other (Explain in Rl = Riffle RA = Rapid
comment section below) OT = Other (Explain n comment section below)
Pre-
served
O
O
Comments
Flag codes: K = No measurement or observation made; U = Suspect measurement or observation; F1, F2, etc. = misc.
flags assigned by field crew. Explain all flags in comment sections.
*Sample Categories: P = Primary, D = Field Duplicate
04/07/2009 NRSA Sample Collection Beatable 2009
3284
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-10
SAMPLE COLLECTION FORM - BOATABLE - (Back)
Reviewed by
SITE ID: FW08
DATE:
./. . ./.2.0
COMPOSITE PERIPHYTON SAMPLE - Primary No Sample Collected O
Sample ID
Assemblage ID .1)
(50-mLtube)
Sample Vol. (mL) Flag Preserved
O
Sample
Category l
OP
OD
Composite Volume (mL)
Chlorophyll (.2)
(GF/F Filter)
Sample Vol. (ir
L) Flag
Frozen
Number of transects sampled (1-11):
Biomass (.3)
(GF/C Filter)
Sample Vol. (mL)
Flag
COMPOSITE PERIPHYTON SAMPLE
Sample ID
Assemblage ID (.1)
(50-mL tube)
Sample Vol. (mL) Flag Preserved
O
Sample
Category '
OP
OD
Composite Volume (mL)
Chlorophyll (.2)
(GF/F Filter)
Sample Vol. (ir
Flag
L) Flag
Frozen
Frozen
Flag
APA (.4)
(50-mL tube)
Sample Vol. (mL)
Flag
Frozen
O
No Sample Collected O
Number of transects sampled (1-11):
Biomass (.3)
(GF/C Filter)
Sample Vol. (mL)
Flag
Comments
Frozen
Flag
APA (.4)
(50-mLtube)
Sample Vol. (mL)
Flag
Frozen
O
SEDIMi ;NT
Sample ID
Sample
Category '
OP
OD
OP
OD
Composite Volump
" -^f
Iran ecis
'HEMISTRY / ENZYMES No Sample Collected Q
Chilled
O
O
Comments
ENTEROCOCCI (Target Volume = 250 mL) No Sample Collected O
Sample ID
One unique ID per line
Flag
Sample
Cate-
gory'
OP
OD
OP
OD
OP
OD
OF
Time
Collected
(hhmm)
Depth
Collected
(m)
Sample
Volume
(mL)
Filt. Start
Time
(hhmm)
Volume Filtered
(Target = 50 mL) **
Filt. 1
Filt. 2
Filt. 3
Filt. 4
Filt. End
Time
(hhmm)
Time
Frozen
(hhmm)
Flag
Comments
" Sample Categories: P = Primary: D = Field Duplicate: F = Filter Blank (Enterococci sample only) Filter blank is collected at vis.it '.vhei« field duplicate sample is NOT taken.
" If <25 ml of buffer solution was used to rinse filter, indicate with an F flag and note in comment section which filter(s) were affected along with the approximate volume(s) of
buffer solution used, 3284
• r^"
04/07.;2009 NRSA Sample Collection Beatable 2009
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-11
PHAB: CHANNEL/RIPARIAN TRANSECT FORM - BOATABLE (FRONT)
Rev'd by(lnlt):
Arrival Time
Leave Time
SITE ID: FW08
DATE:
/ 2,0
TRANSECT: OA OB OC OD OE OF OG OH Ol OJ OK OX
GPS Latitude -dd mm ss.s GPS Longitude - ddd mm ss.s
Chosen bank side:
(Facing down stream) O Left O Right
Transect
Midstream
Transect
Bank
"LITTORAL" SUBSTRATE INFORMATION
DEPTH O ft O m
SHORE
DOM SEC
RS
RR
XB
RS
RR
XB
BOTTOM
DOM SEC
RS
RR
XB
RS
RR
XB
CLASS
BOTTOM SUBSTRATE FROM (X ONE):
O Judgement -or- O OBS. @ 5 Littoral Depths
Flag
RS = Bedrock (Smooth) - (Larger than a car)
RR = Bedrock ( Rough) - (Larger than a car)
XB = Large Boulder (1000 to 4000 mm) - (Meterstick to car)
FLAG
SB
SB
SB
SB
SB = Small Boulder (250 to 1000 mm) - (Basketball to Meterstick)
CB
CB
CB
CB
CB = Cobble (64 to 250 mm) - (Tennis ball to Basketball)
GC
GC
GC
GC
GC = Coarse Gravel (16 to 64 mrn) - (Marble to Tennis ball)
GF
GF
GF
GF
GF = Fine Gravel (2 to 16 mm) - (Ladybug to marble)
SA
SA
SA
SA
SA = Sand (0.06 to 2 mm) - (Gritty • up to Ladybug size)
BANK CHARACTERISTICS
FN
FN
FN
FN
FN = Silt / Clay / Muck - (Not Gritty)
X.XX (m) FLAG
HP
HP
HP
HP
HP = Hardpan - (Firm, Consolidated Fine Substrate)
Wetted Width
WD
WD
WD
WD
WD = Wood - (Any Size)
OT OT OT OT OT = Other (Write comment below)
Bar Width
LARGE WOODY DEBRIS doxiom pioi> TALLy EACH PIECE Flag
CHEC ' IFUNMA. ' cD
Ai ~ZER<-
Bankfull Width
DIAMETER
LARGE END
0.3 -0.6 m
0.6-0.8 m
0.8-1.0m
> 1.0 m
Wood All'Part in Wetted Channel
LENGTH 5-15111
Dry but All/Par* .1 Ban till C. .me!
Bankfull Height
LENGTH 5-15m
Incised Height
BANK
ANGLES
SLOPE/BEARING/DISTANCE (Optional): Deleirnlne slope If feasible In terms of time and distances. Record GPS
coordinates If practical.
INTENDED transect ACTUAL transect
spacing xxx (m): spacing xxx (m):
Slope and Bearing not determined (use map) O
Backslte
Slope Bearing Distance Way
XX.X % o. 359 (m) Point #
GPS Latitude - dd mm ss.s
GPS Longitude • ddd mm ss.s
Flag
Flag
Comments
Flag Codes: K = no measurement made: U = suspect measurement: Ft. F2. etc. = flags assigned by each field crew. Explain all flags in comments
section on tnis side or on Side 2 of tills form.
04/07/2009 NRSA Channel Riparian Boatable Front
24732
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-12
PHAB: CHANNEL/RIPARIAN TRANSECT FORM - BOATABLE (Back)
ReVdby(lnlt.):
SITE ID: FW08 DATE:
TRANSECT: O A OB OC OD OE OF OG OH Ol OJ OK
1 1
OX
/ 2 0
Chosen bank side:
(Facing down stream*
0 Left O Right
0 = Absent (0%) D = Deciduous
VISUAL RIPARIAN 1 = Spare <<10%) c=Coniferous
VKpwni. mri 2 = Moderate (10-40%) E = Broadleaf Evergreen
ESTIMATES 3= Heavy (40-75%) M = Mixecl
4 =Very Heavy (>75«i) N = None
RIPARIAN
VEGETATION COVER
(10m x 20m Plot)
Woody Vegetation Type
BIG Trees {Trunk
>0.3mDBH)
SMALL Tress (Trunk
<0.3 m DBH)
Woody Vegetation Type
Woody Shrubs &
Saplings
Non-Woody Herbs,
Grasses. & Forbs
Woody Shrubs
& Saplings
Non-Woody Herbs.
Grasses and Forbs
Barren. Bare Dirt
or Duff
HUMAN INFLUENCE
Wall/Dike/Revetment
/Riprap/Dam
Buildings
PavemenUCIeared Lot
Road/Railroad
Pipes (Inlel/Outlet)
Landfill/Trash
Park/Lawn
Row Crops
Pasture/RangelHay Field
Logging Operations
Mining Activity
Left Bank
Canopy
D C E M N
01234
01234
Understory
0 C E M N
01234
01234
Ground C
01234
01234
01234
0 -Not Present P = >10 m
Left Bank
0 P C B
0 P C B
0 P C B
0 P C B
Right Bank
>5 m high)
D C E M N
01234
01234
(0.5 to 5m high)
D C E M N
01234
01234
jver (<0.5 m high)
01234
01234
01234
Flag
C = Within 10m B - On Bank
Right Bank
OPCB
0 P C ."
0 P C ..
" P C B
OPCB | 0 ^ C B
0 P C B ^
OPCB
OPCB
OPCB
OPCB
OPCB
k/ P C B
OPCB
OPCB
OPCB
OPCB
OPCB
r—
FISH
COVER/
OTHER
(10m x 20m Plot)
Filamentous Algae
Macrophytes
Woody Debris
>0.3m{BIG)
Brush/Woody Debris
<0.3 m {SMALL)
Live Trees in Stream
Overhanging Veg.
=<1 m of Surface
Undercut Banks
Boulders/Ledges
I L'
COVER CATEGORIES
0 = Absent (0%)
1 = Spare (<10%)
2 = Moderate (10-40%)
3 = Heavy (40-75%)
4 = Very Heavy (>75%)
In-Channel Cover
(circle one)
01234
234
3 4
01234
01234
34
01234
01234
Artificial Structures 01234
Flag
CHANNEL CONSTRAINT
DISTANCE FROM SHORE
TO RIPARIAN VEGETATION (M)xxx
CIRCLE ONE
Channel Is Constrained.
Channel Is In Broad Valley but Constrained hv Incision.
Channel Is in Narrow Valluv but MOT veiv constrained.
Channel Is Unconstrained In Broad Valley.
CHECK ONE
OYES
ONO
I COULD READILY SEE OVER THE BANK.
I COULD NOT READILY SEE OVER THE BANK.
FLAG
Flag
Comments
CANOPY DENSITY ffi BANK
DENS IOMETER (0 TO 17 MAX)
UP
DOWN
LEFT
RIGHT
FLAG
Flag Codes: K = no measurement made: U = suspect or non-standard measurement: F1. F2. etc. = flags assigned by each field crew. Explain al1
flags in comments section on tills side. 22399
04/07/2009 NRSA Channel Riparian Boatable Back
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-13
1
•
PHAB: THALWEG PROFILE FORM - BOATABLE "
i m
SITE ID: FW08 DATE: / / 2 0
TRANSECT: 0 A-B 0 B-C 0 C-D 0 D-E 0 E-F 0 F-G 0 G-H 0 H-l 0 I-J 0 J-K
SUBSTRATE CODES CHANNEL HABITAT CODES OTHER
BH =BEDROCK,HARDPAN (SMOOTH OR ROUGH) -(LARGER THAN A CAR) PO = Pool
BL = BOULDER (250 TO 4000 mm) - BASKETBALL TO CAR) GL = Glide Off Channel = Off
CB= COBBLE (64 TO 250 mm) -(TENNIS BALL TO BASKETBALL) Rl = Riffle Channel Or
GR = COARSE TO FINE GRAVEL (2 TO 64 mm) -(LADYBUG TO TENNIS BALL) RA = Rapid Backwater
SA = SAND (0.06 TO 2 mm) - (GRITTY - UP TO LADYBUG SIZE) CA = Cascade "'
FN = SILT' CLAY /MUCK -(NOT GRITTY) FA = Falls
OT = OTHER (COMMENT ON OTHER SIDE) DR = On/Channel
REMEMBER: A = Upstream end of Reach and K = Downstream end of Reach.
THALWEG PROFILE
STA
TION
0
1
2
3
4
5
6
7
8
9
10
11
SNAG
(circle one)
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
Y
FLAG
N
N
N
N
N
N
N
N
N
N
N
N
DEPTH (Either)
UNITS: O ft O m
SONAR XX
POLE X.X
SUBSTRATE
Circle one Substrate Code
for each station
BH BL CB GR SA FN OT
BH BL CB GR SA FN OT
BH BL CB GR SA FN OT
BH BL CB GR SA FN OT
BH BL CB GR SA F'. i T
BH BL CB GR SA I-.' JT
BH BL CB i R ^A FN OT
BH Bl v,u "" SA FN OT
BH L , Tj GR SA FN OT
BH BL CB GR SA FN OT
BH BL CB GR SA FN OT
BH BL CB GR SA FN OT
CHANNEL HABITAT
Circle one Channel Habitat
Code for each station
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
•0 ',L Rl RA CA FA DR
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
PO GL Rl RA CA FA DR
OFF
CHAN.
(circle one)
Y N
Y N
Y N
Y N
Y N
Y N
Y N
Y N
Y N
Y N
Y N
Y N
FLAG
COMMENT
Flag codes: K = Sample not collected; U = Suspect sample: F1. F2. etc. = flag assigned by field crew. Explain all flags in comment sections.
I
1 04/07/2009 NRSA Thalweg Boatable
37608
\*i
•
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-14
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-15
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-16
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
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Page B-17
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^^ 04/07/2009 NRSA Fish Collection Boatable ^^
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-18
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-19
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-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-20
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ce
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-21
CHANNEL CONSTRAINT FORM - WADEABLE/BOATABLE
Reviewed by (initial):
SITE ID: FW08
DATE:
CHANNEL CONSTRAINT
CHANNEL PATTERN (Fill in one)
O One channel
O Anastomosing (complex) channel - (Relatively long major and minor channels branching and rejoining.)
O Braided channel - (Multiple short channels branching and rejoining - mainly one channel broken up by
numerous mid-channel bars.)
CHANNEL CONSTRAINT(Fill in one)
O Channel very constrained in V-shaped valley (i.e. it is very unlikely to spread out over valley or erode a
new channel during flood)
O Channel is in Broad Valley but channel movement by erosion during floods is constrained by Incision (Flood
flows do not commonly spread over valley floor or into multiple channels.)
O Channel is in Narrow Valley but is not very constrained, but limited in movement by relatively narrow
valley floor (< -10 x bankfull width)
O Channel is Unconstrained in Broad Valley (i.e. during flood it can 'ill of'-channel areas and side channels,
spread out over flood plain, or easily cut new channels by erosion) ^£ S
CONSTRAINING FEATURES (Fill in one)
O Bedrock (i.e. channel is a bedrock-dominated gorge)
O Hillslope (i.e. channel constrained in narrow V-shapod vt'lt/)
O Terrace (i.e. channel is constrained by its own ir-ision 'nto river/stream gravel/soil deposits)
O Human Bank Alterations (i.e. constrained b • rip- '-.p, landfill, dike, road, etc.)
O No constraining features
Percent of channel length with margin
in contact with constraining feature:
(0-100%)
Bankfull width:
(m)
Percent of Channel Margin Examples
100%
100%
Valley width (Visual Estimated Average): (m)
Note: Be sure to include distances between both sides of valley border for valley width.
O
If you cannot see the valley borders, record the
distance you can see and mark this box.
50%
Comments
15186
04/07/2009 NRSA Channel Constraint 2009
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-22
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-23
Reviewed by (Initials):
TORRENT EVIDENCE ASSESSMENT FORM
SITE ID: FW08
DATE:
/ 2 0
TORRENT EVIDENCE
Please fill in any of the following that are evident.
EVIDENCE OF TORRENT SCOURING:
o
01 - Stream channel has a recently devegetated corridor two or more times the width of the low flow channel. This
corridor lacks riparian vegetation with possible exception of fireweed, even-aged alder or cottonwood seedlings.
grasses, or other herbaceous plants.
O
02 - Stream substrate cobbles or large gravel particles are NOT IMBRICATED. (Imbricated means that they lie with flat
sides horizontal and that they are stacked like roof shingles -- imagine the upstream direction as the top of the "roof."
a torrent scour or deposition channel, the stones are laying in unorganized patterns, lying "every which way." In addit
many of the substrate particles are angular (not "water-worn.")
o
03 - Channel has little evidence of pool-riffle structure. (For example, could you ride a mountain bike down the channi
O
04 - The stream channel is scoured down to bedrock for substantial portion of reach.
O
05 - There are gravel or cobble berms (little levees) above bankfull level.
O
06 - Downstream of the scoured reach (possibly several miles), the e a~„• massive deposits of sediment, logs, and othe
debris.
O
07 - Riparian trees have fresh bark scars at many points alom i, <> strp^m at seemingly unbelievable heights above the
channel bed.
o
08 - Riparian trees have fallen into the channel as a res .'* o. scouring near their roots.
EVIDENCE OF TORRENT DEPOSITS:
O
09 - There are massive deposits of sediment, It qs. •"! other debris in the reach. They may contain wood and boulder
that, in your judgement, could not have bet T mi ,ed by the stream at even extreme flood stage.
o
10 - If the stream has begun to erode newly a.~ Deposits, it is evident that these deposits are "MATRIX SUPPORTED."
This means that the large particles ,ike bou ders and cobbles, are often not touching each other, but have silt, sand, s
other fine particles between them (.'•"•.(wp'ght is supported by these fine particles-- in contrast to a normal stream
deposit, where fines, if present, norm, M1 fill-in" the interstices between coarser particles.)
NO EVIDENCE:
O
11 - No evidence of torrent scouring or torrent deposits
COMMENTS
46536
04/07/2009 NRSA Torrent Evidence
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-24
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-25
VISUAL ASSESSMENT FORM - WADEABLE/BOATABLE (Frontf"
levied by (initial):
SITE ID: FW08
DATE:
I
2 0
WATERSHED ACTIVITIES AND DISTURBANCES OBSERVED
(Intensity: Blnnk=Not observed. L=Low, M=Moderate. H=Heavy)
Residential
Recreational
Agricultural
Industrial
Stream Management
L M H Reside,
H Maintained Lawns
H Construction
H Pipes. Drains
H Dumping
H Roads
H Bridge/Culverts
L M H Hiking Trails
L M H Parks. Campgrounds
L M H Primitive Parks. Camping
L M H Trash/Litter
L M H Surface Films
L M H Cropland
L M H Pasture
L M H Livestock Us<
H Orchards
M
M H
M H Irri,
lltry
H Water Withdrawal
M H Sewage Treatment
L M H Industrial Plants
L M H Mines/Quarries
L M H Oil'Gas Wells
M H Power Plants
L M H Logging
L M H Evidence of Fire
M H Odors
M H Commercial
H Liming
H Chemical Treatment
H Angling Pressure
H Dredging
H Channelization
H Water Level Flu ctuatio
H Fish Slocking
L M H Dams
SITE CHARACTERISTICS (200 m radius)
Waterbody
Character
Pristine O 5 Q 4 Q 3 Q2 O 1 Highly Disturbed
Appealing OS O4 O3 O2 O1 Unappealing
Beaver
Beaver Signs: O Absent O Rare O Common
Beaver Flow Modifications: O None O Mimr O Major
Dominant Land Use
Dominant
Land Use
Around'X' O Forest O Agriculture O ~* .nge
IT Forest, Dominant Age ,-. ft „ ~ ~r <*^
Class O 0 - 25 yrs. O 25 - 75 yr* O > 5 yrs.
O Urban
O Suburban/Town
WEATHER
GENERAL ASSESSMENT (Biotic in 90 ,., Vegetation diversity, Local anecdotal information)
37922
04/07/2009 NRSA Visual Assessment
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-26
VISUAL ASSESSMENT FORM - WADEABLE/BOATABLE (Back)""
ewed by limtiah1
SITE ID: FW08
DATE:
J
I 2 0
j / i i i
GENERAL ASSESSMENT (continued)
INVASIVE Or NUISANCE SPECIES OF LOCAL INTEREST
Record species of plants and animals that wer.. ' jser ed but are not on the invasive plant form. Examples would be Zebra Mussel or
New Zealand Mud Snail, or invasive plants or anii. -'.& of concern to a particular state. Indicate your level of confidence in your
identification, and provide some idea of how prevelant it is in the sampling reach or adjacent riparian area.
Species (Common Name)
Confidence
Prevalence
Comments
O LOW
O HIGH
O DOMINANT
O COMMON
O SPARSE
O LOW
O HIGH
O DOMINANT
O COMMON
O SPARSE
O LOW
O HIGH
O DOMINANT
O COMMON
O SPARSE
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT
O COMMON
O SPARSE
O LOW
O HIGH
O DOMINANT
O COMMON
O SPARSE
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT
O COMMON
O SPARSE
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT
O COMMON
O SPARSE
37922
04/07/2009 NRSA Visual Assessment
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-27
TRACKING AND SAMPLE STATUS - WRS
SITE ID: FW08
SENT BY:
State of Site Location:
Visit #: 0102
SENDER PHONE:
Date Collected: / / 2 0
TEAM: DATE SENT: / / 2 0
SHIPPED O FedEx O UPS O Hand Delivery
BY: ~ _.. AIRBILL/TRACKING
O Other: NUMBER:
SAMPLEABLE
O Wadeable
O Beatable
O Partial Wadeable
O Partial Beatable
O Wadeable Interrupted
O Beatable Interrupted
O Altered
NOT SAMPLEABL
O Dry - Visited
O Dry - Not Visile
O Wetland
O Map Error
O Impounded
O Other
Status Comments
Site Status Report
Temporarily
E Not Sampleable
O Not Boatable
d O Not Wadeable o ,
O Other O '
NO ACCESS ° '
O Access Denied O
O Inaccessible
O Temp Inaccessible
SAMPLE STATUS
O No Samples Collected
Vlark the samples that were collected during this site visit:
A/ater Chem {CHEM) O Enterococci (ENTE)
A/ater Chi (WCHL) O Sediment (SEDE)
A/ater Chem (PPCP) O Fish Tissue (FTIS)
3eriphyton Chi (PCHL) O Fish Voucher (VERT)
3eriphyton Bio (PBIO) O Bent Reachwide (BERW)
3e-:phyton ID (PERI) O Bent Low Gradient (BELG)
"erir.iyton APA(PAPA)
Sample ID Sample Type
C H E M
W C H L
.2 PCHL
.3 P B 1 O
C H E M
W C H L
.2 P C H L
.3 P B 1 O
Sample Types
CHEM- Water chemistry
WCHL -Water Column
Chlorophyll
PCHL - Periphyton
Chlorophyll
PBIO- Periphyton
Biomass
"' iments
Condition Codes Chain of Custody
Filled in by recip
C = Cracked jar
F = Frozen
L= Leaking
ML = Missing label
NP = Not preserve
W = Warm
OK = Sample OK
T = Thawed
lent Filled in by recipk
Date Received:
/ /
Received by:
Contact Information
snt Tracking Help:
Marlys Cappaert
PH: 541-754-4467
Lab:
Attn: Phil Monaco, Dynamac
c/o U.S. EPA
1350 Goodnight Ave
Corvallis, OR 97333
PH: 541-754-4787
monaco.phil@epamail.epa.gov
FAX THIS FORM TO 541-754-4637
OR READ TRACKING INFO TO VOICE MESSAGE CENTER:
04/07/2009 NRSA Tracking - WRS 541-754-4663
52109
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-28
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-29
TRACKING -NERL Cincinnati
SITE ID:
SENT BY:
Visit#: 0102
SENDER PHONE:
Date Collected:
State of Site Location:
TEAM:
DATE SENT:
SHIPPED O FedEx O UPS O Hand Delivery
BY:
O Other:
AIRBILL/TRACKING
NUMBER:
/ 2 0
/ 2 0
Sample ID
Sample Type
Condition
Code
P P C P
Sample Types
Condition Codes
Contact Information
PPCP - Water chemistry
Filled in by recipient
C = Cracked jar
F = Frozen
L = Leaking
ML = Missing label
NP= Not preserved
W = Warm
OK= Sample OK
T = Thawed
Tracking Help:
Marlys Cappaert
PH: 541-754-4467
Lab: NERL-Cincinnati
Attn: Dr. Angela Batt
26 W. Martin Luther King Drive
MS 642
Cincinnati. OH 45268
513-569-7284
batt.angela@epa.gov
FAX TK'O FORM TO 541-754-4637
OR READ TRACKING INFO TO VOICE MESSAGE CENTER:
541-754-4663
4197
04/07/2009 NRSA Tracking - NERL
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-30
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-31
TRACKING (BATCHED OR RETAINED SAMPLES) National Rivers and Streams Assessment
Include only all BATCHED or RETAINED samples on one form.
SENDER STATE OF
SENT BY: PHONE: SITE LOCATION: TEAM:
BATCHED SAMPLES - UNPRESERVED samples that will be batched and shipped within 2 weeks.
SHIPPED BY: o FedEx O UPS O Hand Delivery DATE SHIPPED: / / 2 0
AIRBILL/TRACKING
NUMBER:
RETAINED SAMPLES - PRESERVED samples that will be stored longer than a month at a holding facility.
O Held at address:
Site ID
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
Date Sample Collected
MM/DD/YYYY
Visit
O
O
0
o
o
o
0
o
0
o
0
o
0
0
0
o
0
o
0
o
'j
o
c
o
0
o
0
o
0
o
0
0
1
2
1
2
1
2
1
2
1
2
1
2
1
2
1
2
1
2
1
1
1
2
1
2
1
2
1
2
1
2
Sample ID
^^^
Sample Type
#of
Containers
Comments
Cond.
Code
Lab
O ACADEMY OF NATURAL SCIENCES - PHIL. PA
O BENTHIC LAB
O GLEC
O MED - DULUTH. MN
O MICHIGAN STATE UNIV.
Q NERL- CINCINNATI, OH
O OTHER
). MA
Chain of Custody Sample Types Condition Codes
Filled in by recipi
Date Received:
/ /
Received by:
ont PRESERVED - RETAINED: Fj||or| jn hy r«"MP
BERW - Benthos Reach Wide
BELG • Benthos Low Gradient ,- _ ^.._^i
VERT - Fish Vouchers £ Cracked jar
PERI - Perlphyton ID (.1) h ~ l"rozen
L = Leaking
UNPRESERVED - BATCHED: ML = Missing la
SEDE-Sedlm
HAPA - Perlph
Tracking Help: ENTE-EIIK-H:
Marlys Cappaert
p) 541-754-4467
ient
bel
snt Enzyme NP = Not preserved
sue W = Warm
ytoiiAPA(.4) OK = Sample OK
coccl T = Thawed
4?5f)4
• FAX THIS FORM TO 541-754-4637 OR READ TRACKING INFO TO VOICE MESSAGE CENTER: 541-754-4663 • . I ^m
™«_H ^H
rH/n/wnno NP«A Tr^kinn - P='t'-rv'P°Hn ?nnQ • •"! ^"
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-32
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-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-33
WADEABLE
FORMS
PACKET
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-34
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-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-35
Reviewed by (initial);
STREAM VERIFICATION FORM - WADEABLE/BOATABLE (Front)
SITE NAME:
SITE ID: FW08
DATE: / / 2 °
VISIT: O1 O2 O 3
„, . , „.. .. Don't forget to record
State of Site Location: Reach Le*g,h on back TEAM:
STREAM/RIVER VERIFICATION INFORMATION
Stream/River Verified by (fill in all that apply) O GPS O Local Contact O Signs O Roads O Topo. Map
O other (Describe Here): O Not Verified (Explain in Comments)
Coordinates Latitude North
Degrees. Minutes,
and Seconds
MAP OR
Decimal Degrees
Degrees, Minutes,
and Seconds
GPS ' ' ' ' ' '
OR
Decimal Degrees
Longitude West Sa(*e
O
O
of Are GPS Coordinates
lites w/i 10 Sec. of map?
<3 O Yes
ONo
>4
GPS Datum Used
(e.g. NAD27):
DID YOU SAMPLE THIS SITE?
O YES If YES, check one below
SAMPLEABLE (Choose method used)
O Wadeable - Continuous water, greater than 50% wadea
O Beatable
O Partial - Sampled by wading (>50% of reach sampled).
O Partial - Sampled by boat (>50% of reach sampled). Ex
O Wadeable Interrupted - Not continuous water along re;
O Beatable Interrupted - Not continuous water along re"
O Altered - Stream/River Channel Present but differ fro-i
V
GENERAL COMMENTS:
O W-> 1' NO, check one below
NON-^A" .PLEABLE-PERMANENT
ble C Ory- Vis,,ed
C Di, Jot visited
O Wetland (No Definable Channel)
Explain below O Map Error - No evidence channel/waterbody ever present
jlain below. ^ O Impounded (Underneath Lake or Pond)
ch ^ O Other (explain in comments)
h V NON-SAMPLEABLE-TEMPORARY
O Not boatable - Need a different crew - Reschedule for this year
i M. o O Not wadeable - Need a different crew - Reschedule for this year
O Other (Explain in comments)
NO ACCESS
O Access Permission Denied
O Permanently Inaccessible (Unable/Unsafe to Reach Site)
O Temporarily Inaccessible-Fire, etc. - Reschedule for next year
DIRECTIONS TO STREAM/RIVER SITE:
Record information used to define length of reach, and sketch general features of reach on reverse side.
04/07/2009 NRSA Stream Verification 2009
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-36
STREAM VERIFICATION FORM - WADEABLE/BOATABLE (Back) TS
d by
SITE NAME:
DATE:
2 0
VISIT: O1 O2 O 3
SITE ID:
FW08
TEAM:
STREAM/RIVER REACH DETERMINATION
Channel Width
Used to Define
Reach (m)
DISTANCE (m) FROM X-SITE
Upstream
Length
Downstream
Length
Total Reach
Length Intended
Comment
SKETCH MAP - Arrow Indicates North; Mark site L=Launch X=lndex T= Take Out
NOTE: If an outline map is attached here, use a continuous strip of clear tape across the top edge.
You can also attach a separate sheet with the outline map on it.
For boatable sites you can attach topo map with reach, X-site and transect locations marked.
PERSONNEL
NAME
Bio/Chem
Sampling
O
O
O
O
O
Habitat
O
O
O
O
O
Forms
Review
O
O
O
O
O
04^07/2009 NRSA Stream Verification 2009
36530
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-37
FIELD MEASUREMENT FORM - WADEABLE
Reviewed by
(initial*:
SITE ID: FW08 / /
CALIBRATION INFORMATION
Instrument manufacturer and model:
Instrument ID number: Operator:
TEMPERATURE
DO
PH
CONDUCTIVITY
rzam£r sens°rcrding ^ c— ts
Barometric
Elevation OR Pressure(mm Cahbration Displayed
Hg) Value Value r'a9
O ft O mg/L O mg/L
O m ' . . O % . . . . ' . . O %
Cal. STD 1 Description Cal. STD 1 Value Cal. STD 2 Description Cal. STD 2 Value
Calibration Verified with Quality Control Sample (DCS)
QCS Description QCSTrue QCS Measured Flag
I
I • • .' '. . .
Cal. STD 1 Description Cal. STD 1 Value C ,. STD 2 Description Cal. STD 2 Value
Calibration Verif -i v> '*' , Quality Control Sample (QCS)
QCS Description QCS True
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-38
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-39
DISCHARGE FORM - WADEABLE
Reviewed by (Initials):
SITE ID:
FW08
DATE: /
/ 2 0
O Velocity Area
Distance Units
Oft O cm
Depth Units
O ft O cm
Velocity Units
O ft/s XX.X O m/s X.XX
Dist. from Bank Depth Velocity Flag
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
0
OQ Value
Flag
|
1
^^
/^r"
^£/
Repeat
1
2
3
4
5
O Timed Filling
Volume (L)
Time (s)
Flag
O Neutral Bouyant Object
Float Di '.
O ft O m
(s)
I Flag
Float 1
.
Float 2
^
Float 3
.
Cross Sections on Float Reach
Width
O ft O m
Depth 1
O ft O cm
Depth 2
Depth 3
Depth 4
Depth 5
If discharge is determined directly
in field, record value here: Q =
Upper Section
Middle Section
O cfs O m3/s Fl
Lower Section
.AG
Comments
Flag Codes: K = No measurement or observation made: U = Suspect measurement or observation; Q = Unacceptable QC
check associated with measurement: Z = Last station measured (if not Station 20); F1. F2. etc. = Miscellaneous flags
assigned by each field crew. Explain all flags in comments section.
04/07/2009 NRSA Stream Discharge
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-40
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-41
SAMPLE COLLECTION FORM - WADEABLE (Front)
Reviewed by
(Initial):
46387
SITE ID: FW08
DATE:
2 0
WATER CHEMISTRY (4-L CUBITAINER)
No Sample Collected Q
Sample ID
Sample
Category'
Chilled
Comments
OP
OD
o
OD
O
WATER COLUMN CHLOROPHYLL (Target Volume = 1000 mL; max vol = 2000 mL) No Sample Collected Q
Sample ID
Sample
Category'
Volume
Filtered (mL)
Frozen
Comments
OP
O
OP
OD
O
WATER CHEMISTRY PPCP (AmberGlass Bottle)
No Sample Collected O
Sample ID
Sample chi||ed
Category'
Comments
OP
O
OP
O
REACH WIDE BENTHOS S/^/lPI E
No Sample Collected Q
TRANSECT
K
SUBSTRATE CHAN.
OF
OG
Oc
Rapid
O GL
OG
O
Oc
O GL
OG
O GL
O
Oc
OKI
Sample ID
Sample
Category'
OP
OP
OD
OF
OG
Oi
OF
O
Or |O
O.
O- o «
Pre-
served
0_RI _ ,
ORJC^|QRA|Q.
O
OG
O
O
OF
OP
Oc
OF
O GL
OG
O HI
O
O GL
Oo
ORI
O
OF
OP
O GL
OG
O
O
Oo o
OF
OP
O GL
OG
ORI
Oc
Oo o
O GL
ORI
O
o
Comments
LOW - GRADIENT BENTHOS SAMPLE
No Sample Collected O
Transect
Location (LCR):
A
L Oc OR O
B
L Oc OR O
C
LOc OR
D
O LOc OR
E
OUOcOROLOcORO
G
L oc OR o
H
L oc OR o
L Oc
K
LQC OR o
Dominant
Substrate:
ONE PER TRaMSEf
OF Oc
O G O OT
OF Oc
O G O OT
OF O c
O G O OT
OF Oc
O G O OT
OF O c
O G O OT
OF Oc
O G O OT
Of Qc
O G O OT O '
O
O OT
OF Oc
O G O OT
OF O
O G O OT
OF Oc
O G O OT
Channel:
ONEPERTRAMSECl
OP O RA
O RI O GL
O OT
OP O RA
O RI O GL
OOT
OP O RA
O RI O GL
O OT
OP O
O RI O GL
O OT
OP O RA
O RI O GL
O OT
OP O RA
O RI O GL
O OT
OP O RA O P
O RI O GL
O OT
O RI O GL
O OT
OP O
O HI O GL
O OT
OP O
O RI O GL
O OT
OP O RA
O RI O GL
O OT
Dominant
Edge:
(L and R)
NEPERTRiNSECTl
Ou Os
OR O"
OL Oo<
Q OT
O" O s
OR O "
O L O OG
Q OT
O u O s
OR OM
O L O OG
O OT
O " O
OR O
O L O OG
Q OT
O u O s
OR OM
O L O OG
Q OT
Qu Os
OR O"
OL O o<
Q OT
O u Os
OR OM
O L
Q DT
Ou Os
OR OM
) L O OI
Q OT
u Q s
OR OM
O L O OG
Q OT
O" Os
OR OM
O L OO!
Q OT
O" O s
OR O M
O L O OG
Q OT
Edge:
U = Undercut S = Snag R = Rootwad M = Macrophyte bed
L = Leaf Litter OG = Organic deposits OT = Other or Co-Dominant
Substrate:
F = Fine/Sand C = Coarse substrate
G = Gravel OT = Other (Explain In
comment section below)
Channel:
p = Pool RI = Riffle GL = Glide
RA = Rapid OT = Other (Explain In
comment section below)
Sample ID
Sample
Category'| No. Jars
Pre-
served
Comments
OP
OD
O
OP
OD
O
Flag codes: K = No msasursmanf or observation rndtlie: U = Si>:>pw,_-t measurement or ol>:,
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-42
m 33
Reviewed by
46387
SAMPLE
- WADEABLE (Back)
SITE ID: FW08
DATE:
1 /2 0
COMPOSITE PERIPHYTON SAMPLE - Primary No Sample Collected O
Sample ID
Assemblage ID (.1)
(50-mL tube)
Sample Vol. (mL)
Flag Preserved
0
Sample
Category *
OP
OD
Composite Volume (mL)
Chlorophyll (.2)
(GF/F Filter)
Sample Vol. (
nL) Flag
Frozen
0
Number of transects sampled (0-11):
Biomass (.3)
(GF/C Filter)
Sample Vol. (mL)
Flag
COMPOSITE PERIPHYTON SAMPLE
Sample ID
Assemblage ID (.1)
(50-mL tube)
Sample Vol. (mL)
Flag
Flag Preserved
O
Sample
Category '
OP
OD
Composite Volume (mL)
Chlorophyll (.2)
(GF/F Filter)
Sample Vol. (
nL) Flag
Frozen
Frozen
0
Flag
APA (.4)
(50-mL tube)
Sample Vol. (mL)
Flag
Frozen
O
No Sample Collected Q
Number of transects sampled (0-11):
Biomass (.3)
(GF/C Filter)
Sample Vol. (mL)
,
Flag
Comments
Frozen
0
Flag
APA (.4)
(50-mL tube)
Sample Vol. (mL)
Flag
Frozen
O
s3\s*
tX
SEDIMl NT* HEMISTRY/ ENZYMES No Sample Collected O
Sample ID
Sample
Category '
OP
OD
OP
OD
Composite Volup- . Trar ;ects Chilled
. . O
. . 0
Comments
ENTEROCOCCI (Target Volume = 250 mL) No Sample Collected O
Sample ID
One unique ID per line
Flag
Sample
Cate-
gory'
OP
OD
OP
OD
OP
OD
OF
Time
Collected
(hhrnm)
Depth
Collected
(m)
Sample
Volume
(mL)
Fill. Start
Time
(hhmm)
Volume Filtered
(Target = 50 ml) "
Filt. 1
Fill. 2
Filt. 3
Filt. 4
Filt. End
Time
(hhmm)
Time
Frozen
(hhmm)
Flag
Comment
* Sample Categories: P = Primary: D = Field Duplicate; F= Filter Blank (Enterococcl sample only) Filter blank is collected at visit where field duplicate sample is NOT taken.
** If <25 ml of buffer solution was used to rinse filter, indicate with an F flag and note in comment section which filters) were affected along with the approximate volume(S) of buffer
solution used.
04/07/2009 NRSA Sample Collection Wadeable 2009
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-43
m
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-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-44
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-45
•
H PHAB: THALWEG PROFILE & WOODY DEBRIS FORM - WADEABLE Reviewed by (initial
LL. ^;
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-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-46
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-47
3 J2
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-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-48
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-49
•
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-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-50
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-51
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-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-52
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-53
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-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
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-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-55
CHANNEL CONSTRAINT FORM - WADEABLE/BOATABLE
Reviewed by (initial):
SITE ID: FW08
DATE:
CHANNEL CONSTRAINT
CHANNEL PATTERN (Fill in one)
O One channel
O Anastomosing (complex) channel - (Relatively long major and minor channels branching and rejoining.)
O Braided channel - (Multiple short channels branching and rejoining - mainly one channel broken up by
numerous mid-channel bars.)
CHANNEL CONSTRAINTIFill in one)
O Channel very constrained in V-shaped valley (i.e. it is very unlikely to spread out over valley or erode a
new channel during flood)
O Channel is in Broad Valley but channel movement by erosion during floods is constrained by Incision (Flood
flows do not commonly spread over valley floor or into multiple channels.)
O Channel is in Narrow Valley but is not very constrained, but limited in movement by relatively narrow
valley floor (< -10 x bankfull width)
O Channel is Unconstrained in Broad Valley (i.e. during flood it can 'ill of%channel areas and side channels,
spread out over flood plain, or easily cut new channels by erosion) ^£ S
CONSTRAINING FEATURES (Fill in one)
O Bedrock (i.e. channel is a bedrock-dominated gorge)
O HiIIslope (i.e. channel constrained in narrow V-shap^d vt'lt.O
O Terrace (i.e. channel is constrained by its own ir~ision :ntu river/stream gravel/soil deposits)
O Human Bank Alterations (i.e. constrained b • rip- • ^p, landfill, dike, road, etc.)
O No constraining features
Percent of channel length with margin
in contact with constraining feature:
(0-100%)
Bankfull width:
(m)
Percent of Channel Margin Examples
100%
100%
(m)
Valley width (Visual Estimated Average):
Note: Be sure to include distances between both sides of valley border for valley width.
O
If you cannot see the valley borders, record the
distance you can see and mark this box.
50%
50%
Comments
15186
04'07.;2009 NRSA Channel Constraint 2009
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-56
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-57
Reviewed by (Initials):
TORRENT EVIDENCE ASSESSMENT FORM
SITE ID: FW08
DATE:
/ 2 0
TORRENT EVIDENCE
Please fill in any of the following that are evident.
EVIDENCE OF TORRENT SCOURING:
o
01 - Stream channel has a recently devegetated corridor two or more times the width of the low flow channel. This
corridor lacks riparian vegetation with possible exception of fireweed, even-aged alder or cottonwood seedlings,
grasses, or other herbaceous plants.
O
02 - Stream substrate cobbles or large gravel particles are NOT IMBRICATED. (Imbricated means that they lie with flat
sides horizontal and that they are stacked like roof shingles - imagine the upstream direction as the top of the "roof."
a torrent scour or deposition channel, the stones are laying in unorganized patterns, lying "every which way." In addit
many of the substrate particles are angular (not "water-worn.")
o
03 - Channel has little evidence of pool-riffle structure. (For example, could you ride a mountain bike down the channi
O
04 - The stream channel is scoured down to bedrock for su bstantial portion of reach.
05 - There are gravel or cobble berms (little levees) above bankfull level.
O
06 - Downstream of the scoured reach (possibly several miles), the e a-j massive deposits of sediment, logs, and otht
debris.
O
07 - Riparian trees have fresh bark scars at many points alom i, •? strp-im at seemingly unbelievable heights above the
channel bed.
o
08 - Riparian trees have fallen into the channel as a res ,'* o. scouring near their roots.
EVIDENCE OF TORRENT DEPOSITS:
O
09 - There are massive deposits of sediment, li qs. "D other debris in the reach. They may contain wood and boulder
that, In your judgement, could not have bet i mi ,ed by the stream at even extreme flood stage.
10 - If the stream has begun to erode ^ewiy a.« deposits, it is evident that these deposits are "MATRIX SUPPORTED."
This means that the large particles ,ike bou Jers and cobbles, are often not touching each other, but have silt, sand, i
other fine particles between them \^r .< wp'ght is supported by these fine particles- in contrast to a normal stream
deposit, where fines, if present, norm, '!• 'fill-in" the interstices between coarser particles.)
NO EVIDENCE:
O
11 - No evidence of torrent scouring or torrent deposits
COMMENTS
46536
04/07/2009 NRSA Torrent Evidence
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-58
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-59
VISUAL ASSESSMENT FORM - WADEABLE/BOATABLE (Frontf
eviewed by (initial):
SITE ID: FW08
DATE:
2 0
WATERSHED ACTIVITIES AND DISTURBANCES OBSERVED
(Intensity: Blank=Not observed. L=Low, M=Moderate, H=Heavy)
Residential
Recreational
Agricultural
Industrial
Stream Management
H Residences
H Maintained Lawns
H Construction
H Pipes, Drains
H Dumping
H Road.
H Bridge-Culverts
L M H Hiking Trails
L M H Parks. Campgrounds
L M H Primitive Parks. Campii
L M H Trash/Litter
L M H Surface Films
L M H Cropland
L M H Pasture
L M H Livestock Use
L M H Orchards
L M H Poultry
L M H Irrigation Equip.
L M H Water Withdraws
H Industrial Plants
H Mines/Quarries
H Oil' Gas Wells
H Power Plants
H Logging
M H Chemical Treatment
L M H Angling Pr<
H Dredging
H Crmnwllrauon
L M H Sewage Treatment
L M H Evldenc,
M H Co,
of Fir
M H Water L.V.I Flu
M H Fish Stocking
M H Dams
SITE CHARACTERISTICS (200 m radius)
Waterbody
Character
Pristine
Appealing
O5
O5
O4
O4
O3
O3
O2
O2
O1
O1
Highly Disturbed
Unappealing
Beaver
Beaver Signs: O Absent O Rare O Common
Beaver Flow Modifications: O None O Minor O Major
Dominant
Land Use
Dominant Land Use -. ,_ ,-%,..
Around'X1 O Forest O Agriculture
If Forest, Dominant Ago Q „ . 25 yrs O25-75yr«
O "" .nge
O > '5 yrs.
O Urban
O Suburban/Town
WEATHER
GENERAL ASSESSMENT (Biotic in 90 „/ Vegetation diversity, Local anecdotal information)
37922
04/07/2009 NRSA Visual Assessment
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-60
VISUAL ASSESSMENT FORM - WADEABLE/BOATABLE (Backpwetl
SITE ID: FW08
DATE:
2 0
GENERAL ASSESSMENT (continued)
INVASIVE OR NUISANCE SPECIES OF LOCAL INTEREST
Record species of plants and animals that werv. •• jser ed but are not on the invasive plant form. Examples would be Zebra Mussel or
New Zealand Mud Snail, or invasive plants or anii. - s of concern to a particular state. Indicate your level of confidence in your
identification, and provide some idea of how prevelant it is in the sampling reach or adjacent riparian area.
Species (Common Name)
Confidence
Prevalence
Comments
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
37922
04/07/2009 NRSA Visual Assessment
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-61
TRACKING AND SAMPLE STATUS - WRS
SITE ID: FW08
SENT BY:
Visit #: 0102
SENDER PHONE:
Date Collected: / / 2 0
State of Site Location: TEAM: DATE SENT: / / 2 0
SHIPPED O FedEx O UPS O Hand Delivery
BY- ,-, ,.,. AIRBILL/TRACKING
u other: NUMBER:
Site Status Report
SAMPLEABLE
O Wadeable
O Boatable
O Partial Wadeable
O Partial Boatable
O Wadeable Interrupted
O Boatable Interrupted
O Altered
NOT SAMPLEABLE
O Dry - Visited
O Dry - Not Visited
O Wetland
O Map Error
O Impounded
O Other
Status Comments
Temporarily
Not Sampleable
O Not Boatable
O Not Wadeable o \
O Other O '
NO ACCESS ° '
O Access Denied O
O Inaccessible
O Temp Inaccessible
SAMPLE STATUS
O No Samples Collected
Hark the samples that were collected during this site visit:
Water Chern (CHEM) O Enterococci (ENTE)
Water Chi (WCHL) O Sediment (SEDE)
Water Chem (PPCP) O Fish Tissue (FTIS)
3eriphyton Chi (PCHL) O Fish Voucher (VERT)
Deriphyton Bio (PBIO) O Bent Reachwide (BERW)
De--:phyton ID (PERI) O Bent Low Gradient (BELG)
"erir.iyton APA(PAPA)
Sample ID Sample Type
C H E M
W C H L
.2 P C H L
.3 P B I O
C H E M
W C H L
.2 P C H L
.3 P B I 0
Sample Types
CHEM - Water chemistry
WCHL -Water Column
Chlorophyll
PCHL-Periphyton
Chlorophyll
PBIO - Periphyton
Biomass
Condition Codes
Filled in by recipie
C = Cracked jar
F = Frozen
L = Leaking
ML= Missing label
NP = Not preserved
W = Warm
OK= Sample OK
T = Thawed
"'• 11 merits
Chain of Custody
nt Filled in by recipk
Date Received:
/ /
Received by:
Contact Information
snt Tracking Help:
Marlys Cappaert
PH: 541-754-4467
Lab:
Attn: Phil Monaco, Dynamac
c/o U.S. EPA
1350 Goodnight Ave
Corvallis, OR 97333
PH: 541-754-4787
monaco.phil@epamail.epa.gov
FAX THIS FORM TO 541-754-4637
OR READ TRACKING INFO TO VOICE MESSAGE CENTER:
04/07/2009 NRSA Tracking-WRS 541-754-4663
52109
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-62
This page is intentionally blank
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-63
• TRACKING -NERL Cincinnati •
SITE ID: FW08
SENT BY:
Visit #: O1 O2 Date Collected: / / 2 0
SENDER PHONE:
State of Site Location: TEAM: DATE SENT: / / 2 0
SHIPPED O FedEx O UPS O Hand Delivery
BY' ,-. „,.. AIRBILL/TRACKING
O Other: NUMBER:
Sample ID Sample Type Comments
P P C P
Sample Types
PPCP - Water chemistry
Condition Codes
Filled in by recipient
C = Cracked jar
F = Frozen
L = Leaking
ML = Missing label
NP= Not preserved
W = Warm
OK = Sample OK
T = Thawed
•^
Chain of Custody
Filled in by recipi nt
Date Received:
/ /
Received r ••
vP
Condition
Code
Contact Information
Tracking Help:
Marlys Cappaert
PH: 541-754-4467
Lab: NERL -Cincinnati
Attn: Dr. Angela Batt
26 W. Martin Luther King Drive
MS 642
Cincinnati, OH 45268
513-569-7284
batt.angela@epa.gov
FAX TK'r> FORM TO 541-754-4637
OR READ TRACKING INFO TO VOICE MESSAGE CENTER:
541-754-4663
4197
04/07/2009 NRSA Tracking-NERL
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-64
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-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page B-65
TRACKING (BATCHED OR RETAINED SAMPLES) National Rivers and Streams Assessment
Include only all BATCHED or RETAINED samples on one form.
SENDER STATE OF
SENT BY: PHONE: SITE LOCATION: TEAM:
BATCHED SAMPLES - UNPRESERVED samples that will be batched and shipped within 2 weeks.
SHIPPED BY: o FedEx O UPS O Hand Delivery DATE SHIPPED: / / 2 0
AIRBILL/TRACKING
NUMBER:
RETAINED SAMPLES - PRESERVED samples that will be stored longer than a mo ith at a holding facility
O Held at address:
Site ID
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FWOS
FW08
FWOS
FWOS
FWOS
FWOS
Date Sample Collected
MM/DD(YYYY
Visit
O
O
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
'J
o
c
o
o
o
o
o
o
0
o
o
1
2
1
2
1
2
1
2
1
2
1
2
1
2
1
2
1
2
1
1
1
2
1
2
1
2
1
2
1
2
Sample ID
Sample Type
#of
Containers
Comments
Cond.
Code
Lab
Q ACADEMY OF NATURAL SCIENCES - PHIL. PA
O BENTHIC LAB
O GLEC
Q MED - DULUTH. MN
O MICHIGAN STATE UNIV.
O NERL ' CINCINNATI, OH
O OTHER
). MA
Chain of Custody Sample Types Condition Codes
Filled in by recip
Date Received:
/ /
Received by:
ont PRESERVED - RETAINED: pj|M in hy r°"-in
BERW • Benthos Reach Wide
BELG - Benthos Low Gradient ~ „ c***.^\,nri •„..
VERT - Fish Vouchers r " Cracked )ar
PERI - Perlphyton ID (.1) h ~ F"rozen
L = Leaking
UNPRESERVED - BATCHED: ML = Missing la
SEDE - Sedlm
I ••— . 1.311 I u
Tracking Help: ENTE - Entero
Marlys Cappaert
p) 541-754-4467
ient
bel
=mt Enzyme NP = Not preserved
sue W = Warm
ytonAPA(.4) OK = Sample OK
coccl T = Thawed
425 04
FAX THIS FORM TO 541-754^1637 OR READ TRACKING INFO TO VOICE MESSAGE CENTER: 541-754-4663
04/07/2009 NRSA Tracking - Batch/Retain 2009
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page B-66
This page is intentionally blank
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page C-1
APPENDIX C
Shipping and Tracking
Guidelines
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National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page C-2
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-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page C-3
Tracking Forms
If you have access to a computer, fill out the electronic tracking forms
• Be careful to fill out all information accurately and completely
• If you do not have a printer, you will need to include the paper form in the cooler
3 Forms
1 - Tracking and Sample Status -WRS
• This form is filled out for the samples that are shipped immediately after each sampling
event (water chemistry, AFDM, and both chlorophyll samples)
• All of these samples will go together in one cooler to the EPA Corvallis lab
• Save form according to the file naming convention on the bottom of form
• Email to address on bottom of form and print form to include in the shipping cooler
*Emailing the electronic WRS form serves as the "status report" for that sampling event
2 - Tracking (Batched and Retained Samples)
• BATCHED samples are held & shipped within 2 weeks. Send form when SHIPPED.
• RETAINED samples are stored over a month at a holding facility. Send form when
COLLECTED and when SHIPPED
• Do not combine both BATCHED and RETAINED samples on the same form
• Use one tracking form for each laboratory
• Save form according to the file naming convention on the bottom of form
• Email to address on bottom of form and print form to include in the shipping cooler
3 - Tracking - NERL - Cincinnati
• A subset of urban sites that are 5th order or greater will be sampled for PPCP
contaminants.
• Both of the PPCP samples (water and fish tissue) will go to the EPA NERL Cincinnati lab
• Save form according to the file naming convention on the bottom of form
• Email to address on bottom of form and print form to include in the shipping cooler
If you cannot use a computer before shipping:
• Fill out the paper version of the tracking form
• Notify the Information Management Center (contact info on bottom of form) - FAX form
or leave voice message with ALL info from the form
• Include the form in the shipping cooler
• Make sure to FAX or leave a voice message BEFORE the form is sealed in the cooler!
Status Report
• After each site, the Field Team Leader must file a status report with the Information
Management Center and the Field Logistics Coordinator to track visits/samples and to
describe activities, problems, and requests
• Emailing the electronic WRS form serves as the status report!
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page C-4
• If the form cannot be emailed, faxing or phoning the information serves as the status
report
SHIPPING GUIDELINES
Before shipping, it is very important to preserve each sample as directed in the sample
collection portion of this Field Operations Manual.
• Preserve the samples as specified for each indicator before shipping (Fig. C-1).
• Be aware of the holding times for each type of sample (Table C-1):
• Enterococci samples must be filtered and frozen on dry ice within 6 hours of collection
• Fish tissue samples must be frozen on dry ice as soon as possible (hold on wet ice until
freezing on dry ice).
• Fish voucher specimens are held on wet ice until being preserved in formalin in the
laboratory.
• Water chemistry samples (including PPCP water samples) must be shipped within 24
hours of collection.
• Chlorophyll a has a longer holding time, but will be sent with the water chemistry
samples since they are going to the same laboratory.
• The remaining samples must be preserved immediately upon collection; they may then
be sent in batches to the appropriate laboratory.
• The sediment enzyme sample has a two week holding time.
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page C-5
f ENTERO "^
COCCI
(ENTE)
4 filters in
was j
1
f WATER ^1
CHEM STRY
(CHEW
4 I cubrtainer
^ )
' 1
Hold on
wet ice
i
f WATER A
CHLORO-
PHYLL
(WCHL)
Filiei inSOmL
^ wb« J
r i
Hold on
wet ice
F
Prepare 4
filters
1
r 1
Place in bubble
wrap bag
l
PERIPHYTON\
CHLORO-
PHYtL
(PCHL)
Filter in 50 ml
L tube J
r 1
(PERIPHYTONI
BlOrV
ASS
(PBIO)
Filter in
50 ml
1 tube J
' i
f" PPCP ^
WATER
Urban sttes
only
^ 2-500mL J
1
Hold on wet ice
\
' i
' 1
fuRBANFISlA
TISSUE
(FTIS)
In foil & dbl
^ bagged J
' ^
Hold on
wet ice
r
Prepare 1 Filter Each
r 1
Tape lid
r 1
Freeze on
dry ice
i
r 1
' 1
Tape lid. wrap in foil
I 1
Preserve
on wet ice
r 1
Ship in
batches
weeks
1
r 1
' 1
Tape lid
' 1
CNON URBAM^
FISH1
SSUE
(FTIS)
In foil & dbl
^ bagged J
r 1
Hold in live
well
i
/FISH VOUCH"1!
El
S
(VERT)
Jans) with
whole fish
,
Hold in live
well
r T
ID. Measure.
and Dispatch
r 1
Tape lids
' ^
Freeze immediately on dry ice
r 1
r 1
' 1
f SEDIMENT "^
(SEDE)
500 ml bottle
v
,
Preserve in
buffered
formalin
r
ID Measure
and Dispatch
-
Foil wrap /
plastic bag
' ^
Preserve
on wet ice
' ^
SHIP ON WET ICE ASAP AFTER COLLECTION
f 1
OVER-
NIGHT
COURIER
NO
Saturday
delivery!
' 1
F
1
' 1
[
Foil wrap /
plastic bag
>
Freeze on
dry ice
r i
SHIP ON
WET ICE
ASAP
r 1
OVERNIGHT COURIER REQUIRED
Saturday deliver OK
1
s^^.
1
1
y
PERIP
A
HYTOt?
>A
(PAPA)
50 mU tube
r
Hold in
bucket
,
Group by
species.
enclose in
sleeves w/
tags
r
Freeze on
dry ice
'
SHIP ON
DRY ICE
(Maybe
batched)
f 1
OVER-
NIGHT
COURIER
NO
Saturday
delivery!
1
^
1
lYTOr?)
3
(PERI)
50 ml tube
Hold on
wet ice
,
Mm and
composite inlo
500 ml bottle
;
J
^ BENTHOS ^
(BERW)
(B6LG)
X J
,
Hold on
wet ice
-
Mix a draw 50
ml into tube
-
Tape lid
r
Rfjp ace forma-
lin w/ ETOH
after 5-7 days
r
SHIP ON
DRY ICE
(Maybe
batctad]
r
OVER-
NIGHT
COURIER
NO
Saturday
delivery!
L. .J
,
Hold in sieve
buckel(s)
•
Mix D(aw50
rnL, add 1mL
Luasls
'
Tape lid
'
Store re-
frigerated
r
Ship in
batches as
needed
>
OVER-
NIGHT
COURIER
NO
Saturday
delivery!
r
Composite into
!-LboWe(3).
5 1« full
\
Tape lid
'
Freeze on
dry ice
'
SHIP ON
WET ICE
In batches
r
Formalin /
ETOH must be
shipped as
Dariqarous
Gtjudij
via Ground
carrier
,
Fill w/ ETOH
Tape liclisi
,
Store re-
frigerated
,
SHIP ON
WET ICE
In naicivcs
:'.',' • . ..:•:,.•<•-,
r
OVER-
NIGHT
COURIER
NO
Saturday
delivery!
r JL Jt ,
>
Ship in
batches
•.veeks
,
OVER-
NIGHT
COURIER
NO
Saturday
delivery!
. ,
r
Ship in
batches as
needed
r
OVER-
NIGHT
COURIER
NO
Saturday
delivery1
r
Formalin /
shipped as
Da nacrous
Goods
via Ground
earner.
, J^ ^
WRS - EPA Lab
Corvallis, OR
MSU
CIN - EPA Lab
Cincinnati, OH
PPCP urban met
sites only
Sample Type
Sample Code
Container
Field Storage
Sample Prep
Storage after
preparation
Shipment
Lab
Periphyton ID Sample (PERI) will be shipped to one of two labs depending on the state from which the sample was obtained.
Find your state below and ship the sample to corresponding lab:
MSU (Michigan State University in East Lansing, Ml): AL. AR. FL. GA. IL. IN. IA. KS. KY. LA. Ml. MN MS, MO, NE. NC. OH, OK, SC, TN, TX. Wl
PHIL (Academy of Natural Sciences in Philadelphia, PA) AZ. CA. CO. CT. DE. ID. MA, MD, ME. MT. ND. NH. NJ. NM, NV. NY. OR, PA, Ri, SO, UT, VA. vr, WA. wv, WY
Field Forms: All field forms should be reviewed and sent in to the Information Management Coordinator every 2 weeks
Figure C-1. Sample packaging and shipping summary
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page C-6
SAMPLE
Water Chemistry
Chlorophyll a
Periphyton - chlorophyll a
Periphyton Biomass - AFDM
Sediment enzymes
Periphyton - APA
Periphyton - ID
Benthic macroinvertebrates
Fish Vouchers
Fecal Indicator
Fish Tissue (non urban sites)
PRESERVATIVE
Wet ice
Dry ice in field
Dry ice in field
Dry ice in field
Wet ice in field;
refrigerate to hold
Wet ice in field; hold
in freezer
1 ml Lugol's
95% Ethanol
Formalin
Dry ice in field; hold
in freezer; MUST be
filtered & frozen
within 6 hours of
collection
Dry ice in field; hold
in freezer
PACKAGING FOR
SHIPMENT
Ship in cooler with wet ice
Ship in cooler with wet ice
Ship in cooler or sturdy
container
Ship in cooler with DRY ICE
Ship in cooler with DRY ICE
HOLDING TIME
24 hours; ship
these samples
together (Corvallis
lab)
Batch; ship these
samples together
every 2 weeks
(Duluth lab)1
Batch; ship every 2
weeks
Batch; ship every 2
weeks (Region 1
lab)
Batch; ship every 2
weeks to GLEC lab
"Urban fish tissue and PPCP water samples are only collected at pre-selected urban 5 order or greater sites
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page C-7
*PPCP Fish Tissue (urban
sites)
*PPCP Water (urban sites
only)
Dry ice in field; hold
in freezer
Wet ice
Ship in cooler with DRY ICE
Ship in cooler with wet ice
Batch; ship every 2
weeks to EPA
Cincinnati lab
24 hours; ship to
EPA Cincinnati lab
Sediment enzyme samples should not be frozen and must be shipped within two weeks of sampling
When ice is used for shipment (water chemistry, chlorophyll a, sediment enzymes, APA,
AFDM):
• Ensure that the ice is fresh before shipment; pack the entire cooler full with ice.
• Line the cooler with a large, 30-gallon plastic bag.
• Contain the ice separately within numerous 1-gallon self-sealing plastic bags.
Double-bag the ice.
• Use white or clear bags and label with a dark indelible marker. Label all bags of ice
as "ICE" to prevent misidentification by couriers of any water leakage as a possible
hazardous material spill.
• Place bagged samples and bags of ice inside the cooler liner and seal the liner.
• Secure the cooler with strapping tape.
When dry ice is used for shipping (fish tissue and fecal indicator samples):
• Indicate dry ice on shipping airbill.
• Label cooler with a Class 9 Dangerous Goods label.
• Securely tape the cooler drainage open to prevent pressure build-up in the cooler.
• Secure the cooler with strapping tape
• See "Dry Ice Shipping Protocols" at the end of this Appendix.
WATER CHEMISTRY and CHLOROPHYLL-a (from water sample and periphyton sample)
• Water Chemistry
Stored in a 4-L cube container
• Confirm that the cube container is labeled and covered with clear tape.
• Place the cube container in a second bag inside the cooler liner.
• Chlorophyll a
Two filters each stored in a 50-mL steam-top centrifuge tube wrapped with aluminum foil
• Confirm that the labels with sample IDs are completed and covered with clear tape.
• Place the centrifuge tubes in a 1-qt self-sealing plastic bag.
• Place the bag in a1-gal self-sealing plastic bag and place inside cooler liner with
water chemistry sample.
SEDIMENT ENZYMES SAMPLES
Stored in 500 mLjars
• Confirm that the label with sample ID is completed and covered with clear tape.
• Place the 500 mL jar in a 1-gal self-sealing plastic bag and place inside cooler liner.
PERIPHYTON SAMPLES
ID samples preserved with Lugol's solution and sealed at the site.
• Confirm that the label with sample ID is completed and covered with clear tape.
• Verify that the bottle is sealed with electrical tape.
-------
National Rivers and Streams Assessment Final Manual
Field Operations Manual Date: April 2009
Page C-8
• Place the sealed bottles in a gallon-size self-sealing plastic bag.
• Place the bagged samples in the appropriate shipping container.
• Surround the jars with crumpled newspaper, vermiculite, or other absorbent material.
• Samples can be held and shipped in batches to the laboratory for analysis.
AFDM and APA samples held frozen until shipment
• Confirm that the label with sample ID is completed and covered with clear tape.
• Place the frozen samples in a 1-gal self-sealing plastic bag and place inside cooler liner.
BENTHIC INVERTEBRATE SAMPLES
Preserved in 95% ethanol and sealed at the site.
• Confirm that the label with sample ID is completed and covered with clear tape.
• Check to make sure jars are sealed with electrical tape.
• Place up to twenty 500-mL or ten 1-L jars in each cooler.
• Surround the jars with crumpled newspaper, vermiculite, or other absorbent material.
• Samples can be held and shipped in batches to the laboratory for analysis.
NOTE: These samples must be shipped as "DANGEROUS GOODS" and should be
packaged and labeled in accordance with the requirements of the chosen courier.
Alternatively, the ethanol may be decanted from the benthic invertebrate samples so that
they may be shipped using standard overnight shipping:
• Allow the samples to sit for at least 1 week to adequately preserve the organisms.
• Immediately before shipping, decant the ethanol from the samples jars, leaving enough
liquid to keep the samples moist.
• Make sure to use an overnight delivery so that the lab can immediately restore the
ethanol to the sample jars.
• Alert the laboratory so that they are aware they will need to refill the jars immediately
upon receipt.
FISH TISSUE SAMPLES
The samples need to be frozen as soon as possible after collection (within 6 hours).
• Pack the cooler with 50 Ibs of dry ice.
• Refer to the DRY ICE SHIPPING PROTOCOLS at the end of this Appendix.
• Samples may be stored on dry ice for a maximum of 24 hours. Sampling teams have the
option, depending on site logistics, of:
shipping the samples packed on dry ice (50 pounds), via priority overnight delivery
so that they arrive at the sample preparation laboratory within 24 hours of sample
collection, or
• freezing the samples within 24 hours of collection at <-20°C, and storing the frozen
samples until shipment within 2 weeks of sample collection (frozen samples will be
packed on dry ice and shipped to the sample preparation laboratory via priority
overnight delivery service).
FISH VOUCHER SAMPLES
Preserved in a laboratory with formalin
• Confirm that the label with sample ID is completed and covered with clear tape.
• Check to make sure jars are sealed with electrical tape.
• Surround the jars with crumpled newspaper, vermiculite, or other absorbent material.
• Samples can be held and shipped in batches to the laboratory for analysis.
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page C-9
NOTE: These samples must be shipped as "DANGEROUS GOODS" and should be
packaged and labeled in accordance with the requirements of the chosen courier.
FECAL INDICATOR SAMPLES
The sample needs to be filtered and frozen as soon as possible after collection (within 6 hours).
• Confirm that the container is labeled and properly sealed.
• Confirm that the bottle is labeled with the appropriate sample ID and covered with clear
plastic tape.
• Place the container in the cooler and close.
• Pack the cooler with 10-15 Ibs of dry ice (10 Ibs if using dry ice blocks or slices, 15 Ibs if
using dry ice pellets).
• Refer to the DRY ICE SHIPPING PROTOCOLS at the end of this Appendix.
• Samples can be held frozen and shipped in batches to the laboratory for analysis.
DRY ICE SHIPPING PROTOCOLS
1 . Indicate dry ice on shipping airbill
• Fill out Section 1 and Section 3 of the Fed Ex airbill with your Sender and
Recipient address and phone number.
• In Section 4, check "FedEx Priority Overnight."
• In Section 5, check "Other."
• In Section 6, under "Does this shipment contain dangerous goods?":
Check "Yes/Shipper's Declaration not required."
Check "Dry Ice," and fill out " 1 x (ami, of dry ice in kg) kg"
• In Section 7, fill out weight and declared value of package.
2. Label cooler with a Class 9 Dangerous Goods label (available from FedEx) (Fig. C-2).
Shipper's Declaration not Required
Part B is required
Dry Ice amours? must be in
kilograms,
Note: 2 Ibs, = 1 kg.
ls/arbilis must have the following:
1. "Dangerous Goods - Shipper^
Declarators not feqyired".
2. Dry lo; 9; UN 1846; III
\ J. I Kg 904
(Nurnte
Figure
• Place the label on the front
side of the cooler, not the top of the
cooler.
• Fill out #3 in the top right
hand corner of the label with the same
information as in Section 6 of the FedEx
airbill.
• Declare the weight of the
dry ice again in the lower left hand corner.
• Fill out the Sender
("Shipper") and Recipient ("Consignee")
address on the bottom of the label.
C-2. Class 9 Dangerous Goods label.
3. Securely tape the cooler drainage
open to prevent pressure build-up in the cooler. This is critical to ensure proper venting
of the dry ice.
4. Secure the cooler with strapping tape.
5. Place the completed airbill on the top of the cooler.
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Page C-10
NOTE: Not all FedEx locations will accept shipments containing dry ice. Dry ice shipments can
be shipped from "FedEx staffed" locations. You can also arrange for a pick-up from your lab or
hotel. Dry ice shipments usually cannot be shipped from FedEx Kinko's Office and Print
Centers® or Fed Ex Authorized ShipCenter® locations. These types of locations are
differentiated on FedEX.com in the "Find FedEx Locations" feature. Please be sure to call in
advance to ensure your location will accept the package for shipment.
TRACKING FORMS
A Tracking Form must be filled out to accompany each sample shipment. Please refer to
Figures 3.2 and 3.3 for examples of Tracking Forms completed for both unpreserved and
preserved samples. Be very careful to fill in the information correctly and legibly, especially the
airbill number, Site ID, and Sample ID numbers. Use the codes on the bottom of the form to
indicate sample type. The Tracking Form is to be placed in a self-sealing plastic bag and
included inside the shipping container. Before sealing the container, remember to submit the
status report (via email) to sampletracking@epa.gov (see Section 3.2.6); you will need the
information on the tracking form to fill out the status report form. For preserved samples, submit
a status report both when the samples are brought to the holding facility AND when they are
shipped to the appropriate laboratory. For each shipment, you must fill out a scanable tracking
form to include in the cooler and submit the electronic status report.
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Page D-1
APPENDIX D
Common and Scientific
Names of Fishes of the
United States
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Page D-3
Table D-1. Common and Scientific Names of Fishes of the United States
(From: Nelson, J.S., E.J. Grossman, H. Espinosa-Perez, L.T. Findley, C.R. Gilbert, R.N.
Lea, and J.D. Williams. 2004. Common and Scientific Names of Fishes from the United
States, Canada, and Mexico. American Fisheries Society, Special Publication 29, Bethesda,
Maryland.)
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Petromyzontiformes
Carcharhiniformes
Pristiformes
Myliobatiformes
Acipenseriformes
Acipenseriformes
Acipenseriformes
Acipenseriformes
Acipenseriformes
Acipenseriformes
Acipenseriformes
Acipenseriformes
Acipenseriformes
Lepisosteiformes
Lepisosteiformes
Lepisosteiformes
Lepisosteiformes
Lepisosteiformes
Amiiformes
Hiodontiformes
Hiodontiformes
Osteoglossiformes
Elopiformes
Elopiformes
Elopiformes
Anguilliformes
Clupeiformes
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Petromyzontidae
Carcharhinidae
Pristidae
Dasyatidae
Acipenseridae
Acipenseridae
Acipenseridae
Acipenseridae
Acipenseridae
Acipenseridae
Acipenseridae
Acipenseridae
Polyodontidae
Lepisosteidae
Lepisosteidae
Lepisosteidae
Lepisosteidae
Lepisosteidae
Amiidae
Hiodontidae
Hiodontidae
Notopteridae
Elopidae
Elopidae
Megalopidae
Anguillidae
Engraulidae
Ichthyomyzon bdellium
Ichthyomyzon castaneus
Ichthyomyzon fossor
Ichthyomyzon gage;
Ichthyomyzon greeleyi
Ichthyomyzon unicuspis
Lampetra aepyptera
Lampetra appendix
Lampetra ayresii
Lampetra camtschatica
Lampetra hubbsi
Lampetra lethophaga
Lampetra minima
Lampetra richardsoni
Lampetra similis
Lampetra tridentata
Petromyzon marinus
Carcharhinus leucas
Pristis pectinata
Dasyatis sabina
Acipenser brevirostrum
Acipenser fulvescens
Acipenser medirostris
Acipenser oxyrinchus
Acipenser transmontanus
Scaphirhynchus albus
Scaphirhynchus platorynchus
Scaphirhynchus suttkusi
Polyodon spathula
Atractosteus spatula
Lepisosteus oculatus
Lepisosteus osseus
Lepisosteus platostomus
Lepisosteus platyrhincus
Amia calva
Hiodon alosoides
Hiodon tergisus
Chitala ornata
Elops affinis
Elops saurus
Mega/ops atlanticus
Anguilla rostrata
Anchoa mitchilli
Ohio lamprey
chestnut lamprey
northern brook lamprey
southern brook lamprey
mountain brook lamprey
silver lamprey
least brook lamprey
American brook lamprey
river lamprey
Arctic lamprey
Kern brook lamprey
Pit-Klamath brook lamprey
Miller Lake lamprey
western brook lamprey
Klamath lamprey
Pacific lamprey
sea lamprey
bull shark
smalltooth sawfish
Atlantic stingray
shortnose sturgeon
lake sturgeon
green sturgeon
Atlantic sturgeon
white sturgeon
pallid sturgeon
shovelnose sturgeon
Alabama sturgeon
paddlefish
alligator gar
spotted gar
longnose gar
shortnose gar
Florida gar
bowfin
goldeye
mooneye
clown knifefish
machete
ladyfish
tarpon
American eel
bay anchovy
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Page D-4
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Clupeiformes Clupeidae Alosa aestivalis
Clupeiformes Clupeidae Alosa alabamae
Clupeiformes Clupeidae Alosa chrysochloris
Clupeiformes Clupeidae Alosa mediocris
Clupeiformes Clupeidae Alosa pseudoharengus
Clupeiformes Clupeidae Alosa sapidissima
Clupeiformes Clupeidae Dorosoma cepedianum
Clupeiformes Clupeidae Dorosoma petenense
Clupeiformes Clupeidae Harengula jaguana
Clupeiformes Clupeidae Opisthonema oglinum
Cypriniformes Cyprinidae Acrocheilus alutaceus
Cypriniformes Cyprinidae Agosia chrysogaster
Cypriniformes Cyprinidae Campostoma anomalum
Cypriniformes Cyprinidae Campostoma oligolepis
Cypriniformes Cyprinidae Campostoma ornatum
Cypriniformes Cyprinidae Campostoma pauciradii
Cypriniformes Cyprinidae Carassius auratus
Cypriniformes Cyprinidae Clinostomus elongatus
Cypriniformes Cyprinidae Clinostomus funduloides
Cypriniformes Cyprinidae Couesius plumbeus
Cypriniformes Cyprinidae Ctenopharyngodon idella
Cypriniformes Cyprinidae Cyprinella analostana
Cypriniformes Cyprinidae Cyprinella caerulea
Cypriniformes Cyprinidae Cyprinella callisema
Cypriniformes Cyprinidae Cyprinella callistia
Cypriniformes Cyprinidae Cyprinella callitaenia
Cypriniformes Cyprinidae Cyprinella camura
Cypriniformes Cyprinidae Cyprinella chloristia
Cypriniformes Cyprinidae Cyprinella formosa
Cypriniformes Cyprinidae Cyprinella galactura
Cypriniformes Cyprinidae Cyprinella gibbsi
Cypriniformes Cyprinidae Cyprinella labrosa
Cypriniformes Cyprinidae Cyprinella leedsi
Cypriniformes Cyprinidae Cyprinella lepida
Cypriniformes Cyprinidae Cyprinella lutrensis
Cypriniformes Cyprinidae Cyprinella nivea
Cypriniformes Cyprinidae Cyprinella proserpina
Cypriniformes Cyprinidae Cyprinella pyrrhomelas
Cypriniformes Cyprinidae Cyprinella spiloptera
Cypriniformes Cyprinidae Cyprinella trichroistia
Cypriniformes Cyprinidae Cyprinella venusta
Cypriniformes Cyprinidae Cyprinella whipplei
Cypriniformes Cyprinidae Cyprinella xaenura
Cypriniformes Cyprinidae Cyprinella zanema
Cypriniformes Cyprinidae Cyprinus carpio
Cypriniformes Cyprinidae Dionda argentosa
Cypriniformes Cyprinidae Dionda diaboli
blueback herring
Alabama shad
skipjack herring
hickory shad
alewife
American shad
gizzard shad
threadfin shad
scaled sardine
Atlantic thread herring
chiselmouth
longfin dace
central stoneroller
largescale stoneroller
Mexican stoneroller
bluefin stoneroller
goldfish
redside dace
rosyside dace
lake chub
grass carp
satinfin shiner
blue shiner
Ocmulgee shiner
Alabama shiner
bluestripe shiner
bluntface shiner
greenfin shiner
beautiful shiner
whitetail shiner
Tallapoosa shiner
thicklip chub
bannerfin shiner
plateau shiner
red shiner
whitefin shiner
proserpine shiner
fieryblack shiner
spotfin shiner
tricolor shiner
blacktail shiner
steelcolor shiner
Altamaha shiner
Santee chub
common carp
Manantial roundnose
minnow
Devils River minnow
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ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cypriniformes Cyprinidae Dionda episcopa
Cypriniformes Cyprinidae Dionda nigrotaeniata
Cypriniformes Cyprinidae Dionda serena
Cypriniformes Cyprinidae Eremichthys acros
Cypriniformes Cyprinidae Erimonax monachus
Cypriniformes Cyprinidae Erimystax cahni
Cypriniformes Cyprinidae Erimystax dissimilis
Cypriniformes Cyprinidae Erimystax harryi
Cypriniformes Cyprinidae Erimystax insignis
Cypriniformes Cyprinidae Erimystax x-punctatus
Cypriniformes Cyprinidae Exoglossum laurae
Cypriniformes Cyprinidae Exoglossum maxillingua
Cypriniformes Cyprinidae Gila alvordensis
Cypriniformes Cyprinidae Gila atraria
Cypriniformes Cyprinidae Gila bicolor
Cypriniformes Cyprinidae Gila boraxobius
Cypriniformes Cyprinidae Gila coerulea
Cypriniformes Cyprinidae Gila crassicauda
Cypriniformes Cyprinidae Gila cypha
Cypriniformes Cyprinidae Gila ditaenia
Cypriniformes Cyprinidae Gila elegans
Cypriniformes Cyprinidae Gila intermedia
Cypriniformes Cyprinidae Gila nigra
Cypriniformes Cyprinidae Gila nigrescens
Cypriniformes Cyprinidae Gila orcuttii
Cypriniformes Cyprinidae Gila pandora
Cypriniformes Cyprinidae Gila purpurea
Cypriniformes Cyprinidae Gila robusta
Cypriniformes Cyprinidae Gila seminuda
Cypriniformes Cyprinidae Hemitremia flammea
Cypriniformes Cyprinidae Hesperoleucus symmetricus
Cypriniformes Cyprinidae Hybognathus amarus
Cypriniformes Cyprinidae Hybognathus argyritis
Cypriniformes Cyprinidae Hybognathus hankinsoni
Cypriniformes Cyprinidae Hybognathus hayi
Cypriniformes Cyprinidae Hybognathus nuchalis
Cypriniformes Cyprinidae Hybognathus placitus
Cypriniformes Cyprinidae Hybognathus regius
Cypriniformes Cyprinidae Hybopsis amblops
Cypriniformes Cyprinidae Hybopsis amnis
Cypriniformes Cyprinidae Hybopsis hypsinotus
Cypriniformes Cyprinidae Hybopsis lineapunctata
Cypriniformes Cyprinidae Hybopsis rubrifrons
Cypriniformes Cyprinidae Hybopsis winchelli
Cypriniformes Cyprinidae Hypophthalmichthys molitrix
Cypriniformes Cyprinidae Hypophthalmichthys nobilis
Cypriniformes Cyprinidae lotichthys phlegethontis
roundnose minnow
Guadalupe roundnose
minnow
Nueces roundnose minnow
desert dace
spotfin chub
slender chub
streamline chub
Ozark chub
blotched chub
gravel chub
tonguetied minnow
cutlip minnow
Alvord chub
Utah chub
tui chub
Borax Lake chub
blue chub
thicktail chub
humpback chub
Sonora chub
bonytail
Gila chub
headwater chub
Chihuahua chub
arroyo chub
Rio Grande chub
Yaqui chub
roundtail chub
Virgin chub
flame chub
California roach
Rio Grande silvery minnow
western silvery minnow
brassy minnow
cypress minnow
Mississippi silvery minnow
plains minnow
eastern silvery minnow
bigeye chub
pallid shiner
highback chub
lined chub
rosyface chub
clear chub
silver carp
bighead carp
least chub
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ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cypriniformes Cyprinidae Lavinia exilicauda
Cypriniformes Cyprinidae Lepidomeda albivallis
Cypriniformes Cyprinidae Lepidomeda altivelis
Cypriniformes Cyprinidae Lepidomeda mollispinis
Cypriniformes Cyprinidae Lepidomeda vittata
Cypriniformes Cyprinidae Leuciscus idus
Cypriniformes Cyprinidae Luxilus albeolus
Cypriniformes Cyprinidae Luxilus cardinalis
Cypriniformes Cyprinidae Luxilus cerasinus
Cypriniformes Cyprinidae Luxilus chrysocephalus
Cypriniformes Cyprinidae Luxilus coccogenis
Cypriniformes Cyprinidae Luxilus cornutus
Cypriniformes Cyprinidae Luxilus pilsbryi
Cypriniformes Cyprinidae Luxilus zonatus
Cypriniformes Cyprinidae Luxilus zonistius
Cypriniformes Cyprinidae Lythrurus alegnotus
Cypriniformes Cyprinidae Lythrurus ardens
Cypriniformes Cyprinidae Lythrurus atrapiculus
Cypriniformes Cyprinidae Lythrurus bellus
Cypriniformes Cyprinidae Lythrurus fasciolaris
Cypriniformes Cyprinidae Lythrurus fumeus
Cypriniformes Cyprinidae Lythrurus lirus
Cypriniformes Cyprinidae Lythrurus matutinus
Cypriniformes Cyprinidae Lythrurus roseipinnis
Cypriniformes Cyprinidae Lythrurus snelsoni
Cypriniformes Cyprinidae Lythrurus umbratilis
Cypriniformes Cyprinidae Macrhybopsis aestivalis
Cypriniformes Cyprinidae Macrhybopsis australis
Cypriniformes Cyprinidae Macrhybopsis gelida
Cypriniformes Cyprinidae Macrhybopsis hyostoma
Cypriniformes Cyprinidae Macrhybopsis marconis
Cypriniformes Cyprinidae Macrhybopsis meeki
Cypriniformes Cyprinidae Macrhybopsis storeriana
Cypriniformes Cyprinidae Macrhybopsis tetranema
Cypriniformes Cyprinidae Margariscus margarita
Cypriniformes Cyprinidae Meda fulgida
Cypriniformes Cyprinidae Moapa coriacea
Cypriniformes Cyprinidae Mylocheilus caurinus
Cypriniformes Cyprinidae Mylopharodon conocephalus
Cypriniformes Cyprinidae Nocomis asper
Cypriniformes Cyprinidae Nocomis biguttatus
Cypriniformes Cyprinidae Nocomis effusus
Cypriniformes Cyprinidae Nocomis leptocephalus
Cypriniformes Cyprinidae Nocomis micropogon
Cypriniformes Cyprinidae Nocomis platyrhynchus
Cypriniformes Cyprinidae Nocomis raneyi
Cypriniformes Cyprinidae Notemigonus crysoleucas
Cypriniformes Cyprinidae Notropis albizonatus
hitch
White River spinedace
Pahranagat spinedace
Virgin spinedace
Little Colorado spinedace
ide
white shiner
cardinal shiner
crescent shiner
striped shiner
warpaint shiner
common shiner
duskystripe shiner
bleeding shiner
bandfin shiner
Warrior shiner
rosefin shiner
blacktip shiner
pretty shiner
scarlet shiner
ribbon shiner
mountain shiner
pinewoods shiner
cherryfin shiner
Ouachita shiner
redfin shiner
speckled chub
prairie chub
sturgeon chub
shoal chub
burrhead chub
sicklefin chub
silver chub
peppered chub
pearl dace
spikedace
Moapa dace
peamouth
hardhead
redspot chub
hornyhead chub
redtail chub
bluehead chub
river chub
bigmouth chub
bull chub
golden shiner
palezone shiner
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ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cypriniformes Cyprinidae Notropis alborus
Cypriniformes Cyprinidae Notropis altipinnis
Cypriniformes Cyprinidae Notropis amabilis
Cypriniformes Cyprinidae Notropis ammophilus
Cypriniformes Cyprinidae Notropis amoenus
Cypriniformes Cyprinidae Notropis anogenus
Cypriniformes Cyprinidae Notropis ariommus
Cypriniformes Cyprinidae Notropis asperifrons
Cypriniformes Cyprinidae Notropis atherinoides
Cypriniformes Cyprinidae Notropis atrocaudalis
Cypriniformes Cyprinidae Notropis baileyi
Cypriniformes Cyprinidae Notropis bairdi
Cypriniformes Cyprinidae Notropis bifrenatus
Cypriniformes Cyprinidae Notropis blennius
Cypriniformes Cyprinidae Notropis boops
Cypriniformes Cyprinidae Notropis braytoni
Cypriniformes Cyprinidae Notropis buccatus
Cypriniformes Cyprinidae Notropis buccula
Cypriniformes Cyprinidae Notropis buchanani
Cypriniformes Cyprinidae Notropis cahabae
Cypriniformes Cyprinidae Notropis candidus
Cypriniformes Cyprinidae Notropis chalybaeus
Cypriniformes Cyprinidae Notropis chihuahua
Cypriniformes Cyprinidae Notropis chiliticus
Cypriniformes Cyprinidae Notropis chlorocephalus
Cypriniformes Cyprinidae Notropis chrosomus
Cypriniformes Cyprinidae Notropis cummingsae
Cypriniformes Cyprinidae Notropis dorsalis
Cypriniformes Cyprinidae Notropis edwardraneyi
Cypriniformes Cyprinidae Notropis girardi
Cypriniformes Cyprinidae Notropis greenei
Cypriniformes Cyprinidae Notropis harperi
Cypriniformes Cyprinidae Notropis heterodon
Cypriniformes Cyprinidae Notropis heterolepis
Cypriniformes Cyprinidae Notropis hudsonius
Cypriniformes Cyprinidae Notropis hypsilepis
Cypriniformes Cyprinidae Notropis jemezan us
Cypriniformes Cyprinidae Notropis leuciodus
Cypriniformes Cyprinidae Notropis longirostris
Cypriniformes Cyprinidae Notropis lutipinnis
Cypriniformes Cyprinidae Notropis maculatus
Cypriniformes Cyprinidae Notropis mekistocholas
Cypriniformes Cyprinidae Notropis melanostomus
Cypriniformes Cyprinidae Notropis micropteryx
Cypriniformes Cyprinidae Notropis nubilus
Cypriniformes Cyprinidae Notropis orca
Cypriniformes Cyprinidae Notropis ortenburgeri
Cypriniformes Cyprinidae Notropis oxyrhynchus
whitemouth shiner
highfin shiner
Texas shiner
orangefin shiner
comely shiner
pugnose shiner
popeye shiner
burrhead shiner
emerald shiner
blackspot shiner
rough shiner
Red River shiner
bridle shiner
river shiner
bigeye shiner
Tamaulipas shiner
silverjaw minnow
smalleye shiner
ghost shiner
Cahaba shiner
silverside shiner
ironcolor shiner
Chihuahua shiner
redlip shiner
greenhead shiner
rainbow shiner
dusky shiner
bigmouth shiner
fluvial shiner
Arkansas River shiner
wedgespot shiner
redeye chub
blackchin shiner
blacknose shiner
spottail shiner
highscale shiner
Rio Grande shiner
Tennessee shiner
longnose shiner
yellowfin shiner
taillight shiner
Cape Fear shiner
blackmouth shiner
highland shiner
Ozark minnow
phantom shiner
Kiamichi shiner
sharpnose shiner
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ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cypriniformes Cyprinidae Notropis ozarcanus
Cypriniformes Cyprinidae Notropis percobromus
Cypriniformes Cyprinidae Notropis perpallidus
Cypriniformes Cyprinidae Notropis petersoni
Cypriniformes Cyprinidae Notropis photogenis
Cypriniformes Cyprinidae Notropis potteri
Cypriniformes Cyprinidae Notropis procne
Cypriniformes Cyprinidae Notropis rafinesquei
Cypriniformes Cyprinidae Notropis rubellus
Cypriniformes Cyprinidae Notropis rubricroceus
Cypriniformes Cyprinidae Notropis rupestris
Cypriniformes Cyprinidae Notropis sabinae
Cypriniformes Cyprinidae Notropis scabriceps
Cypriniformes Cyprinidae Notropis scepticus
Cypriniformes Cyprinidae Notropis semperasper
Cypriniformes Cyprinidae Notropis shumardi
Cypriniformes Cyprinidae Notropis simus
Cypriniformes Cyprinidae Notropis spectrunculus
Cypriniformes Cyprinidae Notropis stilbius
Cypriniformes Cyprinidae Notropis stramineus
Cypriniformes Cyprinidae Notropis suttkusi
Cypriniformes Cyprinidae Notropis telescopus
Cypriniformes Cyprinidae Notropis texanus
Cypriniformes Cyprinidae Notropis topeka
Cypriniformes Cyprinidae Notropis uranoscopus
Cypriniformes Cyprinidae Notropis volucellus
Cypriniformes Cyprinidae Notropis wickliffi
Cypriniformes Cyprinidae Notropis xaenocephalus
Cypriniformes Cyprinidae Opsopoeodus emiliae
Cypriniformes Cyprinidae Oregonichthys crameri
Cypriniformes Cyprinidae Oregonichthys kalawatseti
Cypriniformes Cyprinidae Orthodon microlepidotus
Cypriniformes Cyprinidae Phenacobius catostomus
Cypriniformes Cyprinidae Phenacobius crassilabrum
Cypriniformes Cyprinidae Phenacobius mirabilis
Cypriniformes Cyprinidae Phenacobius teretulus
Cypriniformes Cyprinidae Phenacobius uranops
Cypriniformes Cyprinidae Phoxinus cumberlandensis
Cypriniformes Cyprinidae Phoxinus eos
Cypriniformes Cyprinidae Phoxinus erythrogaster
Cypriniformes Cyprinidae Phoxinus neogaeus
Cypriniformes Cyprinidae Phoxinus oreas
Cypriniformes Cyprinidae Phoxinus saylori
Cypriniformes Cyprinidae Phoxinus tennesseensis
Cypriniformes Cyprinidae Pimephales notatus
Cypriniformes Cyprinidae Pimephales promelas
Cypriniformes Cyprinidae Pimephales tenellus
Cypriniformes Cyprinidae Pimephales vigilax
Ozark shiner
carmine shiner
peppered shiner
coastal shiner
silver shiner
chub shiner
swallowtail shiner
Yazoo shiner
rosyface shiner
saffron shiner
bedrock shiner
Sabine shiner
New River shiner
sandbar shiner
roughhead shiner
silverband shiner
bluntnose shiner
mirror shiner
silverstripe shiner
sand shiner
rocky shiner
telescope shiner
weed shiner
Topeka shiner
skygazer shiner
mimic shiner
channel shiner
Coosa shiner
pugnose minnow
Oregon chub
Umpqua chub
Sacramento blackfish
riffle minnow
fatlips minnow
suckermouth minnow
Kanawha minnow
stargazing minnow
blackside dace
northern redbelly dace
southern redbelly dace
finescale dace
mountain redbelly dace
laurel dace
Tennessee dace
bluntnose minnow
fathead minnow
slim minnow
bullhead minnow
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ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cypriniformes Cyprinidae Plagopterus argentissimus
Cypriniformes Cyprinidae Platygobio gracilis
Cypriniformes Cyprinidae Pogonichthys ciscoides
Cypriniformes Cyprinidae Pogonichthys macrolepidotus
Cypriniformes Cyprinidae Reronotropis euryzonus
Cypriniformes Cyprinidae Reronotropis grandipinnis
Cypriniformes Cyprinidae Reronotropis hubbsi
Cypriniformes Cyprinidae Reronotropis hypselopterus
Cypriniformes Cyprinidae Reronotropis merlini
Cypriniformes Cyprinidae Reronotropis signipinnis
Cypriniformes Cyprinidae Reronotropis welaka
Cypriniformes Cyprinidae Rychocheilus grandis
Cypriniformes Cyprinidae Rychocheilus lucius
Cypriniformes Cyprinidae Rychocheilus oregonensis
Cypriniformes Cyprinidae Rychocheilus umpquae
Cypriniformes Cyprinidae Relictus solitarius
Cypriniformes Cyprinidae Rhinichthys atratulus
Cypriniformes Cyprinidae Rhinichthys cataractae
Cypriniformes Cyprinidae Rhinichthys cobitis
Cypriniformes Cyprinidae Rhinichthys deacon;
Cypriniformes Cyprinidae Rhinichthys evermanni
Cypriniformes Cyprinidae Rhinichthys falcatus
Cypriniformes Cyprinidae Rhinichthys obtusus
Cypriniformes Cyprinidae Rhinichthys osculus
Cypriniformes Cyprinidae Rhinichthys umatilla
Cypriniformes Cyprinidae Rhodeus sericeus
Cypriniformes Cyprinidae Richardsonius balteatus
Cypriniformes Cyprinidae Richardsonius egregius
Cypriniformes Cyprinidae Scardinius erythrophthalmus
Cypriniformes Cyprinidae Semotilus atromaculatus
Cypriniformes Cyprinidae Semotilus corpora/is
Cypriniformes Cyprinidae Semotilus lumbee
Cypriniformes Cyprinidae Semotilus thoreauianus
Cypriniformes Cyprinidae Snyderichthys cope;
Cypriniformes Cyprinidae Tinea tinea
Cypriniformes Catostomidae Carp/odes carpio
Cypriniformes Catostomidae Carp/odes cyprinus
Cypriniformes Catostomidae Carp/odes velifer
Cypriniformes Catostomidae Catostomus ardens
Cypriniformes Catostomidae Catostomus bernardini
Cypriniformes Catostomidae Catostomus catostomus
Cypriniformes Catostomidae Catostomus clarkii
Cypriniformes Catostomidae Catostomus columbianus
Cypriniformes Catostomidae Catostomus commersonii
Cypriniformes Catostomidae Catostomus discobolus
Cypriniformes Catostomidae Catostomus fumeiventris
Cypriniformes Catostomidae Catostomus insignis
Cypriniformes Catostomidae Catostomus latipinnis
woundfin
flathead chub
Clear Lake splittail
splittail
broadstripe shiner
Apalachee shiner
bluehead shiner
sailfin shiner
orangetail shiner
flagfin shiner
bluenose shiner
Sacramento pikeminnow
Colorado pikeminnow
northern pikeminnow
Umpqua pikeminnow
relict dace
eastern blacknose dace
longnose dace
loach minnow
Las Vegas dace
Umpqua dace
leopard dace
western blacknose dace
speckled dace
Umatilla dace
bitterling
redside shiner
Lahontan redside
rudd
creek chub
fallfish
sandhills chub
Dixie chub
leatherside chub
tench
river carpsucker
quillback
highfin carpsucker
Utah sucker
Yaqui sucker
longnose sucker
desert sucker
bridgelip sucker
white sucker
bluehead sucker
Owens sucker
Sonora sucker
flannelmouth sucker
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-10
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Cypriniformes
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomidae
Catostomus macrocheilus
Catostomus microps
Catostomus occidentalis
Catostomus platyrhynchus
Catostomus plebeius
Catostomus rimiculus
Catostomus santaanae
Catostomus snyderi
Catostomus tahoensis
Catostomus warnerensis
Chasmistes brevirostris
Chasmistes cujus
Chasmistes liorus
Chasmistes muriei
Cycleptus elongatus
Cycleptus meridionalis
Deltistes luxatus
Erimyzon oblongus
Erimyzon sucetta
Erimyzon tenuis
Hypentelium etowanum
Hypentelium nigricans
Hypentelium roanokense
Ictiobus bubalus
Ictiobus cyprinellus
Ictiobus niger
Minytrema melanops
Moxostoma anisurum
Moxostoma ariommum
Moxostoma austrinum
Moxostoma breviceps
Moxostoma carinatum
Moxostoma cervinum
Moxostoma collapsum
Moxostoma congestum
Moxostoma duquesnei
Moxostoma erythrurum
Moxostoma lacerum
Moxostoma lachneri
Moxostoma macrolepidotum
Moxostoma pappillosum
Moxostoma pisolabrum
Moxostoma poecilurum
Moxostoma robustum
Moxostoma rupiscartes
Moxostoma valenciennesi
Thoburnia atripinnis
Thoburnia hamiltoni
largescale sucker
Modoc sucker
Sacramento sucker
mountain sucker
Rio Grande sucker
Klamath smallscale sucker
Santa Ana sucker
Klamath largescale sucker
Tahoe sucker
Warner sucker
shortnose sucker
cui-ui
June sucker
Snake River sucker
blue sucker
southeastern blue sucker
Lost River sucker
creek chubsucker
lake chubsucker
sharpfin chubsucker
Alabama hog sucker
northern hog sucker
Roanoke hog sucker
smallmouth buffalo
bigmouth buffalo
black buffalo
spotted sucker
silver redhorse
bigeye jumprock
Mexican redhorse
smallmouth redhorse
river redhorse
blacktip jumprock
notchlip redhorse
gray redhorse
black redhorse
golden redhorse
harelip sucker
greater jumprock
shorthead redhorse
V-lip redhorse
pealip redhorse
blacktail redhorse
robust redhorse
striped jumprock
greater redhorse
blackfin sucker
rustyside sucker
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-11
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cypriniformes
Cypriniformes
Cypriniformes
Characiformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Catostomidae
Catostomidae
Cobitidae
Characidae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Ictaluridae
Clariidae
Ariidae
Doradidae
Callichthyidae
Loricariidae
Thoburnia rhothoeca
Xyrauchen texanus
Misgurnus anguillicaudatus
Astyanax mexicanus
Ameiurus brunneus
Ameiurus catus
Ameiurus melas
Ameiurus natalis
Ameiurus nebulosus
Ameiurus platycephalus
Ameiurus serracanthus
Ictalurus furcatus
Ictalurus lupus
Ictalurus price!
Ictalurus punctatus
Noturus albater
Noturus baileyi
Noturus elegans
Noturus eleutherus
Noturus exilis
Noturus flavater
Noturus flavipinnis
Noturus flavus
Noturus funebris
Noturus furiosus
Noturus gilbert!
Noturus gyrinus
Noturus hildebrandi
Noturus insignis
Noturus lachneri
Noturus leptacanthus
Noturus miurus
Noturus munitus
Noturus nocturnus
Noturus phaeus
Noturus placidus
Noturus stanauli
Noturus stigmosus
Noturus taylori
Noturus trautmani
Pylodictis olivaris
Satan eurystomus
Trogloglanis pattersoni
Clarias batrachus
Ariopsis felis
Platydoras armatulus
Hoplosternum littorale
Hypostomus plecostomus
torrent sucker
razorback sucker
oriental weatherfish
Mexican tetra
snail bullhead
white catfish
black bullhead
yellow bullhead
brown bullhead
flat bullhead
spotted bullhead
blue catfish
headwater catfish
Yaqui catfish
channel catfish
Ozark madtom
smoky madtom
elegant madtom
mountain madtom
slender madtom
checkered madtom
yellowfin madtom
stonecat
black madtom
Carolina madtom
orangefin madtom
tadpole madtom
least madtom
margined madtom
Ouachita madtom
speckled madtom
brindled madtom
frecklebelly madtom
freckled madtom
brown madtom
Neosho madtom
pygmy madtom
northern madtom
Caddo madtom
Scioto madtom
flathead catfish
widemouth blindcat
toothless blindcat
walking catfish
hardhead catfish
southern striped Raphael
brown hoplo
suckermouth catfish
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-12
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Siluriformes
Siluriformes
Siluriformes
Siluriformes
Esociformes
Esociformes
Esociformes
Esociformes
Esociformes
Esociformes
Esociformes
Esociformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Loricariidae Rerygoplichthys anisitsi
Loricariidae Rerygoplichthys disjunctivus
Loricariidae Rerygoplichthys multiradiatus
Loricariidae Rerygoplichthys pardalis
Esocidae Esox americanus
Esocidae Esox lucius
Esocidae Esox masquinongy
Esocidae Esox niger
Umbridae Dallia pectoralis
Umbridae Novumbra hubbsi
Umbridae Umbra limi
Umbridae Umbra pygmaea
Osmeridae Hypomesus nipponensis
Osmeridae Hypomesus olidus
Osmeridae Hypomesus pretiosus
Osmeridae Hypomesus transpacificus
Osmeridae Osmerus mordax
Osmeridae Spirinchus thaleichthys
Osmeridae Thaleichthys pacificus
Salmonidae Coregonus artedi
Salmonidae Coregonus autumnalis
Salmonidae Coregonus clupeaformis
Salmonidae Coregonus hoyi
Salmonidae Coregonus johannae
Salmonidae Coregonus kiyi
Salmonidae Coregonus laurettae
Salmonidae Coregonus nasus
Salmonidae Coregonus nigripinnis
Salmonidae Coregonus pidschian
Salmonidae Coregonus reighardi
Salmonidae Coregonus sardinella
Salmonidae Coregonus zenithicus
Salmonidae Oncorhynchus clarkii
Salmonidae Oncorhynchus gilae
Salmonidae Oncorhynchus gorbuscha
Salmonidae Oncorhynchus keta
Salmonidae Oncorhynchus kisutch
Salmonidae Oncorhynchus mykiss
Salmonidae Oncorhynchus nerka
Salmonidae Oncorhynchus tshawytscha
Salmonidae Prosopium abyssicola
Salmonidae Prosopium coulterii
Salmonidae Prosopium cylindraceum
Salmonidae Prosopium gemmifer
Salmonidae Prosopium spilonotus
Salmonidae Prosopium williamsoni
Salmonidae Salmo salar
Salmonidae Salmo trutta
southern sailfin catfish
vermiculated sailfin catfish
Orinoco sailfin catfish
Amazon sailfin catfish
redfin pickerel
northern pike
muskellunge
chain pickerel
Alaska blackfish
Olympic mudminnow
central mudminnow
eastern mudminnow
wakasagi
pond smelt
surf smelt
delta smelt
rainbow smelt
longfin smelt
eulachon
Cisco
Arctic Cisco
lake whitefish
bloater
deepwater Cisco
kiyi
Bering Cisco
broad whitefish
blackfin Cisco
humpback whitefish
shortnose Cisco
least Cisco
shortjaw Cisco
cutthroat trout
Gila trout
pink salmon
chum salmon
coho salmon
rainbow trout
sockeye salmon
Chinook salmon
Bear Lake whitefish
pygmy whitefish
round whitefish
Bonneville Cisco
Bonneville whitefish
mountain whitefish
Atlantic salmon
brown trout
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-13
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Salmoniformes
Percopsiformes
Percopsiformes
Percopsiformes
Percopsiformes
Percopsiformes
Percopsiformes
Percopsiformes
Percopsiformes
Percopsiformes
Gadiformes
Gadiformes
Mugiliformes
Mugiliformes
Mugiliformes
Atheriniformes
Atheriniformes
Atheriniformes
Atheriniformes
Atheriniformes
Beloniformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Salmonidae
Salmonidae
Salmonidae
Salmonidae
Salmonidae
Salmonidae
Salmonidae
Percopsidae
Percopsidae
Aphredoderidae
Amblyopsidae
Amblyopsidae
Amblyopsidae
Amblyopsidae
Amblyopsidae
Amblyopsidae
Gadidae
Gadidae
Mugilidae
Mugilidae
Mugilidae
Atherinopsidae
Atherinopsidae
Atherinopsidae
Atherinopsidae
Atherinopsidae
Belonidae
Aplocheilidae
Aplocheilidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Salvelinus alpinus
Salvelinus confluentus
Salvelinus fontinalis
Salvelinus malma
Salvelinus namaycush
Stenodus leucichthys
Thymallus arcticus
Percopsis omiscomaycus
Percopsis transmontana
Aphredoderus sayanus
Amblyopsis rosae
Amblyopsis spelaea
Chologaster cornuta
Forbesichthys agassizii
Speoplatyrhinus poulsoni
Typhlichthys subterraneus
Lota lota
Microgadus tomcod
Agonostomus monticola
Mugil cephalus
Mugil curema
Labidesthes sicculus
Membras martinica
Menidia audens
Menidia beryllina
Menidia extensa
Strongylura marina
Rivulus hartii
Rivulus marmoratus
Fundulus albolineatus
Fundulus bifax
Fundulus blairae
Fundulus catenatus
Fundulus chrysotus
Fundulus cingulatus
Fundulus confluentus
Fundulus diaphanus
Fundulus dispar
Fundulus escambiae
Fundulus euryzonus
Fundulus grandis
Fundulus heteroclitus
Fundulus jenkinsi
Fundulus julisia
Fundulus kansae
Fundulus lineolatus
Fundulus luciae
Arctic char
bull trout
brook trout
Dolly Varden
lake trout
inconnu
Arctic grayling
trout-perch
sand roller
pirate perch
Ozark cavefish
northern cavefish
swampfish
spring cavefish
Alabama cavefish
southern cavefish
burbot
Atlantic tomcod
mountain mullet
striped mullet
white mullet
brook silverside
rough silverside
Mississippi silverside
inland silverside
Waccamaw silverside
Atlantic needlefish
giant rivulus
mangrove rivulus
whiteline topminnow
stippled studfish
western starhead
topminnow
northern studfish
golden topminnow
banded topminnow
marsh killifish
banded killifish
starhead topminnow
russetfin topminnow
broadstripe topminnow
Gulf killifish
mummichog
saltmarsh topminnow
Barrens topminnow
northern plains killifish
lined topminnow
spotfin killifish
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-14
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Fundulidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Poeciliidae
Goodeidae
Goodeidae
Goodeidae
Goodeidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Fundulus notatus
Fundulus nottii
Fundulus olivaceus
Fundulus parvipinnis
Fundulus pulvereus
Fundulus rathbuni
Fundulus rubrifrons
Fundulus sciadicus
Fundulus seminolis
Fundulus stellifer
Fundulus waccamensis
Fundulus zebrinus
Leptolucania ommata
Lucania goodei
Lucania pan/a
Belonesox belizanus
Gambusia affinis
Gambusia amistadensis
Gambusia gaigei
Gambusia geiseri
Gambusia georgei
Gambusia heterochir
Gambusia holbrooki
Gambusia nobilis
Gambusia rhizophorae
Gambusia senilis
Gambusia speciosa
Heterandria formosa
Poecilia formosa
Poecilia latipinna
Poecilia mexicana
Poecilia reticulata
Poecilia sphenops
Poeciliopsis gracilis
Poeciliopsis occidentalis
Xiphophorus hellerii
Xiphophorus maculatus
Xiphophorus variatus
Crenichthys baileyi
Crenichthys nevadae
Empetrichthys latos
Empetrichthys merriami
Cyprinodon arcuatus
Cyprinodon bovinus
Cyprinodon diabolis
Cyprinodon elegans
Cyprinodon eremus
Cyprinodon eximius
blackstripe topminnow
bayou topminnow
blackspotted topminnow
Guadalupe cardinalfish
bayou killifish
speckled killifish
redface topminnow
plains topminnow
Seminole killifish
southern studfish
Waccamaw killifish
plains killifish
pygmy killifish
bluefin killifish
rainwater killifish
pike killifish
western mosquitofish
Amistad gambusia
Big Bend gambusia
largespring gambusia
San Marcos gambusia
Clear Creek gambusia
eastern mosquitofish
Pecos gambusia
mangrove gambusia
blotched gambusia
Tex-Mex gambusia
least killifish
Amazon molly
sailfin molly
shortfin molly
guppy
Mexican molly
porthole livebearer
Gila topminnow
green swordtail
southern platyfish
variable platyfish
White River springfish
Railroad Valley springfish
Pahrump poolfish
Ash Meadows poolfish
Santa Cruz pupfish
Leon Springs pupfish
Devils Hole pupfish
Comanche Springs pupfish
Sonoyta pupfish
Conchos pupfish
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-15
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Cyprinodontiformes
Gasterosteiformes
Gasterosteiformes
Gasterosteiformes
Gasterosteiformes
Gasterosteiformes
Gasterosteiformes
Synbranchiformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Scorpaeniformes
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Cyprinodontidae
Gasterosteidae
Gasterosteidae
Gasterosteidae
Gasterosteidae
Syngnathidae
Syngnathidae
Syn branch idae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cottidae
Cyprinodon macularius
Cyprinodon nevadensis
Cyprinodon pecosensis
Cyprinodon radiosus
Cyprinodon rubrofluviatilis
Cyprinodon salinus
Cyprinodon tularosa
Cyprinodon variegatus
Jordanella floridae
Apeltes quadracus
Culaea inconstans
Gasterosteus aculeatus
Pungitius pungitius
Microphis brachyurus
Syngnathus scovelli
Monopterus alb us
Clinocottus acuticeps
Cottus aleuticus
Cottus asper
Cottus asperrimus
Cottus baileyi
Cottus bairdii
Cottus beldingii
Cottus bendirei
Cottus caeruleomentum
Cottus carolinae
Cottus cognatus
Cottus confusus
Cottus echinatus
Cottus extensus
Cottus girardi
Cottus greenei
Cottus gulosus
Cottus hubbsi
Cottus hypselurus
Cottus klamathensis
Cottus leiopomus
Cottus marginatus
Cottus paulus
Cottus perplexus
Cottus pitensis
Cottus princeps
Cottus rhotheus
Cottus rice;
Cottus tenuis
Leptocottus armatus
Myoxocephalus quadricornis
Myoxocephalus thompsonii
desert pupfish
Amargosa pupfish
Pecos pupfish
Owens pupfish
Red River pupfish
Salt Creek pupfish
White Sands pupfish
sheepshead minnow
flagfish
fourspine stickleback
brook stickleback
espinocho
ninespine stickleback
opossum pipefish
Gulf pipefish
Asian swamp eel
sharpnose sculpin
coastrange sculpin
prickly sculpin
rough sculpin
black sculpin
mottled sculpin
Paiute sculpin
Malheur sculpin
Blue Ridge sculpin
banded sculpin
slimy sculpin
shorthead sculpin
Utah Lake sculpin
Bear Lake sculpin
Potomac sculpin
Shoshone sculpin
riffle sculpin
Columbia sculpin
Ozark sculpin
marbled sculpin
Wood River sculpin
margined sculpin
pygmy sculpin
reticulate sculpin
Pit sculpin
Klamath Lake sculpin
torrent sculpin
spoonhead sculpin
slender sculpin
Pacific staghorn sculpin
fourhorn sculpin
deepwater sculpin
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-16
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Centropomidae
Centropomidae
Centropomidae
Centropomidae
Moronidae
Moronidae
Moronidae
Moronidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Centrarchidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Centropomus ensiferus
Centropomus parallelus
Centropomus pectinatus
Centropomus undecimalis
Morone americana
Morone chrysops
Morone mississippiensis
Morone saxatilis
Acantharchus pomotis
Ambloplites ariommus
Ambloplites cavifrons
Ambloplites constellatus
Ambloplites rupestris
Archoplites interruptus
Centrarchus macropterus
Enneacanthus chaetodon
Enneacanthus gloriosus
Enneacanthus obesus
Lepomis auritus
Lepomis cyanellus
Lepomis gibbosus
Lepomis gulosus
Lepomis humilis
Lepomis macrochirus
Lepomis marginatus
Lepomis megalotis
Lepomis microlophus
Lepomis miniatus
Lepomis punctatus
Lepomis symmetricus
Micropterus cataractae
Micropterus coosae
Micropterus dolomieu
Micropterus notius
Micropterus punctulatus
Micropterus salmoides
Micropterus treculii
Pomoxis annularis
Pomoxis nigromaculatus
Ammocrypta beanii
Ammocrypta bifascia
Ammocrypta clara
Ammocrypta meridiana
Ammocrypta pellucida
Ammocrypta vivax
Crystal/aria asprella
Etheostoma acuticeps
Etheostoma aquali
swordspine snook
smallscale fat snook
tarpon snook
common snook
white perch
white bass
yellow bass
striped bass
mud sunfish
shadow bass
Roanoke bass
Ozark bass
rock bass
Sacramento perch
flier
blackbanded sunfish
bluespotted sunfish
banded sunfish
redbreast sunfish
green sunfish
pumpkinseed
warmouth
orangespotted sunfish
bluegill
dollar sunfish
longear sunfish
redear sunfish
redspotted sunfish
spotted sunfish
bantam sunfish
shoal bass
redeye bass
smallmouth bass
Suwannee bass
spotted bass
largemouth bass
Guadalupe bass
white crappie
black crappie
naked sand darter
Florida sand darter
western sand darter
southern sand darter
eastern sand darter
scaly sand darter
crystal darter
sharphead darter
coppercheek darter
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-17
ORDER
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
FAMILY
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
SCIENTIFIC NAME
Etheostoma artesiae
Etheostoma asprigene
Etheostoma baileyi
Etheostoma barbouri
Etheostoma barrenense
Etheostoma basilare
Etheostoma bellator
Etheostoma bellum
Etheostoma bison
Etheostoma blennioides
Etheostoma blennius
Etheostoma boschungi
Etheostoma brevirostrum
Etheostoma burn
Etheostoma caeruleum
Etheostoma camurum
Etheostoma cervus
Etheostoma chermocki
Etheostoma chienense
Etheostoma chlorobranchium
Etheostoma chlorosoma
Etheostoma chuckwachatte
Etheostoma cinereum
Etheostoma collettei
Etheostoma collis
Etheostoma colorosum
Etheostoma coosae
Etheostoma corona
Etheostoma cragini
Etheostoma crossopterum
Etheostoma davisoni
Etheostoma denoncourti
Etheostoma derivativum
Etheostoma ditrema
Etheostoma douglasi
Etheostoma duryi
Etheostoma edwini
Etheostoma etnieri
Etheostoma etowahae
Etheostoma euzonum
Etheostoma exile
Etheostoma flabellare
Etheostoma flavum
Etheostoma fonticola
Etheostoma forbesi
Etheostoma fragi
Etheostoma fricksium
Etheostoma fusiforme
COMMON NAME
redspot darter
mud darter
emerald darter
teardrop darter
splendid darter
corrugated darter
Warrior darter
orangefin darter
Buffalo darter
greenside darter
blenny darter
slackwater darter
holiday darter
brook darter
rainbow darter
bluebreast darter
Chickasaw darter
vermilion darter
relict darter
greenfin darter
bluntnose darter
lipstick darter
ashy darter
Creole darter
Carolina darter
coastal darter
Coosa darter
crown darter
Arkansas darter
fringed darter
Choctawhatchee darter
golden darter
stone darter
coldwater darter
Tuskaloosa darter
blackside snubnose darter
brown darter
cherry darter
Etowah darter
Arkansas saddled darter
Iowa darter
fantail darter
saffron darter
fountain darter
Barrens darter
Strawberry darter
Savannah darter
swamp darter
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-18
ORDER
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
FAMILY
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
SCIENTIFIC NAME
Etheostoma gracile
Etheostoma grahami
Etheostoma gutselli
Etheostoma histrio
Etheostoma hopkinsi
Etheostoma inscriptum
Etheostoma jessiae
Etheostoma jordani
Etheostoma juliae
Etheostoma kanawhae
Etheostoma kantuckeense
Etheostoma kennicotti
Etheostoma lachneri
Etheostoma lawrencei
Etheostoma lepidum
Etheostoma longimanum
Etheostoma luteovinctum
Etheostoma lynceum
Etheostoma maculatum
Etheostoma mariae
Etheostoma microlepidum
Etheostoma microperca
Etheostoma moorei
Etheostoma neopterum
Etheostoma nianguae
Etheostoma nigripinne
Etheostoma nigrum
Etheostoma nuchale
Etheostoma obeyense
Etheostoma okaloosae
Etheostoma olivaceum
Etheostoma olmstedi
Etheostoma oophylax
Etheostoma osburni
Etheostoma pallididorsum
Etheostoma parvipinne
Etheostoma percnurum
Etheostoma perlongum
Etheostoma phytophilum
Etheostoma podostemone
Etheostoma proeliare
Etheostoma pseudovulatum
Etheostoma punctulatum
Etheostoma pyrrhogaster
Etheostoma radiosum
Etheostoma rafinesquei
Etheostoma ramseyi
Etheostoma raneyi
COMMON NAME
slough darter
Rio Grande darter
Tuckasegee darter
harlequin darter
Christmas darter
turquoise darter
blueside darter
greenbreast darter
yoke darter
Kanawha darter
Highland Rim darter
stripetail darter
Tombigbee darter
headwater darter
greenthroat darter
longfin darter
redband darter
brighteye darter
spotted darter
pinewoods darter
smallscale darter
least darter
yellow/cheek darter
lollypop darter
Niangua darter
blackfin darter
johnny darter
watercress darter
barcheek darter
Okaloosa darter
sooty darter
tessellated darter
guardian darter
candy darter
paleback darter
goldstripe darter
duskytail darter
Waccamaw darter
rush darter
riverweed darter
cypress darter
egg-mimic darter
stippled darter
firebelly darter
orangebelly darter
Kentucky darter
Alabama darter
Yazoo darter
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-19
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Etheostoma rubrum
Etheostoma rufilineatum
Etheostoma rupestre
Etheostoma sagitta
Etheostoma sanguifluum
Etheostoma scotti
Etheostoma sellare
Etheostoma serrifer
Etheostoma simoterum
Etheostoma smith!
Etheostoma spectabile
Etheostoma squamiceps
Etheostoma stigmaeum
Etheostoma striatulum
Etheostoma susanae
Etheostoma swaini
Etheostoma swannanoa
Etheostoma tallapoosae
Etheostoma tecumsehi
Etheostoma tetrazonum
Etheostoma thalassinum
Etheostoma tippecanoe
Etheostoma trisella
Etheostoma tuscumbia
Etheostoma uniporum
Etheostoma variatum
Etheostoma virgatum
Etheostoma vitreum
Etheostoma vulneratum
Etheostoma wapiti
Etheostoma whipplei
Etheostoma zonale
Etheostoma zonifer
Etheostoma zonistium
Gymnocephalus cernuus
Perca flavescens
Percina antesella
Percina aurantiaca
Percina aurolineata
Percina aurora
Percina austroperca
Percina brevicauda
Percina burtoni
Percina caprodes
Percina carbonaria
Percina copelandi
Percina crassa
Percina cymatotaenia
bayou darter
redline darter
rock darter
arrow darter
bloodfin darter
Cherokee darter
Maryland darter
sawcheek darter
snubnose darter
slabrock darter
orangethroat darter
spottail darter
speckled darter
striated darter
Cumberland darter
Gulf darter
Swannanoa darter
Tallapoosa darter
Shawnee darter
Missouri saddled darter
seagreen darter
Tippecanoe darter
trispot darter
Tuscumbia darter
current darter
variegate darter
striped darter
glassy darter
wounded darter
boulder darter
redfin darter
banded darter
backwater darter
bandfin darter
ruffe
yellow perch
amber darter
tangerine darter
goldline darter
pearl darter
southern logperch
coal darter
blotchside logperch
logperch
Texas logperch
channel darter
Piedmont darter
bluestripe darter
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-20
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Percidae
Lutjanidae
Gerreidae
Gerreidae
Gerreidae
Haemulidae
Sparidae
Sparidae
Sciaenidae
Sciaenidae
Sciaenidae
Sciaenidae
Sciaenidae
Sciaenidae
Sciaenidae
Sciaenidae
Elassomatidae
Elassomatidae
Percina evides
Percina fulvitaenia
Percina gymnocephala
Percina jenkinsi
Percina kathae
Percina lenticula
Percina macrocephala
Percina macrolepida
Percina maculata
Percina nasuta
Percina nevisense
Percina nigrofasciata
Percina notogramma
Percina oxyrhynchus
Percina palmaris
Percina pantherina
Percina peltata
Percina phoxocephala
Percina rex
Percina roanoka
Percina sciera
Percina shumardi
Percina squamata
Percina stictogaster
Percina suttkusi
Percina tanasi
Percina uranidea
Percina vigil
Sander canadensis
Sander lucioperca
Sander vitreus
Lutjanus griseus
Diapterus auratus
Eucinostomus harengulus
Eugenes plumieri
Orthopristis chrysoptera
Archosargus probatocephalus
Lagodon rhomboides
Aplodinotus grunniens
Bairdiella chrysoura
Bairdiella icistia
Cynoscion nebulosus
Cynoscion xanthulus
Leiostomus xanthurus
Micropogonias undulatus
Sciaenops ocellatus
Elassoma alabamae
Elassoma boehlkei
gilt darter
Ozark logperch
Appalachia darter
Conasauga logperch
Mobile logperch
freckled darter
longhead darter
bigscale logperch
blackside darter
longnose darter
chainback darter
blackbanded darter
stripeback darter
sharpnose darter
bronze darter
leopard darter
shield darter
slenderhead darter
Roanoke logperch
Roanoke darter
dusky darter
river darter
olive darter
frecklebelly darter
Gulf logperch
snail darter
stargazing darter
saddleback darter
sauger
zander
walleye
gray snapper
Irish pompano
tidewater mojarra
striped mojarra
pigfish
sheepshead
pinfish
freshwater drum
silver perch
bairdiella
spotted seatrout
orangemouth corvina
spot
Atlantic croaker
red drum
spring pygmy sunfish
Carolina pygmy sunfish
-------
National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-21
ORDER
FAMILY
SCIENTIFIC NAME
COMMON NAME
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Elassomatidae
Elassomatidae
Elassomatidae
Elassomatidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Cichlidae
Embiotocidae
Embiotocidae
Eleotridae
Eleotridae
Eleotridae
Eleotridae
Eleotridae
Eleotridae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Gobiidae
Elassoma everglade!
Elassoma okatie
Elassoma okefenokee
Elassoma zonatum
Astronotus ocellatus
Cichla ocellaris
Cichlasoma bimaculatum
Cichlasoma citrinellum
Cichlasoma cyanoguttatum
Cichlasoma managuense
Cichlasoma meeki
Cichlasoma nigrofasciatum
Cichlasoma octofasciatum
Cichlasoma salvini
Cichlasoma urophthalmus
Geophagus surinamensis
Hemichromis letourneuxi
Hems severus
Oreochromis aureus
Oreochromis mossambicus
Oreochromis niloticus
Oreochromis urolepis
Sarotherodon melanotheron
Tilapia mariae
Tilapia zillii
Cymatogaster aggregate
Hysterocarpus traskii
Dormitator maculatus
Eleotris amblyopsis
Eleotris perniger
Eleotris picta
Gobiomorus dormitor
Guavina guavina
Acanthogobius flavimanus
Awaous banana
Clevelandia ios
Ctenogobius boleosoma
Ctenogobius claytonii
Ctenogobius fasciatus
Ctenogobius pseudofasciatus
Ctenogobius shufeldti
Eucyclogobius newberryi
Gillichthys mirabilis
Gobioides broussonetii
Gobiosoma bosc
Lophogobius cyprinoides
Microgobius gulosus
Neogobius melanostomus
Everglades pygmy sunfish
bluebarred pygmy sunfish
Okefenokee pygmy sunfish
banded pygmy sunfish
oscar
butterfly peacock bass
black acara
midas cichlid
Rio Grande cichlid
jaguar guapote
firemouth cichlid
convict cichlid
Jack Dempsey
yellow/belly cichlid
Mayan cichlid
redstriped eartheater
African jewelfish
banded cichlid
blue tilapia
Mozambique tilapia
Nile tilapia
Wami tilapia
blackchin tilapia
spotted tilapia
red belly tilapia
shiner perch
tule perch
fat sleeper
largescaled spinycheek
smallscaled spinycheek
spotted sleeper
bigmouth sleeper
guavina
yellowfin goby
river goby
arrow goby
darter goby
Mexican goby
blotchcheek goby
slashcheek goby
freshwater goby
tidewater goby
longjaw mudsucker
violet goby
naked goby
crested goby
clown goby
round goby
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page D-22
ORDER
Perciformes
Perciformes
Perciformes
Perciformes
Perciformes
Pleuronectiformes
Pleuronectiformes
Pleuronectiformes
Pleuronectiformes
FAMILY
Gobiidae
Gobiidae
Gobiidae
Belontiidae
Channidae
Paralichthyidae
Paralichthyidae
Pleuronectidae
Achiridae
SCIENTIFIC NAME
Proterorhinus marmoratus
Tridentiger barbatus
Tridentiger bifasciatus
Trichopsis vittata
Channa marulius
Citharichthys spilopterus
Paralichthys lethostigma
Platichthys stellatus
Trinectes maculatus
COMMON NAME
tubenose goby
Shokihaze goby
shimofuri goby
croaking gourami
bullseye snakehead
bay whiff
southern flounder
starry flounder
hogchoker
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APPENDIX E
PPCP and PFC Samples at
Selected Urban Sites
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EPA's Office of Science and Technology (OST) within the Office of Water is
collaborating with the Office of Research and Development's National Exposure Research
Laboratory in Cincinnati, Ohio to conduct a study of contaminants of emerging concern (CECs)
within the framework of the National Rivers and Streams Assessment (NRSA). These CECs
include Pharmaceuticals and personal care products (PPCPs), along with perfluorinated
compounds (PFCs). This study involves collection of ambient water (water chemistry) samples
and fish tissue samples at about 150 urban river sites. These sites comprise a statistical subset
within the 1800 sites selected for NRSA sampling. The urban river sites were assigned to the
PPCP and PFC Study based on 5th order or greater Strahler stream order. The majority of
these sites will be boatable, but a few of them will be wadeable. PPCP and PFC water
and fish tissue samples need to be collected at the boatable and wadeable sites in this
subset of urban river locations to maintain the statistical integrity of the data.
PPCP Water Chemistry Samples
The water chemistry protocols for collection of PPCP water samples are identical to the
general water chemistry sample collection protocols for the NRSA water quality indicators. OST
will provide field teams with coolers and 500 ml (0.5 L) amber glass bottles for the PPCP water
samples. Water for the PPCP samples will be collected using the beaker provided for collection
of other water chemistry samples. Field teams will use river water from the beaker to rinse the
sample bottles and caps before filling each of the sample bottles completely with water from the
beaker to eliminate air from the bottle. After fastening the caps tightly on the sample bottles, the
field crews will place the samples in the cooler on wet ice. Field teams will collect two 500 ml
PPCP water samples at all the urban river sites (boatable and wadeable) except the
repeat urban river sites. At the repeat urban river sites, field teams will collect four 500
ml PPCP water samples during the first site visit only.
1. Collect the PPCP water samples mid-channel at the X-site (located via GPS). Samples
are taken mid-channel, at a depth of 0.5 meters or at mid-depth if the site is less than 1
meter deep.
2. Put on nitrile gloves. Avoid touching the inside of the container to prevent contamination.
Make sure not to handle sunscreen or other chemical contaminants until after the sample
is collected.
3. Pre-rinse the beaker with river water 3 times, discarding rinse water downstream. Hold
the container so the opening faces upstream. Collect the sample at a depth of 0.5
meters below the surface, with the beaker slightly angled as you pull it to the surface.
4. Rinse each PPCP sample bottle with a small amount of the sample water before filling
the sample bottle with water from the beaker.
5. Fill the two 500 ml amber glass bottles (or four 500 ml amber glass bottles during the first
visit at urban river repeat sites) using water from the beaker. After filling each sample
bottle completely to eliminate air from the bottle, fasten the cap firmly on the bottle. Make
sure that the label is complete and taped over with clear tape, and then place the sample
bottles in the PPCP water sample cooler on wet ice.
6. Water samples collected at the pre-selected urban PPCP sites must be shipped to
the EPA CINCINNATI lab ON MONDAYS THROUGH THURSDAYS. Do not send
PPCP water samples to the EPA Corvallis lab. Please follow the instructions
provided in the PPCP urban site water sample supply cooler.
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Page E-2
Please use the following special instructions for shipping PPCP water samples:
• PPCP water samples collected from the pre-selected urban river sites must be
shipped on wet ice to the EPA CINCINNATI LAB within 3 days of collection (for
delivery at the lab on the fourth day) using the pre-addressed FedEx airbill provided
in the PPCP water sample cooler.
• There is No Saturday, Sunday, or Federal Holiday Delivery at the EPA
CINCINNATI LAB, so PPCP water coolers must be shipped on Monday through
Thursday.
• IMPORTANT NOTE: PPCP water samples have a holding time of 4 days.
Therefore, PPCP water samples cannot be collected on Friday, held on wet ice over
the weekend, and shipped on Monday or they will exceed the sample holding time.
PFC Water Chemistry Samples
The first 4 steps of the procedures for collecting PFC water samples are identical to the
PPCP water sample collection procedures except that the PFC sample bottles are rinsed 3
times with water from the sampling beaker before filling them. However, there are four
important differences in the remaining procedures for collecting PFC water samples: PFC
samples contain 1 L of water (twice the volume of PPCP samples); water collected for PFC
analysis requires HOPE bottles; PFC samples are preserved with a nitric acid solution; and PFC
sample bottles are shipped in coolers at ambient temperatures with no ice. OST will provide
field teams with coolers and 1 L HOPE bottles for the PFC samples, along with the labels,
stickers, pre-addressed airbills, and other forms necessary for shipping the coolers. As for the
PPCP water samples, water for the PFC samples will be collected using the beaker provided for
collection of other water chemistry samples. Field teams will use river water from the beaker to
rinse the HOPE sample bottles 3 times before filling them almost to the top. Space is left at the
top of the bottle to add 5 ml of a nitric acid solution to preserve the samples. The filled HOPE
water bottles are placed in the cooler with no ice and shipped at the ambient temperature within
3 days to the laboratory designated for PFC analysis. Field teams will collect two 1 L PFC
water samples at all the urban river sites that are sampled in 2009 (both boatable and
wadeable urban sites that are 5th order or greater) except the repeat urban river sites. At
the repeat urban river sites, field teams will collect four 1 L PFC water samples during the
first site visit only.
1. Collect the PFC water samples mid-channel at the X-site (located via GPS). Samples
are taken mid-channel, at a depth of 0.5 meters or at mid-depth if the site is less than 1
meter deep.
2. Put on nitrile gloves. Avoid touching the inside of the container to prevent contamination.
Make sure not to handle sunscreen or other chemical contaminants until after the sample
is collected.
3. Pre-rinse the beaker with river water 3 times, discarding rinse water downstream. Hold
the container so the opening faces upstream. Collect the sample at a depth of 0.5
meters below the surface, with the beaker slightly angled as you pull it to the surface.
4. Rinse each PFC sample bottle 3 times with sample water before filling the sample bottle
with water from the beaker.
5. Fill the two 1 L HOPE bottles (or four 1 L HOPE bottles during the first visit at urban river
repeat sites) using water from the beaker. All sample bottles should only be filled to the
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Page E-3
top of the cylindrical portion of the bottle, leaving the shoulder and the neck empty to
allow room for the preservative (5 ml of 35% nitric acid) to be added.
6. Add 5 ml of 35% nitric acid, supplied in the premeasured ampoules, into the sample, cap
tightly, place an orange EP HNO3 sticker onto the water collection bottles to indicate that
the preservation agent has been added, and mix well. Only the contents of the
ampoule should be added to the sample - the opened ampoule should not be
placed into the sample bottles.
7. Make sure that the labels are complete and taped over with clear tape, and then place
the sample bottles in the RFC water sample cooler. Return sample bottles to the original
shipping container (coolers) and maintain at ambient temperature. Do not cool with wet
or dry ice.
8. Water samples collected at the pre-selected urban PFC sites must be shipped to
the designated lab ON MONDAYS THROUGH THURSDAYS. Do not send PFC water
samples to the EPA Corvallis lab. Please follow the instructions provided in the
PFC urban site water sample supply cooler.
Please use the following special instructions for shipping PFC water samples:
• PFC water samples collected from the pre-selected urban river sites must be
shipped at ambient temperature (without wet or dry ice) to the designated lab
within 3 days of collection (for delivery at the lab on the fourth day) using the pre-
addressed FedEx airbill provided in the PFC water sample cooler.
• There is No Saturday, Sunday, or Federal Holiday Delivery at the designated lab,
so PFC water coolers must be shipped on Monday through Thursday.
PPCP Fish Tissue
A single fish tissue composite sample will be collected at the approximately 150
designated urban river sites, except at the repeat urban river sites where two duplicate fish
tissue samples will be collected during the first site visit. The urban river fish composite
samples will provide tissue for analysis of PPCP chemicals and for analysis of the list of EMAP
chemicals. An important exception is that fish tissue samples will be collected at all
urban sites that are >5th order and wadeable. Field crews will use the protocols outlined in
Section 5.6 of the Field Operations Manual to collect the fish tissue samples at both the
beatable and wadeable urban river sites. These protocols are summarized below. Please note
in step 15 that fish tissue samples collected at urban river sites are shipped directly to the EPA
CINCINNATI LAB.
1. Put on clean nitrile gloves before handling the fish. Do not handle any food, drink,
sunscreen, or insect repellant until after the composite sample has been collected,
measured, and wrapped.
2. Rinse potential target species/individuals in ambient water to remove any foreign
material from the external surface and place in clean holding containers (e.g., livewells,
buckets). Return non-target fishes or small specimens to the river or stream.
3. Retain one predator species composite from each site. The composite must consist of
five fish of adequate size to provide a total of 500 grams of edible tissue for analysis
(refer to Table 5.6-2 for minimum species length guidelines). Select fish for each
composite based on the following criteria:
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• all are of the same species,
• all satisfy legal requirements of harvestable size (or weight) for the sampled river,
or at least be of consumable size if no legal harvest requirements are in effect,
• all are of similar size, so that the smallest individual in a composite is no less than
75% of the total length of the largest individual, and
• all are collected at the same time, i.e., collected as close to the same time as
possible, but no more than one week apart (Note: Individual fish may have to be
frozen until all fish to be included in the composite are available for delivery to the
designated laboratory).
Accurate taxonomic identification is essential in assuring and defining the organisms that
have been composited and submitted for analysis. Under no circumstances should
individuals from different species be used in a single composite sample.
4. Measure each individual fish to determine total body length. Measure total length of
each specimen in millimeters, from the anterior-most part of the fish to the tip of the
longest caudal fin ray (when the lobes of the caudal fin are depressed dorsoventrally).
5. Record sample number, species retained, specimen length, location collected, and
sampling date and time on the Fish Collection Form (Figure 5.5-1) in black ink. Mark
"URBAN" next to the site identification number at the top left of the fish form, and write
primary or duplicate in the comment section. Make sure the sample identification
numbers recorded on the collection form match those on the sample labels.
6. Sign and date the Fish Collection Form.
7. Remove each fish retained for analysis from the clean holding container(s) (e.g., livewell)
using clean nitrile gloves. Dispatch each fish using a clean wooden bat (or equivalent
wooden device).
8. Wrap each fish in extra heavy-duty aluminum foil, with the dull side in (foil provided by
EPA as solvent-rinsed, oven-baked sheets).
9. Prepare a Sample Identification Label for each sample, ensuring that the label
information matches the information recorded on the Fish Collection Form. Be sure to
include fish species and specimen length on each label.
10. Cut a length of food grade tubing (provided by EPA) that is long enough to contain each
individual fish and to allow extra length on each end to secure with cable ties. Place
each foil-wrapped specimen into the appropriate length of tubing. Seal each end of the
tubing with a plastic cable tie. Attach the fish sample label to the outside of the food-
grade tubing with clear tape and secure the label by taping around the entire fish (so that
tape sticks to tape).
11. Place all the wrapped fish in the composite from each site in a large plastic bag and seal
with another cable tie.
12. After each sample is packaged, place it immediately on dry ice for shipment. If samples
will be carried back to a laboratory or other facility to be frozen before shipment, wet ice
can be used to transport wrapped and bagged fish samples in the coolers to a laboratory
or other interim facility.
13. If possible, keep all (five) specimens designated for a particular composite in the same
shipping container (ice chest) for transport.
14. Samples may be stored temporarily on dry ice (replenishing the dry ice daily). You have
the option, depending on site logistics, of:
• shipping the samples packed on dry ice in sufficient quantities to keep samples
frozen for up to 48 hours (50 pounds are recommended), via priority overnight
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National Rivers and Streams Assessment
Field Operations Manual
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Date: April 2009
Page E-5
15.
delivery service (e.g., Federal Express), so that they arrive at the sample
preparation laboratory within less than 24 hours from the time of sample collection,
or
• freezing the samples within 24 hours of collection at <-20°C, and storing the frozen
samples until shipment within 3 weeks of sample collection (frozen samples will
subsequently be packed on dry ice and shipped to the sample preparation
laboratory via priority overnight delivery service).
Fish Tissue samples collected at the pre-selected urban PPCP sites must be
shipped to the EPA CINCINNATI lab. Do not send PPCP fish tissue samples to the
GLEC lab. Please follow the instructions provided in the PPCP site fish tissue
supply cooler. Be sure to include fish species and specimen lengths for all fish
tissue samples on the Sample Tracking Form (Figure E-1).
PPCP Contacts
For any questions about collecting, handling, or shipping PPCP water or fish tissue
samples, please contact Leanne Stahl in the Office of Science and Technology at EPA or Elaine
Snyder of Tetra Tech, Inc. using the information below.
Leanne Stahl
USEPA/OST (4305T)
1200 Pennsylvania Avenue, NW
Washington, DC 20460
(202) 566-0404 (phone)
(202) 566-0409 (fax)
stahl.leanne@epa.gov
Elaine Snyder
Tetra Tech, Inc.
400 Red Brook Blvd., Suite 200
Owings Mills, MD 21117
(410)356-8993 (phone)
(410)356-9005 (fax)
Blaine.Snyder@tetratech.com
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National Rivers and Streams Assessment
Field Operations Manual
Final Manual
Date: April 2009
Page E-1
Please UK Bie tab button to navigate ttnugn W6 fwra.
TRACKING (BATCHED AND RETAINED SAMPLES)
National Rivers and Streams Assessment
Choose One: EJ BATCHED* SAMPLES Q RETAINED** SAMPLES
Do not combine both BATCHED arnJ RETAINED samples on the same foirn - ukase complete a separate farm.
Sender:
Sender Phone:
State of Site Location:
Team:
Date Shipped:
Shippec
By:
Airbill:
For Retained Samples enter
Holding Facility and Address:
DAVID ALTFATER
614-836-8786
OH
1
07/16/2008
EredEx QUPS D Hand Delivery Q Other
If other please specify:
861012765368
Site ID
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FW08OH033
FW08OH033
FW08OH033
FW08OH012
FW08OH012
FW08OH012
FW08QH012
FW08OH012
FW08
FW08
FW08
FW08
FW08
FW08
FW08
FW08
Date Collected
MM/DOfmY
07/Q7/20G8
07/07/2008
07/07/2008
07/15/2008
07/15/2008
07/15/2008
07/15/2008
07/15/2008
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Shipped to Lab
MED - DULUTH
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R5H TISSUE 1MB
PERIPHYTON LAB
BENTHICLAB
FISH MUSEUM
OTHER (list below)
Visrt
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
Sample ID
XXXJOOC3C
524309,1
524309,2
524309,3
522039.1
522039,2
522039,3
522039.4
522039.5
Sample
Twe
FTIS
FTIS
FTIS
FTIS
FTIS
FTIS
FTIS
FTIS
BERW
BERW
BERW
BERW
BERW
BERW
BERW
BERW
#of
Jars
1
1
1
1
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XXX-XXX-XXXX
2 digit state code
MM/DD/YYYY
Street address, Qty, State
& Zip Code
Comments
SMAUJ4«ITH BASS 400 MM
SMAilMOUTH BASS 330 MM
SMALIMOUTH BASS 298 MM
CHAN«L CATFISH 4B2 MH
CWAN«L CATFISH 458 MM
CHANNEL CATFISH 460 MM
CHANNEL CATFISH 498 MM
CHANNEL CATFISH 430 MM
Sample Types
PRESERVED - RETAINS)
BEP.W - BenBro (teach WMe
BELG - Benthos Lew Gradient
VERT-RjhVo«*ers
PERI - Nrfffhyton ID(.l)
UWSfSCRVfD - BATCHED
SD€ - Sediment Enzyme
FTIS - Rsh Tissue
PWA - Peifphptai *PA (.4)
Tracking Assistance
TmtUny Mdp:
Marty* Cappwrt
He S41-7S+4467
Mchete&wer
Ph: S41-75+4793
Completed Forms
Send compfeted electronic tracking
forms to:
sampl-etracking@epa.gov
754-4637
£&<*• info to S4I-754-46&3
Save ftie as BR_Si i t ID_Da6e CdHecoed. rcr Site ID & Date coiiectea i^e the rest Sms listed, Njr ocarpple, & yoy are baloiir^i sarttp
m«h fte flrst ID Uteri as RW08CW123 ostacttrf on 05/06/200% then the Be name weuH be BR,J=VWeaRia_OS,,OS, W.
•B6.TCMED - urratei tti=t will be batched and shipped wtNn 2 weeH. Send sample Monnttit*) when SHIPPED.
> - samplet ttut w:i ix Stored longer tan * men* at a hoUng fadMy. Send sampie nfarirjtioo wtten COUHTTED, and then when shaped
Figure E-1. Example Sample Tracking Form showing fish tissue samples, fish
species, and specimen lengths
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