National Rivers and Streams Assessment 2013/14 Field Operations Manual
Version 1.1, Mayl 2013 Non-Wadeable
United States Environmental Protection Agency
Office of Water
Office of Environmental Information
Washington, DC
EPA-841-B-12-009a
National Rivers and Streams
Assessment 2013/14
Field Operations
Manual
Non-Wadeable
Version 1.0
& the Nati
May 2013
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National Rivers and Streams Assessment 2013/14 Field Operations Manual
Version 1.1, Mayl 2013 Non-Wadeable
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National Rivers and Streams Assessment 2013/14 Field Operations Manual
Version 1.0, May 2013 Non-Wadeable
NOTICE
The complete documentation of overall NRSA project management, design, methods, and
standards is contained in four companion documents, including:
National Rivers and Streams Assessment: Quality Assurance Project Plan EPA-841-B-12-007
National Rivers and Streams Assessment: Site Evaluation Guidelines EPA-841-B-12-008
National Rivers and Streams Assessment: Field Operations Manual EPA-841-B-12-009a and b
National Rivers and Streams Assessment: Laboratory Methods Manual EPA 841-B-12-010
This document (Field Operations Manual (FOM)) 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, microcystins, fish
assemblage, fish tissue plugs, whole fish tissue, Enterococci, and physical habitat. These
methods are based on the guidelines developed and followed in the National Rivers and Streams
Assessment 2008-2009 (EPA 2012), 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 2013/14. 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 suggested citation for this document is:
USEPA. 2013. National Rivers and Streams Assessment 2013-2014: Field Operations Manual -
Non-Wadeable. EPA-841-B-12-009a. U.S. Environmental Protection Agency, Office of Water
Washington, DC.
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TABLE OF CONTENTS
NOTICE Ill
TABLE OF CONTENTS V
LIST OF FIGURES VIM
LIST OF TABLES IX
ACRONYMS/ABBREVIATIONS XI
DISTRIBUTION LIST XIII
1 BACKGROUND 1
1.1 SURVEY DESIGN 1
1.2 TARGET POPULATION AND INDEX PERIOD 1
1.3 REPLACING SITES 2
1.4 SELECTION OF NRSA INDICATORS 2
1.5 SUPPLEMENTAL MATERIAL TO THE FIELD OPERATIONS MANUAL 3
1.6 RECORDING DATA AND OTHER INFORMATION 5
2 INTRODUCTION TO NON-WADEABLE SAMPLING 8
2.1 DAILY OPERATIONS 8
2.2 BASE SITE ACTIVITIES 10
2.2.1 Pre-departure Activities 10
2.2.2 Post Sampling Activities 12
2.3 SAFETY AND HEALTH 14
2.3.1 General Considerations 14
2.3.2 Safety Equipment 16
2.3.3 Safety Guidelines for Field Operations 16
2.4 FORMS (PAPER OR ELECTRONIC) 18
2.4.1 Field Forms 18
2.4.2 Tracking Forms 18
2.4.3 Equipment and Supplies 19
3 INITIAL SITE PROCEDURES 20
3.1 SITE VERIFICATION ACTIVITIES 20
3.1.1 Locating the X-Site 20
3.1.2 Determining the Sampling Status of a Stream 21
3.1.3 Elevation at Transect A 24
3.1.4 Sampling During or After Rain Events 24
3.1.5 Site Photographs 24
3.2 LAYING OUT THE SAMPLING REACH 25
3.2.1 Sliding the Reach 28
3.3 MODIFYING SAMPLE PROTOCOLS FOR HIGH OR Low FLOWS 29
3.3.1 Streams with Interrupted Flow 29
3.3.2 Braided Rivers and Streams 30
H
4 WATER CHEMISTRY / CHLOROPHYLL-^ SAMPLE COLLECTION AND PRESERVATION 32 z
4.1 IN SITU MEASUREMENTS OF DISSOLVED OXYGEN, pH, TEMPERATURE, AND CONDUCTIVITY 32 z
4.1.1 Summary of Method 32 8
4.1.2 Equipment and Supplies 32 Q
4.1.3 Sampling Procedure 33 i-y
4.2 WATER CHEMISTRY SAMPLES 35
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4.2.1 Summary of Method 35
4.2.2 Equipment and Supplies 35
4.2.3 Water Chemistry and Chlorophyll-a Sampling Procedure 35
5 MICROCYSTINS 38
5.1 SUMMARY OF METHOD 38
5.2 EQUIPMENT AND SUPPLIES 38
5.3 SAMPLING PROCEDURE 39
6 BENTHIC MACROINVERTEBRATES 40
6.1 SUMMARY OF METHOD 40
6.2 EQUIPMENT AND SUPPLIES 40
6.3 SAMPLING PROCEDURE 42
6.4 SAMPLE PROCESSING IN FIELD 45
7 PERIPHYTON 47
7.1 SUMMARY OF METHOD 47
7.2 EQUIPMENT AND SUPPLIES 47
7.3 SAMPLING PROCEDURE 47
7.4 SAMPLE PROCESSING INTHE FIELD 49
8 PHYSICAL HABITAT CHARACTERIZATION 50
8.1 EQUIPMENT AND SUPPLIES 50
8.2 SUMMARY OF METHODS APPROACH 50
8.3 COMPONENTS OF THE FIELD HABITAT ASSESSMENT 51
8.4 SUMMARY OF WORKFLOW 52
8.5 WORKFLOW AND REACH MARKING 53
8.5.1 Reconnaissance for Physical Habitat Data Collection 53
8.5.2 Thalweg Profile 55
8.6 CHANNEL MARGIN ("LITTORAL") AND RIPARIAN MEASUREMENTS 59
8.6.1 Channel Margin Depth and Substrate 61
8.6.2 Large Woody Debris 62
8.6.3 Bank Angle and Channel Cross-Section Morphology 63
8.7 VISUAL RIPARIAN ESTIMATES 69
8.7.1 Riparian Vegetation Structure 69
8.8 INSTREAM FISH COVER, ALGAE, AND AQUATIC MACROPHYTES 70
8.9 HUMAN INFLUENCES 71
8.10 CANOPY COVER MEASUREMENTS 73
8.11 CHANNEL CONSTRAINT ASSESSMENT, DEBRIS TORRENTS AND RECENT FLOODS 74
8.11.1 Channel Constraint 74
8.11.2 Debris Torrents and Recent Major Floods 77
9 FECAL INDICATOR (ENTEROCOCCI) 80
9.1 SUMMARY OF METHOD 80
9.2 EQUIPMENT AND SUPPLIES 80
£ 9.3 SAMPLING PROCEDURE 80
•^ 9.4 SAMPLE PROCESSING INTHE FIELD 81
i-
g 10 FISH ASSEMBLAGE 83
u
u- 10.1 SUMMARY OF METHOD 83
LU 10.2 EQUIPMENT AND SUPPLIES 86
m 10.3 SAMPLING PROCEDURES 86
H 10.3.1 Irruptive Species 87
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10.3.2 Secondary Electrofishing 92
10.3.3 Secondary Seining 93
ii) Use two crewmembers, each tending a seine pole. Seine along the near shore area 96
10.4 PROCESSING FISH 99
10.4.1 Identification and Tallying 99
10.4.2 Unknown Specimens 99
10.4.3 Photovouchering 102
10.4.4 Preparing Preserved Voucher Specimen Samples 103
10.4.5 Preserving Voucher Specimen Samples 103
10.4.6 Processing Unknown/Range Extension (UNK/RNG) Voucher Samples 106
10.4.7 Processing QA Voucher Samples 106
11 FISH TISSUE PLUG SAMPLING METHOD Ill
11.1 METHOD SUMMARY Ill
11.2 EQUIPMENT AND SUPPLIES Ill
11.3 SAMPLE COLLECTION PROCEDURES 112
12 WHOLE FISH SAMPLING METHOD 115
12.1 METHOD SUMMARY 115
12.2 EQUIPMENT AND SUPPLIES 115
12.3 SAMPLING PROCEDURES 116
13 FINAL SITE ACTIVITIES 120
13.1 OVERVIEW OF FINAL SITE ACTIVITIES 120
13.2 GENERAL SITE ASSESSMENT 121
13.2.1 Elevation at Transect K 121
13.2.2 Watershed Activities and Disturbances Observed 121
13.2.3 Site Characteristics 121
13.2.4 General Assessment 121
13.3 PROCESSING THE FECAL INDICATOR, CHLOROPHYLL-/*, AND PERIPHYTON SAMPLES 123
13.3.1 Equipment and Supplies (Fecal Indicator Filtering) 123
13.3.2 Procedures for Processing the Fecal Indicator Sample 123
13.3.3 Equipment and Supplies (Chlorophyll-afrom Water Sample Filtering) 125
13.3.4 Procedures for Processing the Chlorophyll-a Water Sample 125
13.3.5 Equipment and Supplies (Periphyton Sample) 126
13.3.6 Procedures for Processing the Periphyton Samples 126
13.4 DATA FORMS AND SAMPLE INSPECTION 129
13.5 LAUNCH SITE CLEANUP 130
14 FIELD QUALITY CONTROL 131
14.1 REVISIT SAMPLING OVERVIEW 131
14.2 REVISIT SAMPLING SITES 131
14.3 FIELD EVALUATION AND ASSISTANCE VISITS 132
14.3.1 Specifications for QC Assurance Field Assistance Visits 132
14.4 REPORTING 133
15 REFERENCES 135 tj
APPENDIX A LIST OF EQUIPMENT AND SUPPLIES A-l ^
O
APPENDIX B SAMPLE FORMS B-l ^
O
APPENDIX C SHIPPING AND TRACKING GUIDELINES C-l y
APPENDIX D COMMON & SCIENTIFIC NAMES OF FISHES OF THE UNITED STATES D-l j<
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LIST OF FIGURES
Figure 1.1 Example NRSASite Maps 4
Figure 1.2 Example Sample Labels for Sample Tracking and Identification 7
Figure 2.1 Field Sampling Scenario (Non-Wadeable Sites) 9
Figure 2.2 Overview of Base Site Activities 10
Figure 3.1 Verification Form (front) 22
Figure 3.2 Verification Form (back) 27
Figure 3.3 Sampling Reach Features (Non-Wadeable Sites) 28
Figure 4.1 Field Measurement Form 34
Figure 4.2 Sample Collection Form (front) 37
Figure 6.1 Sample Collection Form (back) 41
Figure 6.2 Benthic Macroinvertebrate Collection (Non-Wadeable Sites) 42
Figure 6.3 Transect Sample Design for Collecting Benthic Macroinvertebrates (Non-Wadeable
Sites) 43
Figure 8.1 Littoral Riparian Plots for Characterizing Riparian Vegetation, human influences, fish
cover, littoral substrate, and littoral depths 54
Figure 8.2Thalweg Profile Form 58
Figure 8.3 Channel/Riparian Transect Form (front) 60
Figure 8.4 Riparian Zone and Instream Fish Cover Plots for a River Cross-Section Transect 61
Figure 8.5 Schematic Showing Bankfull Channel and Incision for Channels 66
Figure 8.6 Determining Bankfull and Incision Heights for (A) Deeply Incised Channels, and (B)
Streams in Deep V Shaped Valleys (Stick figure included for scale) 67
Figure 8.7 Channel/Riparian Transect Form, page 2 (back side) 68
Figure 8.8 Proximity Classes for Human Influences in Non-Wadeable Rivers 72
Figure 8.9 Schematic of Modified Convex Spherical Canopy Densiometer 73
Figure 8.10 Channel Constraint Form 76
Figure 8.11 Types of Multiple Channel Patterns 77
Figure 8.12 Torrent Evidence Form 79
Figure 9.1 Site Assessment Form 82
Figure 10.1 Fish Gear and Sampling Information (front) 84
Figure 10.2 Fish Collection Form 85
Figure 10.3 Reach Layouts for Fish Sampling at Non-Wadeable Sites 89
Figure 10.4 Seining Information Form 98
Figure 10.5 Unknown/Range Extension Voucher Sample Labels and Voucher Specimen Tags . 104
Figure 10.6 Fish Gear and Sampling Information Form (back) 105
Figure 10.7 QA Voucher Sample Labels and Voucher Specimen Tags 107
Figure 10.8 Fish Identification and Count Update Form 109
Figure 12.1 Whole Fish Tissue Collection Form 119
Figure 13.1 Final Site Activities Summary 120
Figure 13.2 Site Assessment Form 122
Figure 14.1 Summary of the Revisit Sampling Design 131
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LIST OF TABLES
Table 1.1 Summary Table of Indicators for all NRSA 2013/14 Sites 3
Table 1.2 Guidelines for Recording Field Measurements and Tracking Information 5
Table 2.1 Stock Solutions, Uses, and Methods for Preparation 12
Table 2.2 Post-sampling Equipment Care 13
Table 2.3 General Health and Safety Considerations 15
Table 2.4 General Safety Guidelines for Field Operations 17
Table 3.1 Equipment and Supplies: Site Verification 20
Table 3.2 Procedure: Site Verification 23
Table 3.3 Guidelines to Determine the Influence of Rain Events 24
Table 3.4 Procedure: Laying Out the Sampling Reach (Non-Wadeable Sites) 25
Table 3.5 Procedure: Sliding the Sampling Reach 28
Table 3.6 Reach Layout Modifications for Interrupted Streams 29
Table 3.7 Procedure: Modifications for Braided Rivers and Streams 30
Table 4.1 Equipment and Supplies: DO, pH, Temperature, and Conductivity 32
Table 4.2 Procedure: Temperature, pH, Conductivity and Dissolved Oxygen 33
Table 4.3 Equipment and Supplies: Water Chemistry Sample Collection and Preservation 35
Table 4.4 Procedure: Water Chemistry and Chlorophyll-a Sample Collection (Non-Wadeable
Sites) 36
Table 5.1 Equipment and Supplies: Microcystin Sample 38
Table 5.2 Procedure: Microcystin Sample Collection (Non-Wadeable Sites) 39
Table 6.1 Equipment and Supplies: Benthic Macroinvertebrate Collection at (Non Wadeable
Sites) 40
Table 6.2 Procedure: Benthic Macroinvertebrate Sampling (Non-Wadeable Sites) 44
Table 6.3 Procedure: Compositing Samples for Benthic Macroinvertebrates (Non-Wadeable
Sites) 45
Table 7.1 Equipment and Supplies: Periphyton (Non-Wadeable Sites) 47
Table 7.2 Procedure: Collecting Composite Index Samples of Periphyton (Non-Wadeable Sites)48
Table 8.1 Equipment and Supplies: Physical Habitat 50
Table 8.2 Components of Non-Wadeable River Physical Habitat Protocol 51
Table 8.3 Summary of Workflow Physical Habitat Characterization (Non-Wadeable) 52
Table 8.4 Procedure: Thalweg Profile 56
Table 8.5 Channel Unit Categories Used on Thalweg Form 59
Table 8.6 Procedure: Channel Margin Depth and Substrate 61
Table 8.7 Procedure: Tallying Large Woody Debris 63
Table 8.8 Procedure: Bank Angle and Channel Cross-Section 64
Table 8.9 Procedure: Characterizing Riparian Vegetation Structure 69
Table 8.10 Procedure: Estimating Fish Cover 71
Table 8.11 Procedure: Estimating Human Influence 72
Table 8.12 Procedure: Canopy Cover Measurements 74
Table 8.13 Procedure: Assessing Channel Constraint 75
Table 9.1 Equipment and Supplies: Fecal Indicator Sampling (Non-Wadeable Sites) 80
Table 9.2 Procedure: Fecal Indicator (Enterococci) Sample Collection (Non-Wadeable Sites).... 80 £2
Table 10.1 Equipment and Supplies: Fish Sampling (Non-Wadeable Sites) 86 co
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Table 10.4 Procedure: Secondary Electrofishing Methods for Wadeable Areas (Non-Wadeable
Rivers) 92
Table 10.5 Procedure: Secondary Seining Methods for Wadeable Areas (Non-Wadeable Rivers)
94
Table 10.6 Procedure: Processing Fish (Non-Wadeable Sites) 100
Table 10.7 Procedure: Processing Unknown/Range Extension (UNK/RNG) Voucher Samples .. 108
Table 10.8 Procedure: Processing QA Voucher Samples 110
Table 11.1 Equipment and Supplies: Fish Tissue Plug Sample Ill
Table 11.2 Recommended Target and Alternate Species for Fish Tissue Plug Collection 112
Table 11.3 Procedure: Fish Tissue Plug Samples 113
Table 12.1 Equipment and Supplies: Whole Fish Tissue Sample Collection 116
Table 12.2 Recommended Target Species for Whole Fish Tissue Collection 117
Table 12.3 Procedure: Whole Fish Tissue Samples 117
Table 13.1 Equipment and Supplies: Fecal Indicator Sample 123
Table 13.2 Procedure: Processing Fecal Indicator Sample 123
Table 13.3 Equipment and Supplies: Chlorophyll-a Processing 125
Table 13.4 Procedure: Chlorophyll-a Sample Processing 125
Table 13.5 Equipment and Supplies: Periphyton Samples 126
Table 13.6 Procedure: ID/Enumeration Samples of Periphyton 127
Table 13.7 Procedure: Preparing Chlorophyll Samples of Periphyton 127
Table 13.8 Procedure: Preparing Ash-Free Dry Mass (AFDM) Samples of Periphyton 129
Table 14.1 General Information Noted During Field Evaluation 133
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ACRONYMS/ABBREVIATIONS
AFDM Ash-Free Dry mass
ANC Alkialinity
Ca Calcium
CAS Chemical Abstract Service
Cl Chloride
CPR Cardiopulmonary Resuscitation
CSDGM Content Standards for Digital Geospatial Metadata
CSV Comma seperated values
CWA Clean Water Act
DELT Deformities, Eroded Fins, Lesions and Tumors
Dl Deionized water
DO Dissolved Oxygen
DOC Dissolved Organic Compound
DQO Data Quality Objective
EMAP Environmental Monitoring and Assessment Program
EPA Environmental Protection Agency
FGDC Federal Geographic Data Committee
FOIA Freedom of Information Act
FOM Field Operations Manual
GIS Geographic Information System
GPS Global Positioning Device
HQ Head Quarters
IBI Index of Biotic Integrity
IQG Information Quality Guideline
IM Information Management
ITIS Integrated Taxonomic Information System
K Potassium
LIMS Laboratory Information Management System
LOM Lab Operations Manual
LRL Laboratory Recording Levels
LWD Large Woody Debris
ISO International Organization for Standardization
MDL Method Detection Levels (limit)
Mg Magnesium
MMI Multimetric Indicators
MSDS Material Safety Data Sheets
MQO Measurement Quality Objective
Na Sodium
NABS North American Benthic Society
NAD North American Datum
NARS National Aquatic Resources Survey
NAWQA National Water-Quality Assessment Program
NCCA National Costal Condition Assessment
NELAP National Environmental Laboratory Accreditation Program
NERL New England Regional Laboratory
NHD National Hydrology Database
NH3 Ammonia
NH4 Ammonium
NIST National Institute of Standards
NLA National Lakes Assessment
NLCD National Land Cover Dataset
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Version 1.0, May 2013
Field Operations Manual
Non-Wadeable
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NO2
NO3
NRC
NRSA
NWCA
0/E
OSHA
OW
PC
PD
PDE
PDF
PP
PPT
PRISM
PTD
QA
QAPP
QA/QC
QCS
QRG
RBS
RL
RTE
RVT
SAS
SDTD
SEG
SiO2
S04
SOP
SQL
Std
STORET
TL
TOC
TN
TP
TSS
UNK/RNG
USGAO
uses
WED
WT
WSA
WQX
Nitrite
Nitrate
National Research Council
National Rivers and Streams Assessment
National Wetland Condition Assessment
"Observed" over "Expected"
Occupational Safety and Health Administration
Office of Water
Personal Computer
Percent Difference
Percent Difference in Enumeration
Personal Flotation Device
Polypropylene
Parts per thousand
Parameter-elevation Regressions on Independent Slopes Model
Percent Taxonomic Disagreement
Quality Assurance
Quality Assurance Protection Plan
Quality Assurance/Quality Control
Quality Check Solution
Quick Reference Guide
Relative Bed Stability
Reporting Limit
Rare, Threatened and Endangered
Revisit
Statistical Analysis System
Spatial Data Transfer Standard
Site Evaluation Guideline
Silica
Sulfate
Standard Operating Procedures
Standard Query Language
Standard
Storage and Retrieval Data Warehouse
Total Length
Total Organic Carbon
Total Nitrogen
Total Phosphorus
Total Suspended Solids
Unknown Range Extension
United States General Accounting Office
United States Geological Survey
Western Ecology Division
Water Tube
Wadeable Streams Assessment
Water Quality Exchange
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DISTRIBUTION LIST
This Field Operation Manual (FOM) and associated manuals or guidelines will be distributed to
the following EPA, senior staff participating in the NRSA and to State Water Quality Agencies or
cooperators who will perform the field sampling operations. The Quality Assurance (QA) Officers
will distribute the Quality Assurance Project Plan (QAPP) and associated documents to
participating project staff at their respective facilities and to the project contacts at participating
laboratories, as they are determined.
Ellen Tarquinio
NRSA Project Leader
Sarah Lehman
NRSA Project QA Officer
Virginia Fox-Norse
OWOW Quality Assurance
Officer
Steven G. Paulsen
EPA ORD Technical Advisor
Marlys Cappaert,
SRA International Inc.
NARS Information
Management Coordinator
Chris Turner, GLEC, Inc.
Contract Logistics Coordinator.
Leanne Stahl
OST Fish Tissue Coordinator
Robert Shippen
OST Fish Tissue QA Officer
Tom Faber, Region 1
Darvene Adams, Region 2
Louis Reynolds, Region 3
David Melgaard, Region 4
Mari Nord, Region 5
Mike Schaub, Region 6
National Monitoring Coordinators
tarquinio.ellen@epa.gov
202-564-2267
lehmann.sarah@epa.gov
202-566-1379
fox-norse.virginia@epa.gov
202-566-1266
paulsen.steve@epa.gov
541-754-4428
cappaert.marlvs@epa.gov
541-754-4467
541-754-4799 (fax)
cturner@glec.com
715-829-3737
Stahl.leanne@epa.gov
202-566-0404
Shippen.Robert@epa.gov
202-566-0391
Regional Monitoring Coordinators
faber.tom@epa.gov
617-918-8672
adams.darvene@epa.gov
732-321-6700
revnolds.louis@epa.gov
304-234-0244
melgaard.david@epa.gov
404-562-9265
nord.mari@epa.gov
312-353-3017
schaub.mike@epa.gov
214-665-7314
U.S. EPA Office of Water
Office of Wetlands, Oceans, and
Watersheds
Assessment and Watershed Protection
Division
Washington, DC
U.S. EPA Office of Water
Office of Wetlands, Oceans, and
Watersheds
Assessment and Watershed Protection
Division
Washington, DC
U.S. EPA Office of Water
Office of Wetlands, Oceans, and
Watersheds
Oceans and Coastal Protection Division
Washington, DC
Freshwater Ecology Branch Western
Ecology Division, NHEERL, ORD, EPA
200 S.W. 35th St. Corvallis, OR 97330
Computer Science Corporation
200 S.W. 35th Street
Corvallis, OR 9733
Great Lakes Environmental Center
739 Hastings Street
Traverse City, Ml 49686
U.S. EPA Office of Water
Office of Science and Technology
Washington, DC
U.S. EPA Office of Water
Office of Science and Technology
Washington, DC
U.S. EPA- Region 1
11 Technology Drive North Chelmsford,
MA 01863-2431
USEPA- Region II
2890 Woodbridge Ave Edison, NJ 08837-
3679
U.S. EPA -Region III
303 Methodist Building Wheeling WV
26003
U.S. EPA -Region IV
61 Forsyth Street, S.W. Atlanta, GA
30303-8960
U.S. EPA - Region V
77 West Jackson Blvd Chicago, IL 60604-
3507
U.S. EPA -Region VI
1445 Ross Ave -Ste 1200 Dallas, TX
CO
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Gary Welker, Region 7
Karl Hermann, Region 8
Terry Fleming, Region 9
Gretchen Hayslip, Region 10
welker.garv(5)epa.gov
913-551-7177
hermann.karl@epa.gov
303-312-6228
fleming.terrence(5)epa.gov
415-972-3462
havslip.gretchen(5)epa.gov
206-553-1685
75202-2733
U.S. EPA -Region VII
901 North Fifth Street Kansas City, KS
66101
U.S. EPA -Region VIII
1595 Wynkoop St .Denver, CO 80202-
1129
U.S. EPA -Region IX
75 Hawthorne Street San Francisco, CA
94105
U.S. EPA- Region X,
1200 Sixth Avenue Seattle, WA 98101
CO
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1 BACKGROUND
This manual describes field protocols and daily operations for crews to use in the National Rivers
and Streams Assessment (NRSA) 2013/14. 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;
• Evaluate changes in condition from the 2008/09 NRSA; and
• 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.
1.1 Survey Design
EPA selected sampling locations using a probability based survey design. The design is an
unequal probability design that selects 900 sites classified as Strahler order 1-4 and 900 sites
classified as Strahler order 5th and above. To evaluate change from the 2008/09 NRSA, 420 of
the 900 l-4th order sites are resampled from the 2008/09 NRSA and 390, 5th orders and above
sites are revisits from the 2008-2009 NRSA. Approximately 10%, or 200, of the total NRSA sites
are scheduled for repeated sampling (revisit sites) in the same field season. The sample frame
was derived from the National Hydrography Dataset (NHD), NHD-Plus, from 1:100,000 scale
maps. Additional details on the NRSA survey design are found in the National Rivers and Streams
Assessment Survey Design: 2013-2014 documents.
1.2 Target Population and Index Period
The target population consists of all streams and rivers within the 48 contiguous states that
have flowing water during the study index period. This includes major rivers, and small streams.
Sites must have > 50% of the reach length with standing water. Sites with water in less than 50%
of the reach length must be dropped. All sites must be sampled during base flow conditions.
The target population excludes:
• Tidal rivers and streams up to head of salt (defined as < .OSppt for this study).
• Run-of-the-river ponds and reservoirs with greater than 7 day residence time.
• The study index period extends from:
o Beginning of June through end of September for most regions
o Sites in the select ecoregions or States can be sampled starting in the end of
April with approval from the EPA Project Coordinator
Please refer to the Site Evaluation Guidelines (EPA-841-B-012-008) and the NRSA Web site 1
(http://www.epa.gov/owow/riverssurvey/index.html) for more detailed information on the
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1.3 Replacing Sites
All base sites must be evaluated for sampling. If a stream or river site is determined to be un-
sampleable, it must be replaced by another site within the state in the same category. The six
general categories for each state are:
• NRSA 2008-2009 resample l-4th Strahler order sites.
• NRSA 2008-2009 resample 5th and above Strahler order sites.
• Small Stream- new 0-2 Strahler order sites.
• Large Streams- new 3-4 Strahler order sites.
• Major Rivers- new 5 and above order sites. Rivers identified as major rivers or additional
rivers in the book: Rivers of North America.
• Other Rivers- new 5 and above order sites that are not considered Major Rivers.
Please refer to the Site Evaluation Guidelines (EPA-841-B-12-008) for more detailed information.
1.4 Selection of NRSA Indicators
As part of the indicator selection process, EPA worked with state and tribal partners and other
partners through technical conferences and indicator teleconferences. The EPA formed a
National Rivers and Streams Assessment Steering Committee with state, tribal and regional
representatives to provide feedback and evaluate core and supplemental indicators to be
included in the 2013/14 field season. Key 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. The core indicators build upon the work done in the NRSA
2008/09. They have been sampled and analyzed on the national scale and have a known
applicability to Clean Water Act (CWA) programs. Supplemental indicators were selected based
on feedback from the Steering Committee and decisions by EPA management. Supplemental
indicators are either in the research phase and their applicability is still being assessed for CWA
programs or this is the first time they will be sampled at a national scale. For field sampling
purposes, there is no distinction between core and supplemental indicators. Indicators that are
included in the NRSA 2013/14 are briefly described in Table 1.1.
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Table 1.1 Summary Table of Indicators for all NRSA 2013/14 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
Microcystin
Periphyton
Benthic macroinvertebrate
assemblage (Littoral)
Fish Assemblage
Physical habitat assessment
Fecal indicator (Enterococci)
Fish Tissue Plug
Whole Fish Tissue
Core or Supplemental
Indicator
Core Indicator
Core Indicator
Core Indicator
Supplemental Indicator
Core Indicator
Core Indicator
Core Indicator
Core Indicator
Supplemental Indicator
Supplemental Indicator
Supplemental Indicator at
select sites
Specs/Location in Sampling Reach
Measurements taken at Transect A at
mid-channel; readings are taken at 0.5
m depth, or mid-depth if water depth
is less than 1 meter.
Measurements taken at Transect A at
mid-channel; Collected from a depth
of 0.5 m, or mid-depth if water depth
is less than 1 meter.
Collected as part of water chemistry
and periphyton samples
Collected from index site
Collected from 11 locations
systematically placed at each site and
combined into a single composite
sample
Collected from 11 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 and .3m depth
Target species collected throughout
the sampling reach as part offish
assemblage sampling
Target species collected throughout
the sampling reach as part offish
assemblage sampling
1.5 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,
landowner access forms, sampling permits (if needed), site evaluation forms and other
information necessary to ensure an efficient and safe sampling day.
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Site maps have been provided to assist in the site evaluation process. Three maps are available:
an aerial image, topographic map, and road map, see Figure 1.1. These maps provide an overlay
of the NHD waterbody layer with the coordinate and label for the x-site and/or waterbody, if
available. Each site is symbolized by the panel the site is considered within (see Section 1.3).
Other important information that may assist in site evaluation is included on the map including:
state, EPA region, latitude, longitude, ownership, and stream order. These maps will be helpful
in the planning and preparation for visiting and sampling a particular NRSA 2013/14 site. These
maps will become part of your site packet.
2013-2014 National Rivers and Streams Assessment
Oklahoma Site: OKRM-1006
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Oklahoma Site: OKRM-1006
Latitude (DD):
== Longitude |DDf
Stream Order:
Ownership:
2013-2014 National Rivers and Streams Assessment
Oklahoma Site: OKRM-1006
11110105000002
Stale: 1:55,000
Imagery Dale: 2010
Imagery Source: National Geo TOPO!
Figure 1.1 Example NRSA Site Maps
Field crews will also receive a Quick Reference Guide (QRG) that contains tables and figures
summarizing field activities and protocols from the Field Operations Manual (FOM). This
waterproof handbook will be the primary field reference used by field crews after reading the
FOM and completing the required field training session. The field crews are also required to
keep the field operations manual available in the field for reference and for possible protocol
clarification.
Quality assurance is a required element of all EPA-sponsored studies that involve the collection
of environmental data (USEPA 2000a, 2000b). Field crews will be provided a copy of the
integrated Quality Assurance and Project Plan (QAPP). The QAPP contains more detailed
information regarding quality assurance/quality control (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-012-007).
Related NRSA documents include the following: National Rivers and Streams Assessment:
Quality Assurance Project Plan (EPA 841-B-12-007), National Rivers and Streams Assessment:
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Site Evaluation Guidelines (EPA 841-B-012-008), and National Rivers and Streams Assessment:
Laboratory Methods Manual (EPA 841-B-012-010).
1.6 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 1.2).
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 1.2. At the end of each sampling day, the
field crew lead is responsible for reviewing each field form for completeness and legibility. The
field crew lead must initial each field form after reviewed.
Table 1.2 Guidelines for Recording Field Measurements and Tracking Information
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 all primary
sampling visits indicate the event as Visit 1. For revisit sites use Visit 2 to
indicate the second sampling event during the same season.
• 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.
En = Miscellaneous flags (n = 1, 2, etc.) assigned by a field crew during a
particular sampling visit (also used for qualifying samples).
Explain reason for using each flag in comments section on data form.
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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 IDs from a single label sheet are in sequential order. Do not mix
labels from different sheets.
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 acceptability requirements, or collected by non-standard
procedure).
Fn = Miscellaneous flags (n=l, 2, etc.) assigned by a field crew 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 Crew Leader must review all labels and data collection forms for
consistency, correctness, and legibility before transfer to the Information
Management Center.
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CHEMISTRY (CHEM)
Site ID:
Date:
/201 Visit*: 01 02
990000
WATER COLUMN CHLOROPHYLL (CHLA)
Site ID:
Date: / 120 \ Visit #:O1 O2
Volume Filtered: mL
990001
PERIPHYTON ASSEMBLAGE ID (PERI)
Site ID:
Date: / /201 Visit*: O1 O2
Composite Volume: mL
990002
PERIPHYTON CHLOROPHYLL (PCHL)
Site ID:
Date:
/201 Visit*: 01 02
Volume Filtered:
ml
990003
PERIPHYTON B1OMASS (PBIO)
Site ID:
Date:
/201 Visit*: 01 02
Volume Filtered:
ml
990004
BENTHIC MACROINVERTEBRATES (BERW)
Site ID:
Date: / /201 Visit #:O1 O2
Jar 1 of
990006
FISH TISSUE PLUG (FPLG)
Site ID:
Date: / /201
990
O
O2
UNK/RNG VOUCHER (VERT)
Site ID:
Date: / /201 Visit*: O1 O2
Date:
ALGAL i OXIN (MICX)
Sit-? lrj: _
^/ /201 Visit #: O1 O2
mL
\.
*s.
/olume Collected:
990005
;>ITHIC MACROINVERTEBRATES (BETB)
Date:
Site ID:
i
/201 Visit*: 01 02
Jar 1 of
Date:
990007
QA VOUCHER (VERT)
Site ID:
/ /201 Visit*: 01 02
990009
Sample Type:
Site ID: _
Date: /
/201 Visit #: 01 02
Sample ID:
Sample Type:
Site ID: _
Date: /
/201 Visit*: O1 O2
Sample ID:
Sample Type:
Site ID: _
Date: I
/201 Visit*: 01 02
Sample ID:
Figure 1.2 Example Sample Labels for Sample Tracking and Identification
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2 INTRODUCTION TO NON-WADEABLE SAMPLING
2.1 Daily Operations
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.
A field crew for a non-wadeable field crew typically will consist of four or five people in 2 boats.
Additional crew members may either help collect samples, or may remain on the bank to
provide logistical support. 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.
A daily field sampling scenario showing how the work load may be split between crew members
is presented in Figure 2.1. The following sections further define the sampling sequence and the
protocols for sampling activities.
Field crews should define roles and responsibilities for each crew member to organize field
activities efficiently. While crews may choose to allocate resources as they see fit, the sequence
of sampling events presented in the Figure 2.1 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.
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Whole Crew
Locate Xsite
Verify site as target
Determine launch site &set upstaging area
Group A Activities:
Group B Activities:
Prepare forms, equipment & supplies
Calibrate multi-parameter probe
Load equipment and supplies onto boat(s) (if boatable)
LOCATE & TRAVEL TO TRANSECT A
Measure in situ temperature
pH, DO, & conductsiy
Collect water chemistry,
chlorophyll-a and microcystin
samples
LOCATE & TRAVEL TO PHYSICAL HABITAT STATIONS
Collect periphyton samples
*
Collect benthic samples
J
Conduct phy
Characte
1
Collect fecal indicator
sample at last transect
RETURN TO STAGING AREA
Preserve benthic samples
prepare for transport
Filterfecal indicator sample;
prepare for transport
Inspect and clean boat, mo-
tor, & trailerto prevent trans-
ferof nuisance species and
contaminants
Filter chlorophyll-a sample;
prepare for transport
Prepare periphyton samples
fortransport
Clean and organize equipmentfor loading
SHIP SAMPLES
Conduct fish assessment
Collect fish tissue samples
RETURN TO STAGING AREA
Prepare fish issue samplesfor
transport
Review data forms for completeness
Inventory supplies for next sampling event,
Request additional supplies f needed
Report sampling event through Site and Sam-
ple Status Form
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Figure 2.1 Field Sampling Scenario (Non-Wadeable Sites)
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2.2 Base Site Activities
Field crews 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 2.2). Close attention to these activities is required to
ensure that the field crews 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.
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PREDEPARTURE ACTIVITIES
Crew Leader
• Prepare daily itinerary
Crew Members
• Instrument checks & calibration
• Equipment & supplies preparation
Whole Crew
Site verification
SAMPLE SITE
POST-SAMPLING ACTIVITIES
Crew Leader
• Review forms & labels
• File status report by email to
IM Team
Crew Members
• Filter, preserve & inspect samples
• Clean boats/gear with 1-10%
bleach solution
• Make any repairs necessary
• Charge or replace batteries
• Refuel boats, vehicles, etc.
• Obtain ice, dry ice and other con-
sumables.
• Package andshipsamples and
data forms
Figure 2.2 Overview of Base Site Activities
2.2.1 Pre-departure Activities
Pre-departure activities include the development of a daily itineraries, instrument checks and
calibration, and equipment and supply preparation. Procedures for these activities are described
in the following sections.
2.2.1.1 Daily Itineraries
The Field Crew Leaders are responsible for developing daily itineraries. This entails compiling
maps, contact information, copies of permission letters, and access instructions (a "site
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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 crews 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.
2.2.1.2 Instrument Checks and Calibration
Each field crew 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 multi-probe unit for measuring DO, pH, temperature, and conductivity, and electrofishing
equipment. Field crews should have access to backup instruments if any instruments fail the
manufacturer performance tests or calibrations. Prior to departure, field crews 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 crew 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 multi-probe against a Quality Check Solution
(QCS).
• 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.
2.2.1.3 Equipment and Supply Preparation
Field crews 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 or 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 2.1. Follow the regulations of the Occupational Safety
and Health Administration (OSHA). g
Site kits of consumable supplies for each sampling site will be delivered based on the supply o-
requests each crew submits prior to and during the sampling season. Crews will submit an <
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electronic request form to order site kits, forms, labels, etc. Field crew leaders MUST provide a LU
tentative 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, report the change as soon as Q
possible to the Field Logistics Coordinator (Chris Turner, cturner@glec.com, (715)829-3737). The >
site kit will include data forms, labels, sample jars, bottles, and other supplies (see complete list ^
in Appendix A). The crews must inventory these site kits before departure. Container labels §
should not be covered with clear tape until all information is completed during sampling at the O
river/stream. Store at least one extra site kit in the vehicle in the event replacement items are -z.
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Table 2.1 Stock Solutions, Uses, and Methods for Preparation
Solution
Bleach (1%)
Clean nets, other gear, and boat.
Preparation
Add 40 mL bleach to 4 L distilled water.
QCS Solution*
Weekly check of meter calibration
A 1:100 dilution of the standard
solution (RIGHT) produces a solution
with the following theoretical values:
pH 6.98
Conductivity 75.3 u.S/cm @ 25oC
STANDARD SOLUTION:
KH2PO4
Na2HPO4
Deionized water
Mix to dissolve
3.4022 g
3.5490 g
1000 ml
10% Buffered Preservation of periphyton ID sample
Formalint and fixing Fish Vouchers
Formaldehyde (37-40%)
Distilled water
NaH2PO4
Na2HPO4 (anhydrous)
Mix to dissolve
100ml
900ml
4.0 g
6.5 g
95% Ethanol Preservative for benthic invertebrate
samples and fish vouchers.
No preparation needed (use stock solution as
is).
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* QCS or "confidence" solutions can also be purchased pre-mixed from various sources.
f 10% Buffered Formalin can also be purchased pre-mixed from various sources
2.2.2 Post Sampling Activities
Upon return to the launching location after sampling, the crew 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 Crew Leader is to provide
the missing information. The Field Crew 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/reorder, sample and data form shipment, and communications.
2.2.2.1 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 by the field crew lead. 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 (a similar review process is used for
electronic forms). 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.
2.2.2.2 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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2.2.2.3 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 (a 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 2.2 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 rinse thoroughly (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.
2.2.2.4 Supply Inventory
Once a field day is completed, crews should inventory and restock supplies as needed. Ensure
that there is a sufficient quantity of site kits to allow sampling at upcoming sites (for at least the
next 1-2 weeks). Take note of any supplies that are nearing depletion. Also note any items that
may have been lost or damaged during the sampling event. Request additional site kits and/or
supplies as needed via the electronic request form. Note that not all supplies can be replenished
by EPA through the Logistics Contractor, so crews will need to supply some items themselves.
Table 2.2 Post-sampling Equipment Care
Equipment Care after Sampling
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 seines, 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.
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Equipment Care after Sampling
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 sealable plastic bag with a cube container for use at the next stream.
Check nets for holes and repair or locate replacements.
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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).
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 2.3. 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)
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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. They must 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 leader. 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 from the
water, boat, and electrodes via rubber boots and linesman gloves. All personnel should 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 crew 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. Ensure that 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. Crew 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 crew leader has authority, each crew
member has the responsibility to question and modify an operation or decline participation if it
is unsafe.
Table 2.3 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)
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
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• Cell (or satellite) phone and VHP radio if possible
Personal Safety
• Field clothing and other protective gear including lifejackets for all crew members
• Medical and personal information (allergies, personal health conditions)
• Personal contacts (family, telephone numbers, etc.)
• Physical exams and immunizations
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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 provided
to field crews working in remote areas in case of an emergency. Supplies (e.g., clean water,
antibacterial 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 2.4. 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 crews 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
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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:
1. 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.
2. 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 crew 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.
3. For most environmental incidents, the following emergency telephone numbers should
be provided to all field crews: 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.
Table 2.4 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 on foot 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.
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
lOoC or 50oF) if he or she is exposed to wind or becomes wet.
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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.
2.4 Forms (Paper or Electronic)
Forms are the key to data collection and tracking for the NRSA 2013/14.Electronic forms have
been developed as well as paper forms. These electronic forms should streamline data
collection. Field crews will have the option of using paper or electronic forms.
2.4.1 Field Forms
Field forms are the primary documents where we record measures, observations, and collection
information during the course of the field day. Additional information regarding specifics of data
entry is contained in Section 1.6.
• Paper Field Forms: A paper field form packet (wadeable or non-wadeable) for each site
will be provided by the NARS IM Coordinator if you have elected to use paper field data
collection. You will need to add these forms to the site packet prior to going in the field.
After a site is sampled, the completed NRSA 2013/14 paper field forms are checked for
completeness and organized sequentially into a Data Packet. The Data Packets from
several sites are batched together and sent every 1-2 weeks to the NARS IM Center and
accompanied by a Tracking form to track which data packets have been shipped. Extra
paper field forms will be provided to field crews to serve as backup copies in case of lost
forms or problems with electronic field forms.
• Electronic Field Forms: This form of data collection can be collected through 3
platforms: an iOS, Android or a Windows portable electronic device (tablets, phones).
This will require a field crew to download or install the developed Application (or "App")
onto the device. The field forms will be optimized for tablet devices. Once downloaded
and the App launched, the field forms will be split into sections or "form-lets" for easier
data entry. It is important for a field crew to familiarize themselves with the App prior
to field sampling.
2.4.2 Tracking Forms
^ Tracking forms describe the status and location of all samples collected during NRSA 2013/14.
% Field crew leaders will typically transmit these forms electronically (by emailing a fillable PDF
< form) to the NARS IM Center at specified times and you will pack hard copies in shipping
^ containers with the samples. See APPENDIX C: SHIPPING GUIDELINES for more information.
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O site (immediate shipment and batch shipments). This also serves as the tracking form for
Q sample shipped to the WRS lab.
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p and shipped every 1 or 2 weeks. These are sent to the NARS IM Center.
Q • Tracking -Samples: Accompany samples that are batched together from multiple sites
oc and shipped every 1 or 2 weeks. Whenever batched samples are shipped to their
^ designated lab for analysis, the appropriate tracking form, which lists the Sample ID
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numbers for all samples packed in a shipping container, is included in the shipping
package and is also transmitted electronically to the NARS IM Center. Separate forms
exist for the tracking of frozen batched samples, non-chilled batched samples and whole
fish samples
2.4.3 Equipment and Supplies
2.4.3.1 Request Form
Field Crews will submit requests for field forms, labels and site kits via an electronic Request
Form. This form will be submitted to the NARS Information Management (IM) Coordinator who
will ensure that the request reaches the appropriate entity. Crews must submit sampling
schedules at or before the time of submitting request forms. Crews should submit the Request
Form at least 2 weeks prior to their desired sampling date.
2.4.3.2 Base Kit
The Base Kit is comprised of the subset of durable equipment and supplies needed for NRSA
2013/14 sampling that is provided by USEPA through the Contract Field Logistics Coordinator.
Typically one Base Kit is provided to each Field Crew and contains some of the equipment that is
used throughout the field season. See APPENDIX A: EQUIPMENT & SUPPLIES for a list of the
items provided by USEPA in the Base Kit. We anticipate that this equipment will be available for
use in future NRSA efforts.
2.4.3.3 Site Kit
A Site Kit contains the subset of consumable supplies (i.e., items used up during sampling or
requiring replacement after use) provided by USEPA through the Contract Field Logistics
Coordinator. The site kit will contain all the sample bottles necessary for sampling a single site. A
new Site Kit is provided for each site sampled. See APPENDIX A: EQUIPMENT & SUPPLIES for the
consumable items that will be provided by USEPA.
2.4.3.4 Field Crew Supplied Items
The field crew will also supply particular items for the field sampling day. These might include
supplies from the previous NRSA, typical field equipment (like a GPS), or boat equipment. See
APPENDIX A: EQUIPMENT & SUPPLIES for the items that the field crew will need to provide. ^
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3 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
Guidleines EPA-841-B-12-008). 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.
3.1 Site Verification Activities
3.1.1 Locating 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 to each
field crew leader. Table 3.1 provides the equipment and supplies needed for site verification.
Note that the coordinates provided on the site spreadsheet may not be located in the middle of
the stream or river; and in some cases, the coordinates may be on dry land next to the stream or
river. In these cases, it is important for crews to locate the x-site at a point that is in the middle
of the stream or river (e.g. midway between the left and right banks). To do this, simply
measure the distance between banks and move the point perpendicular to the nearest bank
until it is half-way between the left and right banks.. Record these coordinates as the x-site on
the verification form. If the coordinates are located on dry land near a stream, move the
coordinates to the nearest blue line during the desktop reconnaissance. Note this movement on
the site recon tracking form and in the comments section of the Site Verification form
Table 3.1 Equipment and Supplies: Site Verification
For locating and
verifying site
Sampling permit and landowner access (if required).
Field Operations Manual and laminated Quick Reference Guide.
Site dossier, including access information, site spreadsheet with map coordinates,
street and/or topographic maps with "X-site" marked.
NRSA Community Fact Sheets.
GPS unit (preferably one capable of recording waypoints) with manual, reference
card, extra battery pack.
Surveyor's flagging tape (to mark transects).
Laser rangefinder.
50 m or 100 m measuring tape with reel (if not using rangefinder).
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Fine-tipped indelible markers to write on flagging
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3.1.2 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 (see Table 3.2). 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 an access to the site.
There must be greater than 50% water throughout the channel reach. If the channel is dry at
the X-site, determine if water is present within 75 m upstream and downstream of the X-site. 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 3.1.1. Do not drop
the site if it is dry at the X site, as long as there is greater than 50% water throughout the
channel. 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 3.2.1) 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 3.1). 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 same
category based on Strahler order and whether it is a 2008/09 resample site (Section 1.3).
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Site ID:
NRSA 2013/14 VERIFICATION (Front)
Visit: O 1 Oz Date: /
Site Name:
State of Site Location:
Field Crew:
STREAM/RIVER VERIFICATION INFORMATION
Stream/River verified by (Mark all that apply): O GPS
O Other (Describe Here):
O Local Contact O Signs
O Roads O Topo. Map
Coordinates
Decimal Degrees
Latitude
Longitude
Type of GPS Fix Elevation at transect A
C)2D Q3D
Location: O X-5lte (wadeable) O Transect A (non-wadeable)
DID YOU SAMPLE THIS SITE?
O YES |f Yes, check one below
SAMPLEABLE {Choose method used)
O Wadeable-Continuous water,greaterthan 50% wadeable
O Scalable
O Partial - Sampled by wading |>50% of reach sampled). Explain below,
O Partial -Sampled by boa! (>50'A of reach sampled). Explain below.
O Wadeablelnterrupted-Not continuous water along reach
O Boatablelnterrupted -Not continuous water along reach
O Altered-Stream/RiverChannBl PresenlbutdiffersfromMap
ADDITIONAL SITE CHARACTERISTICS
O Tidally Influenced O Blackwater O Not Applicable
O NO if No, Chech one below
NON-SAMPLEABLE-PERMANENT
O Dry-Visited
O Dry-Not visited
O Wetland (No Definable Channel)
O Map Error (No evldencechannel/waterhody ever present)
O Impounded (>7dayresldencellme)
O Tidal (exceeds salinity threshold)
NON-SAMPLEABLE-TEMPORARY
O Other (explain In comments)
O Not bodtable • Need a different crew- Reschedule for this year
O Not wadeable • Need a different crew- Reschedule for this year
NO ACCESS
O Other (Explain in comments)
Q Access Permission Denied
O Permanently Inaccessible tUnablefUnsafetaReachSlte)
O Temporarily Inaccessible-Fire, etc. - Reschedulefornextyear
GENERAL COMMENTS
DIRECTIONS TO SITE
04/08/2013 2013 Verification
5409334985
Figure 3.1 Verification Form (front)
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Table 3.2 Procedure: Site Verification
Site Verification Procedures
1. Find the stream/river location in the field corresponding to the X-site coordinates. 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 83). 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 United States Geological Society (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 bubble on the verification form.
5. 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, sampled by wading.
Boatable: Continuous water, too deep to sample by wading.
Partial wadeable: Sampled by wading (>50% of reach sampled).
Partial boatable: Sampled by boat (>50% of reach sampled).
Wadeable Interrupted: not continuous water along reach, >50% water in reach.
Boatable Interrupted: not continuous water along reach, >50% water in 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 manmade 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, and 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.
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No Access to Site Categories
Access Permission Denied: You are denied access to the site by the landowner.
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).
6. Do not sample non-target or "Non-sampleable" or "No Access" sites. Fill in the "NO" bubble for
"Did you sample this site?" and mark the appropriate bubble in the "Non-Sampleable" or "No
Access" section of the Verification Form; provide a detailed explanation in comments section.
3.1.3 Elevation at Transect A
Record elevation at Transect A using your GPS device. To record this information, record the
elevation holding the GPS at approximately 3 feet above the surface of the water. Ensure that
the numbers are properly recorded from Transect A on the Site Verification form.
3.1.4 Sampling During or After Rain Events
Avoid sampling during high flow rainstorm events. Use your best professional judgement to
determine if the stream has risen above baseflow during this recent rain event. 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 crew. Some guidelines to help make
this decision are presented in Table 3.3. 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.
Table 3.3 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.
3.1.5 Site Photographs
Taking site photographs is an optional activity, but should be considered if the site has unusual
natural or manmade 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
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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.
3.2 Laying out the sampling reach
Many 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
wetted 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 Table 3.4 (non-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.
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 the Verification Form (back)
as shown in Figure 3.2.
Figure 3.3 illustrates the principal features of the established sampling reach for non-wadeable
sites, including the location of the 11 cross section transects used for collecting samples and
physical habitat measurements. The figures also show the specific sampling stations on each
transect for collection of 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 3.2). 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 3.4 Procedure: Laying Out the Sampling Reach (Non-Wadeable Sites)
Laying out the sampling reach at the base site (recommended at boatable sites)
1. Using GIS, an aerial photo or a 1:100:000 topographic map; locate the X-site using the
coordinates provided for the site.
2. Determine the average wetted width of the channel at the X-site using GIS or if not
available 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 GIS. 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 transects into a GPS unit so 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).
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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
3.3). Note that left and right sides of the stream are determined while you are facing
downstream. It is at these locations that you will collect benthic macroinvertebrate and
periphyton samples.
7. When you are at the site, "ground truth" the wetted width measurements and proceed
to Table 3.5 to see if the layout needs to be adjusted.
Laying out the sampling reach in the field
1. Locate the X-site using the coordinates provided for the site.
2. 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.
3. 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).
4. Check the condition of the stream about the X-site by having one crew 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 2.
5. Determine if the reach needs to be adjusted about the X-site due to confluences with
higher order streams (downstream), transitions into 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 3.5 for specific instructions.
6. Starting at the X-site (or the new midpoint of the reach if it had to be adjusted as
described in Step 8), 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.
7. 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. Note that left and right sides of the stream are determined while
you are facing downstream. It is at these locations that you will collect benthic
macroinvertebrate and periphyton samples.
8. 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 3.3. 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.
9. 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 downstream) at
intervals equal to 1/10 of the reach length. Continue to assign the sampling stations
systematically.
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V
Site ID:
NRSA 2013/14 VERIFICATION (Back)
Visit: 01 O2 Date: /
STREAM/RIVER REACH DETERMINATION
Channel Width
Used to Define
Reach(m)
DISTANCE (ra) FROM X-SITE
Upstream Length:
Downstream Length:
Total Reach
Length Intended (m):
Comment:
SKETCH MAP
Arrow Indicates North; Mark site L=Launch X=l ndex T= Take Out
NOTE: If an outline map Is attached here, use a continuous slrlp 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-slte and transect locations marked.
PERSONNEL
Crew Leader:
Fish Taxonomist
Name;
Name:
Name:
Name:
Name:
Name:
04/08/2013 2013 Verification
4426334983
Figure 3.2 Verification Form (back)
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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 5m up & downstream from
each transect (10m x 15m)
Total reach length = 40 x mean wetted width (min = 150 m; max = 4 km)
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Figure 3.3 Sampling Reach Features (Non-Wadeable Sites)
3.2.1 Sliding the Reach
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. Reasons for sliding the reach include:
• Landowner access;
• Confluence with higher order waterbody;
• Impoundment;
• Impassable barrier.
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 3.5 describes when you should and should not slide the
sampling reach.
Table 3.5 Procedure: 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. Slide the reach if you run into a confluence (another stream meeting the water-body you are
sampling) with a higher Strahler Order.
4. When you are denied access permission to a portion of the reach, you can slide the reach to make
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it entirely accessible; use the point of access restriction as the endpoint of the reach.
5. 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.
6. Do not slide the reach so that the X-site falls outside of the reach boundaries.
7. 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).
8. Do not slide a reach to avoid man-made obstacles such as bridges, culverts, rip-rap, or
channelization. These represent important features and effects to study.
9. Do not slide a reach to gain more water to sample if the flow is interrupted.
10. Do not slide a reach to gain better habitat for benthos or fish.
3.3 Modifying Sample Protocols for High or Low Flows
3.3.1 Streams with Interrupted Flow
You cannot collect the full complement of field data and samples from streams that are
categorized as "Interrupted" (Table 3.6). Note that no data should be collected from streams
that are completely "Dry" as defined in Table 3.6. 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 3.6. Samples for water chemistry (Section 4) will be collected at Transect A (even if the
reach has been adjusted by "sliding" it). If Transect A 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 3.6 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 Transect A 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 Transect A. Note that the
sample wasn't collected at the Transect A 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 water 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 and Benthic Macroinvertebrates
Obtain a complete thalweg profile for the entire reach. At points where the 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
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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 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 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, benthic macroinvertebrate, and fish indicators, using standard procedures.
3.3.2 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 3.7. Figuring the
mean width of extensively braided rivers and streams for purposes of setting up the sampling
reach length is challenging. For braided channels, measure the mean width and bankfull width
as defined in the physical habitat protocols (Section 8). 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 (Figure 3.1 and Figure 3.2) about what you did. It is
important to remember that the purpose of the 40 channel width reach length is to sample
enough streams 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 3.7 Procedure: Modifications for Braided Rivers and 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, using the mean bankfull width for your calculations.
• 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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4.1
WATER CHEMISTRY / CHLOROPHYLL-a SAMPLE COLLECTION
AND PRESERVATION
In Situ Measurements of Dissolved Oxygen, pH, Temperature, and
Conductivity
4.1.1 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 Transect A.
Take the readings at 0.5 m depth. Measure the site depth accurately before taking the
measurements. If the depth at the x-site is less than 1 meter, take the measurements at mid-
depth. Take care to avoid the probe contacting bottom sediments, as the instruments are
delicate. Record the measurements on the Field Measurement Form, as seen in Figure 4.1.
4.1.2 Equipment and Supplies
Table 4.1 provides the equipment and supplies needed to measure dissolved oxygen, pH,
temperature, and conductivity.
Table 4.1 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 (Dl) and tap water
• Calibration cups and standards
• Barometer or elevation chart to use for calibration
• Field Measurement Form
• Pencils (for data forms)
4.1.2.1 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. Follow your manufacturer's guidelines for calibration of the DO probe.
pH Meter
Calibrate the pH meter prior to each sampling event. Calibrate the meter in accordance with the
manufacturer's instructions and with the crew agency's existing SOP.
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.
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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.
4.1.3 Sampling Procedure
Table 4.2 presents step by step procedures for measuring dissolved oxygen, pH, temperature,
and conductivity.
Table 4.2 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 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 Transect A, 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
(or at mid-depth if less than 1 meter deep).
2. Record the measurements on the Field Measurement Form
3. Flag any measurements that the crew feels needs further comment or when a measurement
cannot be made.
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• NRSA 2013/14 FIELD MEAS
Site ID:
IRFMFNT "-•' — n-fn-i-n. •
Date: / /
CALIBRATION INFORMATION
Instrument manufacturer and model:
Instrument ID number:
TEMPERATURE
DO
pH
CONDUCTIVITY
Thermometer
Reading ("C) Sensor Rea ding {"C> Comments
i i it it i it i
Barometric
Elevation Pressure (mm Hg) Calibration
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4.2 Water Chemistry Samples
4.2.1 Summary of Method
The water chemistry samples will be analyzed for total phosphorus (TP), total nitrogen (TN),
total ammonium (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). All field crews must collect a grab sample in one 4-L cube container and in
one 2-L amber Nalgene bottle from Transect A at the midpoint of the reach. Store all samples on
ice in a closed cooler. After Collection, store all samples in darkness on ice in a closed cooler.
After you filter the chlorophyll-o sample, the filters must be kept frozen until ready to ship.
4.2.2 Equipment and Supplies
Table 4.3 provides the equipment and supplies needed to collect water samples at Transect A.
Record the Water Sample Collection and Preservation data on the Sample Collection Form, as
seen in Figure 4.2.
Table 4.3 Equipment and Supplies: Water Chemistry Sample Collection and Preservation
For collecting samples
For recording
measurements
Nitrile gloves
4-L cube container
2-L amber Nalgene bottle
3-L Nalgene beaker
Cooler with ice
Dl water (for cleaning beaker between sites)
Field Operations Manual and laminated Quick Reference Guide
Sample Collection Form
Water Chemistry sample label with pre-printed Sample ID
Clear tape strips
Field Measurement Form
Pencils (for data forms)
Fine tipped indelible markers
4.2.3 Water Chemistry and Chlorophyll-a Sampling Procedure
Table 4.4 describes the sampling procedures for collecting water chemistry samples in non-
wadeable streams and rivers.
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Table 4.4 Procedure: Water Chemistry and Chlorophyll-a Sample Collection (Non-Wadeable Sites)
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1. Fill out the pertinent information (Site ID and date) on the water chemistry label and affix the label
to the cube container. Completely cover the label with clear tape.
2. Collect the water samples from Transect A in a flowing portion near the middle of the site.
3. Put on nitrile gloves. Make sure not to handle sunscreen or other chemical contaminants until after
the sample is collected.
4. Rinse the 3-L Nalgene beaker three times with water, and discard the rinse downstream.
5. Remove the cube container lid and expand the cube container by pulling out the sides. NOTE: DO
NOT BLOW into the cube container or place your fingers inside the opening to expand it, this will
cause contamination.
6. 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.
7. 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.
Chlorophyll-a
8. 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.
9. Fill the beaker with water and pour into the 2 L amber Nalgene bottle, filling the bottle. Cap the
bottle tightly. This sample will be filtered later and the bottle will be reused at future sites,
therefore it is not necessary to label this bottle.
Place the cube container and Nalgene 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.
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 the sample was collected at Transect A, darken the Transect A
bubble in the "STATION COLLECTED" field. If you had to move to another part of the reach to collect the
sample, place the letter of the nearest transect in the "STATION COLLECTED" field. Record more detailed
reasons and/or information in the Comments section.
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Site ID:
MRS A 2013/14 SAMPLE CO
LLECTION (Front) R*Vi™BdhyrinEb»i>: ^m
Date: / /
CHEMISTRY (CHEM) STATION COLLECTED: . .
(Target Volume = 4L) Q X-SIt* (wadeablej O Transect A (non-wadeable) O OtherTranseet: | | No ^""P18 Collected Q
Sample ID
1 1 1 1
Chilled Comments
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WATER COLUMN CHLOROPHYLL (CHLA)
(Target Volume = 1000mL; max vol = 2000 mL)
Sample ID
till
Volume Filtered c ,,
(mil Frozen comments
O
I I
COMPOSITE PERIPHYTON
Composite Volume
No Sample Collected Q
No Sample Collected Q
No of
Transects Commenls
Assem Wage ID (PERI) (SO-mL tube) Chlorophyll (PCHL) (GF
Sample ID
i i
Volume (ml)
Sample ID
i i i i i iii
Preserved Flag Volume ( ml ) Froze
0 0
IF Filter) Biomass (PBIO) (GF/C Filter)
Sample ID
i i i i i i i i i i i
i Flag Vdume(ml) Frozen Flag
0
Flag Comments
ALGAL TOXIN
(Target Volume
Sample ID
lilt
(Microcystin) (MICX)
= 500 ml)
Frozen Comments
O
1 1
ENTEROCOCCI (ENTE)
(Target Volume = 250 mL)
Sample ID
1 1 1 1
Time Depth Sample Ftlt. Sta
Collected Collected VoLme Time
(hhmm) (m) (mL) (hhmm
1 1 1
Comments
Filer blank is colected during visit 1 at at revisil sites.
^ 04/08/2013 2013 Sample Colled on
No Sample Collected Q
No Sample Collected O
Blank Collected Q
t Volume Filtered Flit. End Time
(Target = 50 mL) Time Frozen
1 Filt 1 Fit. 2 (hhmm) (hhmm)
3054051198 |
Figure 4.2 Sample Collection Form (front)
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5 MICROCYSTINS
Microcystis is a microscopic organism that is found naturally at low concentrations in freshwater
systems. Under optimal conditions (such as high light and calm weather, usually in summer),
Microcystis occasionally forms a bloom, or dense aggregation of cells, that floats on the surface
of the water forming a thick layer or "mat." At higher concentrations, Microcystis blooms are so
dense that they resemble bright green paint that has been spilled in the water. These blooms
potentially affect water quality as well as human health (Microcystis produces microcystin, a
potent liver toxin) and natural resources. Decomposition of large blooms can lower the
concentration of dissolved oxygen in the water, resulting in hypoxia (low oxygen) or anoxia (no
oxygen). Sometimes, this results in fish kills. The blooms can also be unsightly, often floating at
the surface in a layer of decaying, odiferous, gelatinous scum.
Although the likelihood of people being affected by a Microcystis bloom is low, minor skin
irritation can occur with contact, and gastrointestinal discomfort can also occur if water from a
bloom is ingested. People recreationally exposed (e.g., swimmers or personal watercraft
operators) to microcystins have also reported minor skin irritation. Health problems may occur
in animals if they are chronically exposed to fresh water with Microcystis present. Just as
livestock and domestic animals can be poisoned by drinking contaminated water, fish and bird
mortalities have been reported in waterbodies with persistent Microcystis blooms.
5.1 Summary of Method
The microcystin (algal toxin) sample is a grab sample taken from Transect A. All field crews must
collect a grab sample using the 3 L beaker to fill a 500ml bottle. Collect this sample after the in
situ measurements and water chemistry sample are collected. Store all samples on ice in a
closed cooler.
5.2 Equipment and Supplies
Table 5.1 provides the equipment and supplies needed to collect microcystin sample at the
index site ( Transect A if reach has not been slide).. Record the Water Sample Collection and
Preservation data on the Sample Collection Form, as seen in Figure 4.2.
Table 5.1 Equipment and Supplies: Microcystin Sample
For collecting samples
Nitrile gloves
3-L Nalgene beaker
500 ml Nalgene bottle
Cooler with ice
Field Operations Manual and laminated Quick Reference Guide
For recording
measurements
Sample Collection Form
Microcystin sample label with pre-printed Sample ID
Clear tape strips
Field Measurement Form
Pencils (for data forms) and Fine tipped indelible markers for labels
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5.3 Sampling Procedure
Table 5.2 presents step-by-step procedures for collecting microcystin samples at non-wadeable
sites.
Table 5.2 Procedure: Microcystin Sample Collection (Non-Wadeable Sites)
Microcystin Sample Collection
1. Fill out the pertinent information (Site ID and date) on the microcystin label and affix the label to
the 500 mL Nalgene bottle. Completely cover the label with clear tape.
2. Collect the microcystin samples from Transect A in a flowing portion near the middle of the site.
3. Put on nitrile gloves. Make sure not to handle sunscreen or other chemical contaminants until
after the sample is collected.
4. Rinse the 3-L Nalgene beaker three times with water, and discard the rinse downstream.
5. Fill the 3 liter beaker with water and slowly pour 30 - 50 ml into the 500 ml 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.
6. Fill the beaker with water and pour into the 500 ml Nalgene bottle. Fill bottle, leaving one inch of
head space in the bottle to allow for expansion when frozen.
Place the 500 ml Nalgene 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.
Record the Sample ID on the Sample Collection Form along with the pertinent site information (site
name, ID, date, etc.).
As soon as you return to your base site (hotel, lab, office, etc.), freeze the entire sample and keep frozen
until shipping.
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6 BENTHIC MACROINVERTEBRATES
6.1 Summary of Method
Collect benthic macroinvertebrate composite samples using a D-frame net with 500 u.m mesh
openings. Take the samples from the sampling stations at the 11 transects equally distributed
along the targeted reach. Multiple habitats will be encountered and sampled using this
approach. They include bottom substrate as well as woody debris, macrophytes, and leaf packs.
Composite all sample material and field preserve with ~95% ethanol.
6.2 Equipment and Supplies
Table 6.1 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 back of Sample Collection Form (Figure 6.1).
Table 6.1 Equipment and Supplies: Benthic Macroinvertebrate Collection at (Non Wadeable Sites)
For collecting
samples
Modified kick net (D-frame with 500 u.m
mesh) and 4-ft handle
Watch with timer or stopwatch
Buckets, plastic, 8 to 10 qt (optional)
Sieve bucket with 500 u.m 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 Composite benthic sample labels with &
measurements without preprinted ID Sample 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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Site ID:
NRSA 2013/14 SAMPLE COLLECTION (Back)
Date: /
BENTHIC MACROINVERTEBRATES (BERW) - WADEABLE
No Sample Collected Q
Presence! No. of
Number of jars (ETQH) Transects Comments
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REACH-WIDE BENTHOS - WADEABLE
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BENTHIC MACROINVERTEBRATES (BETB) - BOATABLE
No Sample Collected Q
Sample ID
Preserved No. of
Number of jars (ETON) Transects Comments
O
TRANSECT BENTHOS - BOATABLE
Habitat: C = Coarse Substrate / LWD L = Leaf Pach F = Organic Fine Muds / Sand M = Macropnyte beds OT = Other (Flag and expiain m ccmment *«ton below)
Substrate: F = Fine / Sand G = Gravel C = Coarse substrate OT = Other [Flag and explain in comment secLon below)
Channel: P = Pool GL = Glide RI = Rlffle RA= Rapid OT = Other (Flag and explain in comment secDon betow)
TRANSECT
Locallon (LIR|:
A
PL OR
B
OL OR
C
OL QH
D
OL OR
E
OL OR
F
OL OR
G
OL OR
H
OL OR
I
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K
OL OR
Dominant
Habitat:
OF O
Q OT
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Q OT
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Q QT
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Oc OL
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OF Oc
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QQT
Flag
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Flag
Flag
Flag
0629051193
04/08/2013 2013 Sample Collection
Figure 6.1 Sample Collection Form (back)
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6.3 Sampling Procedure
Figure 6.2 summarizes how samples will be collected from non-wadeable sites. Collect benthic
macroinvertebrate samples at the 11 transects and within the sampling stations for non-
wadeable streams (Figure 6.3). The process for selecting the sample stations is described in the
Initial Site Procedures (Section 3). 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.
The procedure for collecting samples at each transect is described in Table 6.2. Take 1 linear
meter sweep at the dominant habitat type. Record the benthic macroinvertebrate collection
data on the Sample Collection Form as seen in Figure 6.1. As you proceed from transect to
transect, combine all samples into a bucket.
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NON-WADEABLE
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At Transect A, locate 1st sampling sta-
tion and determine primary and secon-
dary habitat types
(only primary is to be sampled)
Sweep 1 linear meter of the dominant
habitat type
Transfer sample into bucket
I
Mark bubble for substrate & channel
habitat types on the Sample Collection
Form.
I
Thoroughly rinse net and proceed to the
next sampling location.
I
Composite the samples from all stations
to create a single sample for the site.
Figure 6.2 Benthic Macroinvertebrate Collection (Non-Wadeable Sites)
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Figure 6.3 Transect Sample Design for Collecting Benthic Macroinvertebrates (Non-Wadeable Sites)
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Table 6.2 Procedure: Benthic Macroinvertebrate Sampling (Non-Wadeable Sites)
Collecting the Benthic Macroinvertebrate Sample
1. After locating the sampling station at Transect A according to procedures described in the reach
layout section, identify the dominant and secondary habitat type within the plot:
• Rocky/cobble/gravel/large woody • Organic fine mud or sand
debris
• Macrophyte beds • Leaf Pack
2. Use the D-frame dip net (equipped with 500 u.m 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.
a) 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.
b) Because a dip-net is being used for sampling, the maximum depth for sampling will be
approximately 0.5 to 1 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 side of the river on which the sampling station is located (left or right).
5. Record the sampled habitat type (Dominant) as well as the Secondary habitat type on the Sample
Collection Form.
• Fine/sand (F): not gritty (silt/clay/muck <0.06 mm diam.) to gritty, up to ladybug sized (2
mm)
• eravel (G): fine to coarse gravel (ladybug to tennis ball sized; 2 mm to 64 mm)
• Coarse (C): Cobble to boulder (tennis ball to car sized; 64 mm to 4000 mm)
• Other (OT): 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.
6. Identify the channel habitat type where the sampling station was located. Indicate the
appropriate channel habitat type for the transect on the Sample collection Form. The channel
habitat types are:
• Pool (P): Still water; low velocity; smooth, glassy surface; usually deep compared to other
parts of the channel
• Glide (GL): Water moving slowly, with smooth, unbroken surface; low turbulence
• Riffle (Rl): Water moving, with small ripples, waves, and eddies; waves not breaking, and
surface tension is not broken; "babbling" or "gurgling" sound.
• Rapid (RA): Water movement is rapid and turbulent; surface with intermittent "white
water" with breaking waves; continuous rushing sound.
• Other (OT): Note type of "other" channel habitat in comments on field form.
7. Proceed to the next sampling station and repeat steps 1-6. 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. Record the number of transects that were sampled throughout the reach.
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6.4 Sample Processing in Field
Use a 500 u.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. 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/£ full of material. Multiple jars may be required if detritus is heavy
(Table 6.3). If there is lots 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.
Try to use no more than 4 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.
Record information for each composite sample on the Sample Collection Form as shown in
Figure 6.1. Place the samples in a cooler or other secure container for transporting and/or
shipping to the laboratory (see Appendix C).
Table 6.3 Procedure: Compositing Samples for Benthic Macroinvertebrates (Non-Wadeable Sites)
Compositing Benthic Macroinvertebrate Sample
1. Pour the entire contents of the bucket into a sieve bucket with 500 u.m mesh size. Remove any
large objects and wash off any clinging organisms back into the sieve before discarding. Remove
any large 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 many 1-L jars will be
required. Try to use no more than 4 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 Vi full, or use a second jar if necessary. 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).
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"
6. 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
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7.
8.
Compositing Benthic Macroinvertebrate Sample
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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7 PERIPHYTON
7.1 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 transect location as the benthic macroinvertebrate samples (Section
6), directly after the benthic macroinvertebrate samples have been collected. Prepare one
composite sample of periphyton for each site. At the completion of the day's sampling activities,
but before leaving the site, prepare three types of laboratory samples (an ID/enumeration
sample to determine taxonomic composition and relative abundances, a chlorophyll sample,
and a biomass sample (for ash-free dry mass [AFDM])) from the composite periphyton sample.
7.2 Equipment and Supplies
Table 7.1 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 to ensure that all of the required equipment is
brought to the river.
Table 7.1 Equipment and Supplies: Periphyton (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 graduated plastic bottle for the composite sample with
marked volume gradations
60-mL plastic syringe with tip removed, and length of tubing
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 (3 per set) with the sample ID number
Clear tape strips for covering labels
7.3 Sampling Procedure
At each of the 11 transects, collect samples from the sampling station assigned during the layout
of the reach (left or right). 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
7.2. 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 composite sample on the Sample
Collection Form as shown in Figure 4.2.
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Table 7.2 Procedure: Collecting Composite Index Samples of Periphyton (Non-Wadeable Sites)
Periphyton Composite Sample
1. Starting with Transect "A", collect a single sample from the assigned sampling station using the
procedure below.
If coarse substrate (cobbles, woody materials, etc.) are present that can be removed from the
water:
(a) Collect a sample of substrate (rock or wood) that is small enough (< 15 cm diameter) and can be
easily removed from the water. 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 water from this bottle, wash the dislodged periphyton
from the funnel into the 500-mL bottle. Use an amount of water (~45 mL) that brings the
composite volume up to the next graduation mark on the bottle.
(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.)
If large coarse substrate is present that is too large to remove from the water (bedrock, large
woody materials, boulders, etc.):
(a) 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 using the tip of the syringe in a
scraping motion.
(b) While dislodging periphyton with the syringe tip, simultaneously pull back on the syringe
plunger to draw the dislodged periphyton into the syringe.
(c) Empty the syringe into the same 500-mL plastic bottle as above. If the volume of the vacuumed
sediment is not enough to raise the composite volume to the next graduation on the bottle
(~45 mL), add additional stream water to the bottle to raise the level to the next graduation.
(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.)
If no coarse sediment (cobbles or larger) are present:
(a) Use the area delimiter to confine a 12-cm2 area of soft sediments.
(b) Vacuum the top 1 cm of sediments from within the delimited area into a de-tipped 60-mL
syringe.
(c) Empty the syringe into the same 500-mL plastic bottle as above. If the volume of the vacuumed
sediment is not enough to raise the composite volume to the next graduation on the bottle
(~45 mL), add additional stream water to the bottle to raise the level to the next graduation.
(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.
Storage
3. After samples have been collected from all 11 transects, thoroughly mix the 500-mL bottle
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regardless of substrate type. Record the total volume of the composite sample in the periphyton
section of the Sample Collection Form.
4. If all 11 samples are not collected, record the number of transects collected and reason for any
missed collection on the field forms.
7.4 Sample Processing in the Field
You will prepare three different types of laboratory samples from the composite index samples:
an ID/enumeration sample (to determine taxonomic composition and relative abundances), a
chlorophyll o sample, and a biomass sample (for ash-free dry mass (AFDM)). All of the methods
for processing the three samples are found in the Final Site Activities section of the manual. 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.
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8
PHYSICAL HABITAT CHARACTERIZATION
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Field measurements for physical habitat 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.
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.
8.1 Equipment and Supplies
Table 8.1 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 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 8.1 Equipment and Supplies: Physical Habitat
For making
measurements
For recording
data
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
Surveyor's telescoping leveling rod
Sounding rod
Field Operations Manual and/or laminated quick reference guide
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: Channel Constraint
1+ extras field Form: Visual Assessment Form
8.2 Summary of Methods Approach
Physical habitat in rivers includes all those physical attributes that influence or provide
sustenance to organisms within the stream. The physical habitat of a river varies naturally, thus
expectations differ even in the absence of anthropogenic disturbance. River sample reach
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lengths are defined as 40 times the wetted width at the x-site, with a minimum of 150m and
maximum of 4k, as described in Section 3. Measurement points are systematically placed to
statistically represent the entire reach. River 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. 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.
8.3 Components of the Field Habitat Assessment
Field data collection for the physical habitat assessment is accomplished in a single float down
each sampling reach.. The physical habitat methods are made up of the following components:
Thalweg profile, Littoral/Riparian Cross-Sections, and assessments of the entire reach. Table 8.2
describes the components of physical habitat in non-wadeable systems and gives an overview of
how the data is collected. Measurements are recorded on 11 copies of a two-sided field form,
and separate forms for assessing the degree of channel constraint, and recording evidence of
debris torrents or recent major flooding.
Table 8.2 Components of Non-Wadeable River Physical Habitat Protocol
1. 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.
• Record the presence of mid-channel snags
• Measure thalweg (maximum) depth using Sonar or rod
2. Littoral/Riparian Cross-Sections: at 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
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• 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.
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.
8.4 Summary of Workflow
Table 8.3 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 8.3 Summary of Workflow Physical Habitat Characterization (Non-Wadeable)
A. At the chosen bank on first transect (farthest upstream):
1. Complete the header on the front of the Channel/Riparian Transect form including the Site ID,
Date, Transect (A-K) and bank (left or right) that was assigned during reach layout.
2. Read the GPS coordinates and record them in the Transect (Bank) space on the field form.
3. Record the dominant and subdominant littoral substrate, based on visual observations.
4. Move boat in a "loop" within the 10 x 20 m littoral plot, measuring 5 littoral depths and probing
substrate.
5. Estimate dominant and subdominant littoral substrate, based on probing the 5 locations.
6. Estimate areal cover offish concealment features in the 10 x 20 meter littoral plot.
7. Tally LWD within or partially within the 10 x 20 meter littoral plot.
8. Collect densiometer measurements at bank (facing upstream, downstream, left, right).
9. Choose bank angle class, estimate bankfull height, width and channel incision. (Note that width
and incision estimates incorporate both left and right banks.).
10. Tally LWD entirely out of water but at least partially within the bankfull channel.
11. Estimate and record distance to riparian vegetation on the chosen bank.
12. 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).
13. 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", use the close rating; "C" if disturbance is present
within the riparian plot. If the item is only observed beyond (outside) the riparian plot, the
proximity code is "P". If the disturbance is not present in or adjacent to the plot, mark as "0", not
present.
14. Proceed to a midstream point on the transect and record the GPS coordinates in the Transect
(Midstream) space on the field form
15. If the next transect has not already been marked during reach layout, 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:
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1. 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.
2. 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 thalweg measurement is l/10th the distance between
transects, which can help you keep track of your downstream progress).
C. Repeat the Whole Process (for the remaining 10 transects and subreaches).
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.
8.5 Work Flow and Reach Marking
In a single midstream float down the 40 channel-width reach, the 2-person habitat crew
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) and/or a laser range finder. 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 perform cross-section, littoral, and riparian measurements, recording the actual GPS
latitude/longitude of the transect position (bank and midstream). They will also collect biological
samples at each transect.
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 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 rivers.
8.5.1 Reconnaissance for Physical Habitat Data Collection
The habitat crew will also record reconnaissance and safety notes at this time. They will inform o
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 K
the crew 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 5£
consideration multiple channels, blind channels, backwaters, alcoves, impassible riffles, rapids, u
jams, and hazards such as dams, bridges and power lines. Habitat Sampling Locations within the <
Reach 55
Measurements are made at two scales along the length of the reach; the results are later -i
aggregated for the entire reach. Figure 8.1 illustrates the locations within the reach where data y
for the different components of the physical habitat characterization are collected. Most >
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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 = l/10th the total reach length). The
thalweg profile measurements must be spaced evenly over the entire reach.
The sampling reach is 40 times the wetted width at the X- site, with a 500m minimum and 4
kilometer maximum reach. Section 3.1.1 describes the procedures for locating the X-site, or the
midpoint of the sample reach. Section 3.2 describes 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 8.1).
The thalweg profile measurements are spaced as evenly as practicable over the entire sample
reach length. They must also 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:
• Divide the reach length by 100 to set the thalweg increment distance. You will be
making 100 evenly-spaced thalweg profile measurements, 10 between each detailed
channel cross-section where littoral/riparian observations are made (the 11 transect
locations already established).
• If the thalweg is too deep or not physically possible to be measured, estimate the depth
to the best of your ability and flag it on the field form.
_ 20m
UPSTREAM END
River Flow
Riparian
Plot
v
10m
RIGHT
1 BANK
20m
DOWNSTREAM END
Figure 8.1 Littoral Riparian Plots for Characterizing Riparian Vegetation, human influences, fish cover,
littoral substrate, and littoral depths
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8.5.2 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 8.1). 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 Table 8.4. Record data on the Thalweg Profile Form as shown in Figure 8.2.
8.5.2.1 Thalweg Depth Profile
A thalweg depth profile of the entire 40 channel-width reach is approximated by a sonar or
sounding rod 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. 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.
8.5.2.2 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 8.4. 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
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 8.2). Substrate characteristics are
recorded at every thalweg depth measurement (e.g., 10 determinations between each ^
transect). In shallow, fast-water situations, where pole-dragging might be hazardous, crews will P
estimate bottom conditions the best they can visually and by using paddles and oars. If Cd
unavoidable, suspend measurements until out of Whitewater situations, but make notes and ^
appropriately flag observations concerning your best judgments of depth and substrate. <
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8.5.2.3 Channel Habitat Classification J
Classify and record channel habitat types shown in Table 8.5 at a spatial resolution of about 0.5 <
channel-widths and check presence of off-channel and backwater habitat at every 0.4 channel- m
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it as a pool unless it occupies an area about half as wide or long as the channel is wide). 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).
Table 8.4 Procedure: Thalweg Profile
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1. Record GPS coordinates (Lat/Long) midstream and at shoreline location on the Channel/Riparian
Transect Form at the transect located at the upstream end of the thalweg subreach about to be
assessed.
2. 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.
3. Complete header information on the Thalweg Profile Form, noting transect pair (up- to downstream).
4. Begin at the upstream transect (station "0" 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 may 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
a) 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.)
b) Lightly drag the rod on the river bottom to "feel" the substrate and detect snags.
c) 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.
d)Circle the appropriate "SUBSTRATE" type and record the presence/absence of "SNAGS".
e) If it is too deep to safely measure the substrate type, estimate the type based on knowledge and
surrounding measurements and flag the data.
Channel Habitat Classification
a) Classify and record the channel habitat type at increments of every 0.4 channel width.
b) Check for off-channel and backwater habitat at increments of every 0.4 channel width.
c) If channel is split by a bar or island, navigate and survey the channel with the most flow.
d) When a side channel is encountered, fill in "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.
5. Proceed downriver to the next station, and repeat the above procedures.
6. 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 transect mid-stream,
and transect shoreline locations on the Channel/Riparian Transect Form corresponding to the
downstream end of the thalweg sub-reach you just traversed.
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NOTE: if you have taken measurements at all 10 primary thalweg stations and there is still a significant
amount of river before the next transect, you should evenly space 1 or 2 additional thalweg profile stations
in the remaining area to maintain the stations as evenly spaced as possible and not miss thalweg data in the
downstream end of the subreach. Record these additional stations in the data rows labeled 10 and/or 11 on
the form.
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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• NRSA 2013/14 PHAB: THALWEG PROFILE - BOATABLE ONLY
Site ID: Date: / /
al>
•
TRANSECT: 0 A-B OB-CO C-D 0 D-E 0 E-F O F-G O G-H O H-l 0 I-J 0 J-K
SUBSTRATE CODES
BH = BEDROCK'HARDPAN (SMOOTH OR ROUGH) • (LARGER THAN A CAR)
BL = BOULDER (250 TO 4000 mm) • BASKETBALL TO CAR)
CB = COBBLE (64 TO 250 mm) • (TENNIS BALL TO BASKETBALL)
GR = COARSE TO FINE GRAVEL (2 TO 64 mm) • (LADYBUG TO TENNIS BALL)
SA = SAND (0.06 TO 2 mm) • (GRITTY - UP TO LADYBUG SIZE)
FN = SILT/ CLAY / MUCK • (NOT GRITTY)
OT = OTHER (Flag and write comment below)
CHANNEL HABITAT CODES
PO =POOI
GL = Glide
Rl = Rime
RA = Rapid
CA = Cascade
FA = Falls
OR = Dry Channel
OTHER
Off Channel = Off
Channel or
Backwater
REMEMBER: A = Upstream ond of Reach and K = Downstream end of Reach.
THALWEG PROFILE
STATION
D
1
2
3
4
i
e
7
S
a
10
11
Flag
SNAG
O
O
O
O
O
O
0
O
0
0
0
0
DEPTH (Elthsr)
UNITS: O ft O m
SONAR XX.X
POLE X.X
SUBSTRATE
Fir Inont Substnt* Cod*
for each station
O BH O BL OCB O GR
O SA O FN O OT
O BH O BL OCB O GR
QSA Q FN O°T
O BH O BL OCB O GR
O SA O FN O OT
O BH O BL Oca OGR
O SA O FN O OT
O BH O BL O CB O GR
O SA O FN O OT
O BH O BL O CB OCR
O SA O FN O OT
OBH OBU OCB OS"
O SA O FN O OT
OBH QBL QCB OGR
O SA O FN O OT
OBH OBL OCB OGR
O SA O FN O OT
O BH O BL O CB O GR
O SA O FN O OT
O BH O BL O CB O GR
O SA O FN O OT
O BH O BL O CB O GH
O SA O FN O OT
CHANNEL HABITAT
FIIHn on» Cnanml HibKat
code foraach station
O PO O GL O R| O RA
O CA O FA O DR
O PO O <3L O HI O "A
O CA OF* O DR
O PO o GL O Ri O "A
O CA O FA O DR
O PO O GL O Rl O R*
O CA O FA O DR
O PO O GL OKI O RA
O CA O FA O DR
O PO O GL O R! O RA
O CA O FA O DR
O PO O GL ORi O RA
O CA O FA O DR
O PO O GL O Rl O RA
O CA O FA O DR
OPO OGL Om ORA
O CA O FA O DR
O PO O GL O Rl O RA
O CA O FA O DR
O PO O GL O Rl O RA
O CA O FA O DR
O PO O GL O Rl O RA
O CA O FA O DR
OFF
CHAN.
O
o
o
o
o
o
0
0
0
0
0
0
FLAG
Comments
Rag codes: U = Suspect sample Fl. F2. ete. = Flag assigned by Held crew. Explain all flags in comment sections.
O 04/15/2013 2013 PhabThalweg Profile -Beatable 6700391398 £
Figure 8.2 Thalweg Profile Form
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Table 8.5 Channel Unit Categories Used on Thalweg Form
Class (Code)3 Description
Pools (PO)
Glide (GL)
Riffle (Rl)
Rapid (RA)
Cascade (CA)
Falls (FA)
Dry channel (DR)
Off-channel
Still water, low velocity, smooth, surface, deep compared to other parts of channel
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 & very turbulent over steep channel bottom. Most of the water
surface is broken in short, irregular plunges, mostly Whitewater. Sound: roaring.
Free falling water over vertical or near vertical drop into plunge, water turbulent and
white over high falls. Sound: splash to roar. (Don't navigate raft over a waterfall!).
No water in the channel.
Side-channels, sloughs, backwaters, and alcoves separated from the main channel.
0 In order for a channel habitat unit to be distinguished, it must be at least 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 8.6.3 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, fill in the "Y" bubble on the Thalweg Profile form in the "Off-
Channel" column. These notations 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" notation 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" notations and indicate in the comments section that the feature is
a side channel.
8.6 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 (Figure 8.3
and Figure 8.7).
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NRSA 2013/14 PHAB: CHANNEL/RIPARIAN TRANSECT (Front) - BOATABLE ONLY
Site ID:
Date:
TRANSECT: O A OB OC OD OE OF OG OH Ol OJ OK
Chosen bank side:
O Lefl O RigM
Midstream Decimal Degrees Latitude
Bank Decimal Degrees Latitude
Longitude
Longitude
•LITTORAL" and SHORELINE SUBSTRATE INFORMATION
BOTTOH SUBSTRATE FROM (X ONE):
Judgement or- O °BS. @ 5 Lilloral Depths
Flag
Bedrock (Smooth) -(Largerthan a car)
RR = Bedrock ( Rough) - (Larger than a car)
XB = Large Boulder 11000 to 4000 mm) - (Meterstick to car)
SB - Small Boulder (250 to 1000 rnm) - (Basketball to Meterstick)
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 mm) • (Ladybug to marble)
SA = Sand |0.06to 2 mm)-(Gritty-up to Ladybug size)
FN = Silt I Clay I Muck - (Not Gritty)
HP- Hard pan - (Firm. Consolidated Fine Substrate)
WD = Wood - (Any Size)
QT - Other (Write comment below)
LARGE WOODY DEBRIS
[10 * 20m PW) Ts:V eacn pi*cr
FiLL IN IF UNMARKED BOXES ARE ZERO
O
FLAG:
DIAMETER
LARGE END
O.tvoem
Wood AWPirt In w«t*d Ch»nn»l
Length 5-15m
15-30fn
Length 5-15m
Dry by Aiyp»n m Banfcftjll C
15-30m
SLOPE/BEARING/DISTANCE(Optlonal): SlopeandBearingnotdeterminedlusemap)
Determine slope if feasible In terms of lime and distances. Record GPS coordinates it practical.
INTENDED Iransecl ACTUAL transect
spacing xxx (m): spacing xxx (m):
Supplemental Waypolnls
DEPTH O (I O m
BANK CHARACTERISTICS
Wetted Width
Bar Width
Banktull Width
Banhfull Height
Incised Height
BANK
ANGLE
CIRCLE ONE
Ov
OS
OG
OF
X.XX (m| FLAG
Backsltt
Slope Bearing Distance
XXX % 0-334 (m|
Way
Point *
1ST
2ND
3RD
Flag
GP5 Latitude - decimal degrees
GPS LongiliKJe - decimal degrees
Hag
Flag Codes U = suspect or norr-standard measurement; F1, F2 etc = flags assigned by each Reid crew Explain all flags in comments section
04/08/2013 2013 Phab Channel Riparian - Boat able (Front) 7231169289
Figure 8.3 Channel/Riparian Transect Form (front)
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8.6.1 Channel Margin Depth and Substrate
Channel margin depths are measured along the designated shoreline at each transect within a
10m x 20m littoral plot that is centered on the transect (Figure 8.4). 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 8.6. Record these
measurements on the Channel/Riparian Transect Form as shown in Figure 8.3. 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 8.6.
"~ "zP&fre-r- tft*.
Riparian
Plot
LEFT
10m BANK
Bankfull line
WrtSEaOM&tftrsi-
Thalweg Profile
r River Flow ^—z*,
\
Riparian
Plot
Vj&ffltys jfHfttr?? f f~/ ^--
10m
RIGHT
1 BANK
20 m
Figure 8.4 Riparian Zone and Instream Fish Cover Plots for a River Cross-Section Transect
Table 8.6 Procedure: Channel Margin Depth and Substrate
1.
2.
3.
Fill in the header information on page 1 of a Channel/Riparian Transect Form. Be sure to indicate the
letter designating the transect location. Indicate the assessed bank (left or right) as designated
during reach layout activities. Also ensure that GPS coordinates have been recorded for each
transect at the bank and midstream locations.
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 10m x 20m plot as well as on the
shoreline 1m 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
Size Class
Bedrock (Smooth)
Bedrock (Rough)
Large Boulders
Size Range (mm)
>4000
>4000
>1000 to 4000
Description
Smooth surface rock bigger than a car
Rough surface rock bigger than a car
Meter stick to Car size
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4.
5. Re
SB
CB
GC
GF
SA
FN
HP
WD
OT
Small Boulders
Cobbles
Gravel (Coarse)
Gravel (Fine)
Sand
Fines
Hardpan
Wood
Other
>250 to 1000
>64 to 250
>16 to 64
> 2 to 16
>0.06 to 2
<0.06
Regardless of Size
Regardless of 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, indicate the appropriate shore and bottom
substrate type and record the depth measurements ("SONAR" or "POLE" columns).
jpeat Steps 1 through 4 at each new cross-section transect.
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8.6.2 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 8.7. For each tally (Wood All/Part in
Wetted Channel and Dry but All/Part in Bankfull Channel), the field form (Figure 8.3) 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. Tally all LWD that is within each 10x20
littoral plot. Do not tally woody debris in the area between channel cross-sections, but the
presence and location of 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 8.3) 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.
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Table 8.7 Procedure: Tallying Large Woody Debris
Note: Tally pieces of large woody debris (L WD) within the 11 transect littoral plots of the river reach at
the same time the shoreline measurements are being determined. Include all pieces whose large end is
located within the littoral 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 sampling reach. These
plots are each 20 m long in the upstream-downstream direction (10m up, 10m down from the
transect). They are positioned along the chosen bank and extend from the shore 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 [1 ft.]; length 5 m [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 for a 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.
8.6.3 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 8.3). 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 8.3). The procedures for obtaining bank angle and
measurements of channel cross-section morphology are presented in Table 8.8.
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-
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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 8.8 Procedure: Bank Angle and Channel Cross-Section
Bank Angle and Cross Section Methods
1. Record the wetted width of the river and the width of exposed mid-channel bars (if present) in the
BANK CHARACTERISTICS section of the field data form. Also determine the bankfull channel width.
2. 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. Mark the angle in the
"BANK ANGLES" section of the Channel/Riparian Transect Form.
3. 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.
4. 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.
5. Repeat Steps 1 through 4 at each cross-section transect. 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 the NRSA. 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
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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 deposi-
tional 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 8.5) 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. ^
Q
If the channel is not greatly incised, bankfull channel height and incision height will be the same. H
However, if the channel is incised greatly, the bankfull level will be below the level of the first Cd
terrace of the valley floodplain, making "Bankfull Height" smaller than "Incision" (Figure 8.6). t|
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 u
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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 back of the Channel/Riparian Transect form (Figure 8.7) regarding the
relationships among channel incision, valley sideslope, and width of the valley floodplain. Mark
whether you could or could not readily see over the bank. You will also do an overall assessment
of channel constraint for the whole river reach; see Section 8.12 for a discussion of constraint
concepts and assessment procedures.
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
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Downcutting over
geologic time
Former second
terrace becomes
Fnrmer firc-t third terrace
Former active floodplain
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
>
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Figure 8.5 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)
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A) Deeply Incised Channel
Hill Slope
Incision Height (Always
equal to or greater than
bankfull height)
Second Terrace
First Terrace
From Figure 7-5 (B)
Dwfi cuffing cv«r
51
/
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,—Bankfull
/ Height
/ (When
/ channel form
' is not a good
indicator, use
recent
floodina)
B) Small stream constrained in V-shaped valley
Bankfull Height
(when channel form is
not a good indicator,
use evidence of recent
flooding, lack of
permanent flood-
intolerant vegetation)
Incision Height=
Bankful Height
No incision:
No evidence of
downcutting,
vertical bank
angle, etc.)
Figure 8.6 Determining Bankfull 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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Rtvtnvtd by [initial):_
NRSA 2013/14 PHAB: CHANNEL/RIPARIAN TRANSECT (Back) - BOATABLE ONLY
Site ID:
Date:
TRANSECT: O A OB OC OD OE OF O6 OH Ol OJ
/
OK
Chosen bank side:
: r .r;.: .-. !•- B •
O Left O Right
VISUAL RIPARIAN ESTIMATES
0= Absent (()•%] 1 = Sparse (*1Cr%J 2 = Mod*m*i )10-4(hi,[ 3 = Heavy [40-761*,)
4 = VcryHcavy (>75*X.)
O = 0*ciduous C = Conifer-out E = B*e>**»«f Ev*rflr«*n M = Mlxad N = Mont
RIPARIAN VEGETATION COVER
Canopy (>5 m high) Left Bank
Woody Vegelaoon Type Q O O O O
BIG Trees (Trunk
SMALL Trees (Trunk
«0.3mDBHI
OOOOO
Flag
OOOOO
Rlahl Bank
ooooo
ooooo
Flag
Underslory (0.5 lo 5 m high)
Woody Vegetation Type
Woody Sfiruds & Saplings
Non-Woody Herts,
Grasses. & Forbs
Ground Cover (<0.5 m
woooy Shrubs
.'. saplings
Non-woody Hems,
Grasses and Forbs
Barren, Bare Din
orOuff
ooooo
ooooo
ooooo
high)
ooooo
ooooo
ooooo
ooooo
ooooo
ooooo
ooooo
ooooo
ooooo
HUMAN INFLUENCE 0= Not Present
>10m C = IrVllritn 10 mplot B = On Bank
wall, Dik«.'R*v*r ne m
Buildings
Pipes (InletVOutirtj
Undfi ID Trash
PastureiHangeJH ay Field
Logging Operations
Flag
0000
0000
0000
0000
0000
0000
O O O O
O O O O
O O O O
0000
Mining Activity Q O O O
0000
0000
0000
0000
0000
0000
O O O O
O O O O
O O O O
0000
O O O O
FISH COVER/OTHER (10m x 20m Plot)
0 = Absent (0%) 1
<<10%| 2 = Mod*r«t«
ryRf avy [»76*.J
In-channal Cover
Flag
Woody Debris >0.3 m |BIC|
Brush/Woody Debris <0.3 m (SMALL)
Live Trees In Strean
Overtianglng Veg. =<1 m oT Surface
Undercut Banks
Boulders/Ledges
OQOQQ
OOOOO
ooooo
ooooo
ooooo
OOQOQ
OQQOQ
00000
Artificial Slmctures Q O O O O
CHANNEL CONSTRAINT
Distance from shore to riparian vegetation (m) XXX
Mark only one:
O Channel is Constrained
O Channel isin Broad Valley but constrained byincision
O Channel isin Narrow Valley but NOT very constrained
O Channel is Unconstrained in Broad Valley
Flag
Mark only one:
O Yes. I could readily see over the bank
O No, I could not readily see over the hank
CANOPY DENSITY @ BANK
DENSIOMETER <0-17Max)
Flag
Flag Codes U = suspect or non-standard measurement: Fl F2. etc, = flags assigned by each field crew. Explain all flags in comments section.
04/08/2013 2013 Phab Channel Riparian - Boatable (Back) 7154355572
Figure 8.7 Channel/Riparian Transect Form, page 2 (back side).
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8.7 Visual Riparian Estimates
8.7.1 Riparian Vegetation Structure
Riparian vegetation observations apply to the riparian area upstream 10 m and downstream 10
m 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 8.1). The riparian plot dimensions are
estimated, not measured. Table 8.9 presents the procedure for characterizing riparian
vegetation structure and composition. Figure 8.7 illustrates how measurement data are
recorded in the "Visual Riparian Estimates" section of the Channel/Riparian Transect Form
(back).
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.
Table 8.9 Procedure: Characterizing Riparian Vegetation Structure
Riparian Vegetative Cover
1. Anchor or tie up at the river margin at a cross-section transect; then make the following observa-
tions 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 JDeciduous, Coniferous, broadleaf £vergreen, JVHxed, 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 "None". Indicate the appropriate vegetation type in the "VISUAL RIPARIAN
ESTIMATES" section of the Channel/Riparian Transect 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 "None".
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.
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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", but no more than one cover type
could receive a rating of 4.
8.8 Instream Fish Cover, Algae, and Aquatic Macrophytes
Over a defined length and distance from shore at the sampling locations (Figure 8.4), crews shall
estimate by eye and by sounding the proportional cover of fish cover features and trophic level
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 8.10. Record
data in the "Fish Cover/Other" section of the Channel/Riparian Transect Form as shown in
Figure 8.8. 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 only on the side of the river
previously chosen for assessment during reach layout (Figure 8.1). The areal cover classes offish
concealment and other features are the same as those described for riparian vegetation
(Section 8.8.1).
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, which 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
O debris tally [Section 8.7.2]). 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
tl 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 (e.g., cars or tires) or purposefully placed for diversion, impoundment, channel stabilization, or
^
in other purposes. Record the cover of these structures on the form
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Table 8.10 Procedure: 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 of fish 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,
l=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.
8.9 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 Figure
8.8. Record human influences on the Channel/Riparian Transect Form (Figure 8.7). Relate your
observations and proximity evaluations to the river and riparian area within 10 m upstream and
10 m downstream from the transect (Figure 8.8). Four proximity classes are used: In the river or
on the bank within 10 m upstream or downstream of the cross-section transect, present within
the 10 m x 20 m 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 8.7. 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).
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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C—within riparian plot
B—on bank or in stream
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P—outside plot
(but do not sight through next
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Figure 8.8 Proximity Classes for Human Influences in Non-Wadeable Rivers
Table 8.11 Procedure: Estimating Human Influence
Human Influence
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.
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).
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.
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 10 m stream segment and located in the stream
or on the stream bank.
C (Close) Present within the 10 x 20 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 the 20 m stream segment or the riparian
plot area at the transect
Repeat Steps 1 through 4 for the opposite bank.
Repeat Steps 1 through 5 for each cross-section transect, recording data for each transect on a
separate field form.
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8.10 Canopy Cover Measurements
Canopy cover over the river is determined at each of the 11 cross-section transects. A spherical
densiometer (model A convex type) is used (Lemmon 1957) and is provided in the base kit to
each crew. Mark the densiometer with a permanent marker or tape exactly as shown in Figure
8.9 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). Measure vegetative cover over the reach at the chosen bank at each of the
11 transects (A-K). 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 8.10. 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 8.9. 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 four measurement points, record the number of intersection
points (maximum=17) that have vegetation covering them in the CANOPY DENSITY @ BANK section
of the Channel/Riparian Cross-section Form as shown in Figure 8.7.
TAPE
o
1
BUBBLE LEVELED"
Figure 8.9 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."
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Table 8.12 Procedure: Canopy Cover Measurements
Canopy Cover Methods
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.
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8.11 Channel Constraint Assessment, Debris Torrents and Recent
Floods
8.11.1 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 8.13. Record data on the Channel
Constraint Assessment Form (Figure 8.10). First, classify the stream reach channel pattern as
predominantly a single channel, an anastomosing channel, or a braided channel (Figure 8.11J:
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
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blocked by vegetation), record the distance you can see and mark the appropriate bubble on the
field form.
Table 8.13 Procedure: Assessing Channel Constraint
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.
1. Classify the stream reach channel pattern as predominantly a single channel, an
anastomosing 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.
2. 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.
3. 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.).
4. Based on your determinations from Steps 1 through 3, select and record one of the
constraint classes shown on the Channel Constraint Form.
5. Estimate the percent of the channel margin in contact with constraining features (for
unconstrained channels, this is 0%). Record this value on the Channel Constraint Form.
6. 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
bubble on the field form.
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NRSA 2013/14 CHANNEL CONSTRAINT
Date: / I
CHANNEL PATTERN (Fill in one):
O One Channel
O Anastomosing (complex) channel - (Relatival/ 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(FMI in one):
O Channel very constrained in V-shaped valley (i.e. rt 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 Inasion
(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 fill off-channel areas and side channels,
spread out over flood plain, or easily cut new channels by erosion)
CONSTRAINING FEATURES (Fill in one):
O Bedrock (i e channel is a bedrock-dominated gorge)
O Hillslope (i.e. channel constrained in narrow V-shaped valley)
O 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, dike, road, etc.)
O No constraining features
Percent of channel length with margin
in contact with constraining feature: >
(0-100%)
Bankfull width:
(m)
Valley width (Visual Estimated Average): im\
Note- Be sure to include distances between both sides of valley border for valley width
If you cannot see the valley borders, record the distance /"j
you can see and fill this bubble. *"*
Percent of Channel Margin Examples
100%
50%
100%
50%
COMMENTS
04/08/2013 2013 Channel Constraint
1742268066
Figure 8.10 Channel Constraint 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.
Figure 8.11 Types of Multiple Channel Patterns
8.11.2 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.
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,
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including its substrate, banks, and riparian corridor, checking the presence of features described
on the Torrent Evidence Assessment Form (Figure 8.12). 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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Site ID:
NRSA 2013/14 TORRENT EVIDENCE ASSESSMENT
Date: / /
RutlomdbyltnUlBj] _
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 How channel. This
corridor lacks riparian vegetation with possible exception of fireweed, even-aged alder or cottonwood seedlings, grasses,
or other herbaceous plants.
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.") In
a torrent scour or deposition channel, the stones are laying in unorganized patterns, lying "every which way." In addition
many of the substrate particles are angular (not "water-worn.")
03 - Channel has little evidence of pool-riffle structure. (For example, could you ride a mountain bite down the channel?)
04 - The stream channel is scoured down to bedrock for substantial portion of reach.
05 - There are gravel or cobble berms (little levees) above bankfull level.
06 - Downstream of the scoured reach (possibly several miles), there are massive deposits of sediment, logs, and other
debris.
07 - Riparian trees have fresh bark scars at many points along the stream at seemingly unbelievable heights above the
channel bed.
08 - Riparian trees have fallen into the channel as a result of scouring near their roots.
EVIDENCE OF TORRENT DEPOSITS:
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 moved by the stream at even extreme flood stage.
10 - If the stream has begun to erode newly laid deposits, it is evident that these deposits are "MATRIX SUPPORTED."
This means that the large particles, like boulders and 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 "fill-in" the interstices between coarser particles.)
NO EVIDENCE:
11 - No evidence of torrent scouring or torrent deposits.
COMMENTS
04/08/2013 2013 Torrent Evidence
3362134147
Figure 8.12 Torrent Evidence Form
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9 FECAL INDICATOR (ENTEROCOCCI)
9.1 Summary of Method
Collect a fecal indicator sample at the last transect (Transect K) after all other sampling is
completed. 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. Following collection, place the sample in a cooler and maintain on ice prior to
filtration of two 50 ml volumes. Again, 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 9.1).
9.2 Equipment and Supplies
Table 9.1 provides the equipment and supplies needed to collect the fecal indicator sample.
Record the sample data on the Sample Collection Form (Figure 4.2).
Table 9.1 Equipment and Supplies: Fecal Indicator Sampling (Non-Wadeable Sites)
For collecting samples
For recording measurements
nitrile gloves
pre-sterilized, 250 ml sample bottle
sodium thiosulfate tablet
Wet ice
cooler
Sample Collection Form
Fecal Indicator sample labels (2 vial labels and 1 bag label)
Pencils (for data forms)
Fine tipped indelible markers (for labels)
Clear tape strips
9.3 Sampling Procedure
The procedure for collecting the fecal indicator sample is presented in Table 9.2.
Table 9.2 Procedure: Fecal Indicator (Enterococci) Sample Collection (Non-Wadeable Sites)
Enterococci Sample
1. Put on sterile, 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 uncapped, inverted 250 ml sample bottle to a depth of 1 foot below the water surface,
avoiding surface scum, vegetation, and substrates.
4. 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.
5. If the depth does not reach 0.3m along transect at 1 m from the bank, take the sample and flag it on
the field form.
6. After removing the container from the water, discard a small portion of the sample to allow for
proper mixing before filtering.
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7. Add the sodium thiosulfate tablet, cap, and shake bottle 25 times.
8. Store the sample in a cooler on ice to chill (do not freeze immediately). Chill for at least 15 minutes.
9. Sample must be filtered and all filters frozen within 6 hours of collection.
9.4 Sample Processing in the Field
You will need to process two separate filters for the Enterococci sample. All the filters required
for an individual site should be sealed in plastic bags until use to avoid external sources of
contamination. Please refer to Section 13.3 for information regarding processing the Enterococci
samples.
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NRSA 2013/14 ASSESSMENT (Front)
Date: /
RtilivndbytaiUalr.
Elevation at transect K:
Oft
WATERSHED ACTIVITIES AND DISTURBANCES
(Intensity: HlanK=Noc observed. L=Low. MsModerate. H=Heavy)
OBSERVED
BLANK FIELD INDICATES ABSENCE:
O
O O O Restdencas
O O O MamtamedLamis
O G O Construction
O O O PiP'S- Dr«m
O O O Oumfing
O O O Roads
O O O Bridges/Causeway
O O O Sewage Treatment
Recreational
O © O H.kmgTuili
O O O Parts. Campgrounds
O 0 O PrimitiveParks. Camping
O O O TrasnlUWr
O O O Surface Films Scums.
or Slicks
Agricultural
O O O Cropland
O O 0 Pasture
O © 0 Livestock Use
O 0 O Oichar*
O 0 0 Poultry
O O O Feedlot
O 0 O ••"'•<"<• '-'-i i ;.>•.•/.•
Industrial
O ©
O 0
0 InduslrialPJants
0 Mines/Quarries
0 © 0 OiyGas Wells
O 0 0 Power Plants
O 0 0 Loggmg
O 0 O Evidence of Fire
O 0 O oi"*
O O O Commercial
Stream Management
O 0 O Limmg
O 0 0 ChemicalTreatment
O 0 0 AngtngPressure
O 0 O Dredgons
0 0 O Channelization
O 0 O W«ter Level Fluctuations
O © O Fl5n Stock.no,
O © O Dafns
SITE CHARACTERISTICS (200m radius)
WATERBODY CHARACTER
PRISTINE: OS O^ O3 O2 O1 Highly Disturbed
APPEALING: O 5 O4 O3 O2 Q1 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 'X' O Forest O Agriculture O Range O Urban O SuburbanfTown
If Forest, Dominant Age Class O 0 - 25 yrs O 25 - 75 yrs O > 75 yis.
WEATHER
CONDITIONS AND LOCAL CONTACTS
OBSERVATIONS (e.g. accessibility, boating, fishing, swimming, health concerns):
04/OS/2013 2013 Assessment
0351596807
Figure 9.1 Site Assessment Form
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10 FISH ASSEMBLAGE
10.1 Summary of Method
The fish sampling method is designed to provide a representative sample of the fish community,
collecting all but the rarest taxa inhabiting the site. It is intended to accurately represent species
richness, species guilds, relative abundance, size, and presence of anomalies. The intended uses
of the fish assemblage data are to calculate predictive models of multimetric indicators (MMIs;
similar to an Index of Biotic Integrity [IBI]; Pont et al. 2009, USEPA 2013a) and possibly
Observed/Expected (O/E) taxa richness. In addition, the fish assemblage data provides a starting
point for developing potential indicators of ecosystem services related to fish.
In non-wadeable rivers, collect fish using boat (or raft) electrofishing over a defined sampling
reach ( 40 times the wetted width) within the support reach established for the site. Use
secondary fish collection methods in habitat that cannot be adequately sampled by boat.
Secondary methods may include backpack or tote barge electrofishing, using your boat as a
barge, or seining as a last option only if conductivity is too high for electrofishing. Conduct
sampling in a downstream direction, allocating effort (button time) within subreaches (areas
between the cross-section transects). At medium and large wadeable streams, if you have not
collected 500 individuals at the end of the defined fish sampling reach, sample additional
subreaches in their entirety until you obtain at least 500 individuals. Record information related
to sampling effort on the front of the Fish Gear and Sample Information Form (Figure 10.1).
Record species identification and enumeration data on one or more pages of the fish collection
form (Figure 10.2).
There are numerous revisions and clarifications to the non-wadeable sampling protocol from
that used in the NRSA 2008/09. Separate collection forms for each subreach are no longer
required. The layout of the fish sampling reach and how to allocate effort across the reach is
clarified. Guidance is provided to use supplemental gear to more effectively sample areas
inaccessible to boats or rafts. Guidance is provided to deal with irruptive species (e.g., shad,
certain shiners, etc.) that may artificially skew count and relative abundance results. The fish
gear form is modified to include the specific protocol used to sample (beatable, large wadeable,
or wadeable), clarify reasons for not sampling, and include information for any secondary gear
used. The collection form is modified to include information about introduced species and to
break down total counts by major size classes (to begin to look at ecosystem service related
indicators). Measuring minimum and maximum lengths for each species collected is not
required. An additional form to record seining effort information is used to evaluate the
sufficiency of the seining results and the ability to combine them with electrofishing data at a
site. The procedure is clarified for preparing voucher specimens to distinguish unknown/range
extension vouchers (those taken back to a local facility for identification) and QA vouchers (a
sample of all species from a site sent to an external facility for confirmation of all field
identifications). Guidance for recording unknown taxa is clarified to minimize ambiguity. The
procedure to update species identifications based on the results from unknown/range extension
voucher specimens is clarified and uses an additional data form.
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g NRSA 2013/14 FISH GEAR AND SAMPLING INI
Site ID:
FISH SAMPLING PROTOCOL (select one):
O BOATABLE(20to40 Channel Widths (CW))
O LG. WADEABLE • (>=12.5 m wide) (500 m to 40 C\
O WADEABLE - (<\2S mwide) |40 CW)
Final Length of
Fishing Reach (m):
O Fast flowing high gradient site Sampli
FISH GEAR INFORMATION
WaterVisibllity: O Good O Poor
Primaiy Electrofishmg Gear
8ofNetters(1):
Anodes
O RAFT (No motor) Number
O BACKPACK
Diameters:
O BANK OR TOWED &ARGE
Secondary Electrofishing Gear
# of Metiers (1):
Anodes
ORAFT(Nomotor) Number
O BACKPACK
Diameters:
O BANK OR TOWED BARGE
-ORMATION (Front) H
Date: / /
FISH SAMPLING - NOT CONDUCTED OR SUSPENDED (select one):
O Fished - None Collected
V) O Not Fished/Fishing suspended - Cant sample >/= 50% of
required reach:
-Boa table (10 CW)
• Lg. Wacleable (250m or 10 CW, whichever is longer)
- Wadeable (20 CW)
ng ProtocolComments:
O Not Fished • No Permit
O Not Fished - Site Conditions Prohibit Sampling (Describe in comments)
O Fishing Suspended - Permit Restriction (Listed species encountered)
O Not Fished - Equipment Failure
O Not Fished -Other Explain:
Water Temp ("CJ: Cond(uSfcm):
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Total Shock (button) Time (s):
Total FishingTime(min):
% of Fishing Reach Sampled:
Pulse Width (ms):
Total Shock (button) Time (s):
!: Total Fishing Time (min):
% of Fishing Reach Sampled:
Primary Seine Net: O BAG SEINE Q MINNOW SEINE No. of crew members:
Height(rn): Length(m): Mesh(mm): Lenoth^ml' Hauls' ^Thrie'fminV V, of Fishing Reach Sampled:
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_ NRSA 2013/14 FISH COLLECTION
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FISH ASSEMBLAGE
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10.2 Equipment and Supplies
Table 10.1 shows the checklist of equipment and supplies required to complete the non-
wadeable fish sampling. 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.
Table 10.1 Equipment and Supplies: Fish Sampling (Non-Wadeable Sites)
For collecting
fish
Boat, motor, and trailer (and necessary
safety equipment)
Gasoline and oil (if using a 2 cycle
generator)
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
Pulsator control box (5.0 GPP or less)
Backpack electrofisher (as used for
wadeable streams)
Dip nets (non-conductive handles) %" mesh
Scientific collection permit(s)
10 ft x 6 ft minnow or bag seine with % in.
mesh (additional 4 ft depth seine may also
be used)
GPS with transect waypoints
preloaded
Several Leak proof HOPE jars for fish
voucher specimens (various sizes
from250mL-4L)
1 scalpel for slitting open large fish
before preservation
1 container of 10% buffered formalin
1 aquarium net for dipping small fish
from live well
2 measuring boards (3 cm size
classes) (optional; needed only if
quantitative length data are needed)
1 set Fish ID keys
Field Operations Manual and/or
laminated Quick Reference Guide
Digital camera with extra memory
card & battery
20 ft x 6 ft minnow or bag seine with
% inch mesh (additional 4 ft depth
seine may also be used)
Polarized sunglasses and hats
For recording Sheet of sample labels and voucher
measurements specimen tags (for unknown/range
extension voucher samples)
Sheet of sample labels and voucher
specimen tags (for QA voucher samples)
Fish Gear and Information form
Fish collection forms (several per site,
depending on expected species richness)
Seining information form
Clear tape strips
Soft (#2) lead pencils
Fine-tip indelible markers
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10.3 Sampling Procedures
Table 10.2 describes the procedure for collecting fish in non-wadeable streams. The sampling
crew should consist of one boat operator (who also controls the electrofishing unit) and one dip
netter (equipped with a 1/4" mesh dip net) situated at the bow. Begin sampling at the upstream
end of the support reach defined for the site and proceed downstream.
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The reach length sampled for fish varies based on the width of the river used to establish the
length of the support reach and on the number of individuals collected (Figure 10.3). In very
small rivers (< 12.5 m wide), conduct sampling in the same fashion as for small wadeable
streams, sampling the entire support reach and moving the boat (or raft) within each subreach
to sample both shorelines as well as the mid-channel.
In medium rivers (mean CW between 12.5 and 25 meters) the minimum length for fish sampling
is 500 m, which will include between 5 and 10 subreaches. If you reach the required distance
within a subreach, extend the length of the fish sampling reach to the end of the subreach so
you end fishing efforts at a transect. For large rivers (mean CW > 25 meters), the minimum
length for fish sampling is 5 of the 10 subreaches. In both medium and large rivers, if a
minimum of 500 fish are not collected after sampling the minimum fishing reach, additional
subreaches will be sampled until 500 fish are collected or all 10 subreaches have been sampled.
Table 10.2 summarizes the fishing protocols for each of three sizes of non-wadeable rivers.
For rivers > 12.5 m wide, restrict sampling to shoreline habitats within each subreach. Start on
the same bank as the habitat crew, and move to the opposite bank after every two subreaches.
Within each subreach, sample for ~700 seconds of "button time." Prior to sampling each
subreach, determine the most appropriate gear for the subreach (e.g., boat or raft vs. barge or
backpack electrofishing units, or seines if the conductivity is too high). When electrofishing,
proceed downstream at a pace equal to or slightly greater than the prevailing current to
maximize capture efficiency. Maneuver the electrofishing unit in and around complex habitat
when necessary; however, use discretion in sampling these habitats to maintain equal effort
among subreaches.
Whenever possible, process fish at the end of each subreach. You can use multiple lines per
species on the fish collection form if necessary (e.g., you collect a large number of individuals
and need additional space for tallying, or collect the same species at non-adjacent subreaches
[e.g., A-BandG-H]).
If seining is used, record fish collected with seining protocols on a separate line.
At the end of the designated fish sampling reach, determine if you have collected at least 500
individuals. If so, stop sampling. If not, sample additional subreaches (one at a time) until at
least 500 individuals are captured. Stop sampling when you reach Transect K (the end of the
entire support reach), regardless of the number of individuals collected. Once the decision is
made to fish an additional subreach, it should be completely fished as described above (do not
stop sampling partway through a subreach).
10.3.1 Irruptive Species
For the purposes of NRSA, the term irruptive species will be used to describe fish species which
are found in locally abundant "patches" in one or two small places within the sampling reach.
These are distinct from dominant species which are in abundance throughout most of the reach.
As such, irruptive species may artificially skew necessary effort to reach 500 individuals; and, if
included the overall assemblage counts, may artificially skew the calculations of relative LU
abundance offish species in the reach. To avoid the impact of irruptive species, move quickly <
through large isolated schools of a single species (e.g., shad, certain shiners, etc.). Also, when £
tallying total fish at the end of the designated fish sampling reach, calculate the percentage of ^
irruptive species to total individuals captured. If any single irruptive species comprises greater <
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than 50% of the total sample, continue fishing one or more additional subreaches until the
percentage of the irruptive species decreases to less than 50%.
Table 10.2 Summary of Non-wadeable Fishing Protocols
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Small Non-Wadeable (less than 12.5 meters wide)
Sampling reach will be between 150 and 500 meters
Subreaches will be between 15 and 50 meters each
•S Sample all 10 subreaches in their entirety from bank to bank starting at Transect A
S Total button time will range from 500-700 seconds per subreach
• You do not have to expend equal button time among the 10 subreaches—you can devote
more button time to subreaches with more complex habitat.
•S No minimum fish number
Medium Non-Wadeable (12.5 to 25 meters wide)
Sampling reach will be between 500 and 1000 meters
Subreaches will be between 50 and 100 meters each
Minimum fishing length = 500 meters which will between 5 and 10 subreaches
If needed, extend fishing length to end at a transect
•S Fish each subreach along bank in pairs of subreaches starting at the random PHab bank at
Transect A
•S Button time is roughly 700 seconds per subreach
• Depending upon the habitat complexity, you can vary the distance actively fished to
allocate the available button time throughout the subreach.
S Minimum fish number is 500 unless all 10 subreaches have been fished.
> After fishing at least 500 meters, if 500 fish have not been collected, add subreaches
one at a time (but fish them in their entirety) until 500 fish are collected or all 10
subreaches have been fished.
Large Non-Wadeable (25 + meters wide)
Sampling reach will be between 1000 and 4000 meters
Subreaches will be between 100 and 400 meters each
Minimum fishing length = 5 subreaches (which will equal between 500 and 2000 meters)
S Fish each subreach along bank in pairs of subreaches starting at the random PHab bank at
Transect A
S Button time is roughly 700 seconds per subreach
• Depending upon the habitat complexity, you can vary the distance actively fished to
allocate the available button time throughout the subreach.
S Minimum fish number is 500 unless all 10 subreaches have been fished.
> After fishing 5 subreaches, if 500 fish have not been collected, add subreaches one at a
time (but fish them in their entirety) until 500 fish are collected or all 10 subreaches
have been fished.
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Small Non-wadeable River: Channel Width < 12.5 m
< Fish Entire Reach (40 x Channel Width)
K
Medium Non-wadeable River: Channel Width 12.5 m to 25 m
B
A
Fish a minimum of 500 m*, but do not
stop in middle of a subreach
K
Example: If average CW = 20 meters,
each subreach equals 80 meters.
500 meters would fall between Transects G and H.
Initial fishing reach would stop at Transect H and equal 560 meters.
Large Non-wadeable River: Channel Width > 25 m
Fish a minimum of 5
subreaches*
(20 x Channel Width)
K
*At medium & large rivers, if < 500 individuals have been collected after minimum sampling reach, continue fishing to
next transect (alternating banks) until 500 individuals are collected or Transect K is reached, (10 subreaches fished)
Figure 10.3 Reach Layouts for Fish Sampling at Non-Wadeable Sites
Dark shaded areas indicate the minimum length of the fish sampling reach. Light shaded areas are
sampled as needed to meet the required 500 individuals.
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1) Table 10.3 Procedure: Electro-fishing (Non-Wadeable SitesjComplete the header section of
the fish gear form (Site ID and date).
2) Decide if you will be able to sample the site.
a) 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. If you cannot sample at all because of permit restrictions, mark Wot
Fished - No Permit.
b) If site conditions prevent boat or raft electrofishing (e.g., no access for boat or raft, safety
concerns, conductivity is too high or too low to use a boat electrofishing unit), determine if
you can sample with the use of secondary methods (backpack or barge shocker, or by
seining).
i) If yes, follow the procedures presented in Section 10.3.2 and/or Section 10.3.3.
ii) If not, mark Wot Fished - Site Conditions Prohibit Sampling. Note the conditions in the
Sampling Protocol Comments.
c) If you determine that > 50% of the required fish sampling reach is physically inaccessible or
otherwise unsampleable by any means, mark Wot Fished/Fishing Suspended - Can't sample
>=50% of required reach.
d) If you cannot sample because of equipment problems, mark Wot Fished - Equipment
Failure.
e) If you cannot sample for any other reason, mark Wot Fished - Other and note the reason in
the Explain field.
3) If you can begin to sample, mark Bootable in the Fish Sampling Protocol section. For safety,
everyone must wear personal floatation devices, foot protection, and insulated linesman's gloves
a) To aid vision while netting fish, wear polarized sunglasses and a hat or visor.
4) Determine the minimum length for the fish sampling reach based on the width used to define the
support reach for the river (recorded on the stream verification form).
a) If the width is < 12.5 m, the fish sampling reach length is the same as the support reach (40
channel widths).
b) If the width is between 12.5 and 25 m, the minimum fish sampling reach length is -500 m. If
500 m falls within a subreach, extend the length to the end of the subreach.
c) If the width is > 25 m, the minimum fish sampling reach length is 20 channel widths (5
subreaches).
5) Mark the appropriate Water Visibility conditions on the form. Poor implies that your ability to
electrofish effectively is compromised because of poor visibility. Record the water temperature
and conductivity.
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6) Mark either Boat or Raft as the Primary Electrofishing Gear. Boats use outboard motors and can
travel upstream, while rafts are oar-powered and generally cannot move upstream.
a) Only one netter should be used. Record this along with the number of anodes, and their
diameter (mark the units as either inches or centimeters) on the fish gear form.
b) Set the boat electrofishing unit to pulsed DC and mark it in the Wave Form section of the fish
gear form.
c) Test the other settings outside of the sampling area. Start the electrofisher, set the timer, and
depress the switch to begin fishing. Typical settings are: 500-1000 V DC; 8-20 A; 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 the voltage, then the pulse width.
d) Record the final settings that will be used for sampling on the fish gear form.
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7) Begin sampling at the upstream end of the support reach (Transect A). Start on the bank (right or
left as you face downstream) selected at random to begin the physical habitat sampling.
a) For small rivers (< 12.5 m wide), move the boat (or raft) within each subreach to sample both
shorelines as well as the mid-channel areas (similar to what is done for a small wadeable
stream using a backpack electrofishing unit).
b) For rivers > 12.5 m wide, sample along the shoreline only. Move to the opposite bank after
every two subreaches. Within each subreach, proceed downstream in close proximity to the
bank and at a pace equal to or slightly greater than the prevailing current to maximize capture
efficiency. You can "nose in" your boat or raft into shoreline habitat to effectively sample but
do not remain in that habitat for too long.
8) Generally effort (i.e., button time) should be 500 -700 seconds per subreach.
a) At sites with maximum reach length (4000 m) it is likely that the entire length within each
subreach (400 m) will not be fished. Depending upon the habitat complexity, you can vary the
distance actively fished to allocate the available button time throughout the subreach. You do
not have to expend equal time among the subreaches.
b) Avoid the temptation to focus sampling in the richest habitat types.
9) Use a 6mm (1/4 inch) mesh dip net to collect stunned fish. Actively capture stunned fish from the
electric field and immediately place them into the live well. Devote special attention to net small
and benthic fishes as well as fishes that may respond differently to the electric current.
a) Irruptive species: If you encounter a large school of a single species (e.g., shad, certain shiners,
etc.), quickly move through it to ensure you can sample the entire subreach within the allotted
button time.
10) Sampling with Secondary Gear: If shallow habitat exists within a subreach that is inaccessible to
your boat or raft, use secondary collection methods to sample the habitat thoroughly. If you use a
secondary electrofishing gear to sample some portion of a subreach, follow the directions provided
in Table 10.4. As a last option, if the conductivity is too high to use a backpack or barge
electrofishing units, seine those areas you cannot access with a boat, following the directions
presented in Table 10.5.
11) Process fish at the completion of each transect to reduce mortality and track sampling effort.
Release fish in a location that eliminates the likelihood of recapture.
12) Repeat Steps 7 through 10 until you have sampled the minimum reach length determined in Step
4.
13) If you have sampled the required minimum fishing reach length but have not sampled all 10
subreaches, determine the total number of individuals collected.
a) If the total is < 500, sample one or more additional subreaches until at least 500 individuals
have been collected and processed, or you sample all 10 subreaches. Go to Step 14.
b) If you collect > 500 individuals, determine if a single irruptive species comprises > 50% of the
total number of individuals.
i) If irruptive species make up > 50% of the sample, sample one or more additional
subreaches to bring the proportion of the irruptive species below 50%. Go to Step 14.
ii) If not, go to Step 14.
c) If you have sampled all 10 subreaches (i.e., you have reached Transect K), go to Step 14.
14) After sampling all the required subreaches, record the final length of the fish sampling reach (as
tracked by GPS or measured by range finder) in the Fish Sampling Protocol section of the fish gear
form.
a) If you suspend sampling before completing the minimum distance, record the length that was
sampled, and mark the reason for the suspension in the Fish sampling - Not Conducted or
Suspended section of the fish gear form.
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b) It is acceptable to take >1 day to complete the fish sampling.
15) In the Primary Electrofishing Gear section of the fish gear form, record the total button time
expended for boat electrofishing, the total time spent sampling, and the percentage of the total
fish sampling reach sampled by the boat electrofishing unit.
a) In the Secondary Electrofishing Gear section of the fish gear form, record the total button time
expended for secondary electrofishing, the total time spent sampling, and the percentage of
the total fish sampling reach sampled by the secondary electrofishing method
b) If seining was used as an additional collection method, record the total number of hauls, the
average haul length, the total time spent seining, and the proportion of the total fish sampling
reach sampled (recorded in the Fish Sampling Protocol section of the fish gear form) for each
type of seine.
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10.3.2 Secondary Electrofishing
If shallow habitat exists within a subreach that is inaccessible to your boat or raft, it will be
necessary to use secondary collection methods to sample the habitat thoroughly. Table 10.4
presents the procedure for collecting fish from large wadeable areas of subreaches through the
use of secondary electrofishing techniques. The intent of the secondary electrofishing methods
is to provide comparable data to boat electrofishing. Do not use secondary methods to sample
"microhabitats" within a subreach. Do not use wadeable electrofishing if your electrofishing unit
is larger than 5000 V (e,g,. a GPP 5.0). Record information about the gear used in the Secondary
Electrofishing Gear section of the fish gear form (Figure 10.1). At the end of sampling, record the
button time of the secondary gear, the total amount of time spent using the secondary method,
and the proportion of the total fish sampling reach that was sampled with the secondary
method. The total button time for boat and wadeable electrofishing should be ~700 seconds per
subreach.
Table 10.4 Procedure: Secondary Electrofishing Methods for Wadeable Areas (Non-Wadeable Rivers)
1) Use secondary electrofishing as a collection method whenever an area within a subreach is
inaccessible by boat (e.g., a large riffle area). The area must be safe to wade. Do not use wadeable
electrofishing if your electrofishing unit is larger than 5000 V (e.g., GPP 5.0).
2) Mark either Backpack or Bank or Towed Barge in the Secondary Electrofishing Gear section of the
fish gear form.
a) Only one netter should be used. Record this along with the number of anodes, and their
diameter (mark the units as either inches or centimeters) on the fish gear form.
b) For backpack electrofishing there may be from 2 to 3 people involved (depending upon the
crew size). When using a barge or pram, the minimum crew size for electrofishing is three. The
barge operator must remain at the control box and navigate the barge. Use only one probe
operator, but there may be 1-2 additional crewmembers involved (depending upon crew size).
3) Operation of Backpack Electrofisher—
a) Set unit to pulsed DC and mark it in the Wave Form section of the fish gear form.
b) Select the initial voltage setting based on the conductivity of the river.
i) 150-400 V for high conductivity (>300 u.S/cm).
ii) 500-800 V for medium conductivity (100 to 300 u.S/cm).
iii) 900-1100 V for low conductivity (< 100 u.S/cm).
c) Select the initial pulse rate and width.
i) In waters with strong swimming fish (length >200 mm), use a pulse rate of 30 Hz with a
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pulse width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-70 Hz.
d) Turn the electrofisher on, 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.
e) Once you have determined the appropriate settings, record them on the fish gear form. Start
cleared clocks and resume fishing.
i) Note: some electrofishers do not meter all the requested settings; provide what you can.
f) If button time is not metered, estimate it with a stop watch and flag the data.
4) Operation of Tote Barge Electrofisher—
a) Set unit to pulsed DC and mark it in the Wave Form section of the fish gear form.
b) Test settings outside of the sampling area. Start the electrofisher, set the timer, and depress the
switch to begin fishing. Typical settings are:
i) 500-1000 V DC
ii) 8-20 A
iii) 120 Hz.
iv) 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.
5) Once the settings on the electrofisher are adjusted properly to sample effectively and minimize
injury and mortality, begin sampling at the wadeable area at the downstream end and work
upstream.
a) Depress the switch and slowly sweep the electrode from side to side sampling the wadeable
area.
b) In slack water areas, 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.
c) In fast, shallow water, sweep the anode and fish downstream into a net. Keep the cathode near
the anode if fish catch is low.
6) Use a 6mm (1/4 inch) mesh dip net to collect stunned fish. Actively capture stunned fish from the
electric f ie/d and immediately place them into the live well. Devote special attention to net small
and benthic fishes as well as fishes that may respond differently to the electric current.
7) The total button time for boat and wadeable electrofishing should be -700 seconds per subreach.
8) After you finish sampling the wadeable area, return to Step 11 of Table 10.2.
9) You need to track the button time, total fishing time, and the length of stream sampled by wading in
each subreach. At the end of all sampling, record the total button time and total fishing time
expended for wadeable electrofishing in the Secondary Electrofishing Gear section of the fish gear
form. Estimate the proportion of the fish sampling reach (recorded in the Fish Sampling Protocol
section of the fish gear form) that was sampled by wadeable electrofishing.
10.3.3 Secondary Seining
In waters where high conductivity would require the use of an electrofishing unit larger than
5000 V (e.g., a 7.5 or 9.0 GPP system), use seining as a secondary method in large wadeable
areas that are inaccessible to a boat. Use seining only as the last option for collecting fish at a
non-wadeable site. If conditions are such that seining is the only method used, provide a
justification in the Sampling Protocol Comments section of the fish gear form (Figure 10.1).
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Table 10.5 presents the procedure for seining large wadeable areas of subreaches. The intent of
the supplemental seining is to provide comparable data to boat electrofishing. Do not use
seining to sample small "microhabitats" within a subreach.
Record any fish collected with seining protocols on a separate line in the field forms than the
fish electroshocked fish.
Although wadeable electrofishing techniques typically work best in an upstream direction,
seining may work best moving downstream. Depending upon habitat types and complexity, use
2 to 3 crew members. Two crewmembers move the seine; a third person creates and maintains
a bag in the seine in areas with higher velocities, or agitates rocks in riffles or snags. To avoid
mortality, process fish after each seine haul. Use additional lines on the fish collection form
(Figure 10.2) to record species collected by seining (i.e., do not combine results for a single
species from boat electrofishing and seining on the same line). To track effort, seine
characteristics and haul length, habitat, and time should be tallied after each seine haul.
If you seine, record information for each seine haul on the Seining Information Form to track
effort (Figure 10.4). Denote the bank as right or left as you face downstream. Restrict each haul
to a single habitat type. After fish sampling is completed for the site, use the information from
the seining information form to complete the information in the Primary and Secondary Seine
Net sections of the fish gear form (Figure 10.1). Include the seining information form with the
packet of completed field forms you submit to the NRSA information management staff.
Table 10.5 Procedure: Secondary Seining Methods for Wadeable Areas (Non-Wadeable Rivers)
1) Use secondary seining as a last option only (e.g., when electrofishing is ineffective due to high
conductivity or extremely high turbidity). Area must be safely wadeable.
a) If site conditions are such that only seining is used, note the reason in the Sampling Protocol
Comments section.
b) At the end of each seine haul, immediately place all fish in one or more live wells to minimize
injury and mortality, and so that most fish can be returned to the river alive.
2) Complete the header section of the fish gear form (Site ID and date).
3) Mark the pertinent protocol and size class in the Fish Sampling Protocol section.
a) Proceed to the downstream end of the support reach (Transect A).
i) At some sites, seining may be more effective while working downstream (from Transect K)
instead of upstream.
(1) If working downstream in a large wadeable stream, reverse the transects in Figure 10.3
and move to the opposite bank where indicated.
b) For safety, everyone must wear personal floatation devices and foot protection.
c) To aid vision while seining, wear polarized sunglasses and a hat or visor.
4) Mark the appropriate Water Visibility conditions on the form. Poor implies that your ability to seine
effectively is compromised because of poor visibility. Record the water temperature and
conductivity..
5) Mark the type of seine being used (Bag Seine or Minnow Seine) in the Primary Seine Net section of
the fish gear form. This is the seine that will be used for sampling the majority of the fish sampling
reach.
a) Record the number of crewmembers (2-3), and the net dimensions (height, total length, and
mesh size) on the fish gear form.
b) If you have to use a second type of seine for parts of the sampling reach, Mark the type and
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6)
8)
record the dimensions in the Secondary Seine Net section of the fish gear form.
Determine the length of the fish sampling reach and the number of subreaches that should be
sampled (refer to Table 10.2)
7) To maximize capture efficiency, please do the following:
a) Always use 10 and 20 ft. seines. When necessary, reduce the width by rolling seine poles and
floats into the net.
b) When narrowing seines, always keep lead line outside of the pole.
c) When working edge habitats, only roll the inner side of the seine, while keeping the near bank
pole extended.
d) As a default, use seines that are 2 meters in depth. A 1.25 meter seine may be used in shallow
habitats.
e) Keep the float line above the surface (avoid dragging it below the surface while pulling).
f) Maintain the lead line along the river bottom.
g) Either tie the seine to the poles tightly, or roll the seine into the poles.
h) Always maintain the bag behind the poles.
Seining habitats include large riffles or gravel bars, pools (which include backwater areas), glides or
runs, edges, and snags. Seine width and haul length is dependent upon the water velocity, depth,
and/or complexity of the habitat.
a) The objective of the seining effort is to acquire a comparable collection of fish (in terms of
species richness and relative abundance, and allocation of effort throughout the fish sampling
reach) to that obtained if the site was electrofished.
i) Avoid extended seine hauls that collect hundreds of individuals.
ii) Seine as many available habitat types as possible within each subreach (one haul each).
iii) Total time spent seining a site should be comparable to what would have been spent
electrofishing.
Riffle Habitats
b)
Use two crewmembers, each tending a seine pole. Place the seine perpendicular to the
current across the downstream end of the riffle. Ensure that the lead line is on the
bottom. Tilt the net slightly downstream to form a bag to trap aquatic vertebrates.
Starting no more than 3 m upstream, a third crewmember kicks the substrate and
overturns rocks, proceeding quickly downstream toward the net.
When the area is thoroughly kicked, quickly raise and bag the net. Process fish (i.e.,
enumerate, identify and voucher fish) and record tally information on the fish collection
form (Figure 10.2). You may use separate lines on the fish collection form to record
species information from seine hauls.
Record seine characteristics and haul length, habitat, and time on the seining
information form
c) Pool, Backwater, and Bar Habitats (Slack water)
i) Use two crewmembers, each tending a seine pole. Pull the seine across the pool using
shallow riffles or banks as barriers. A third crewmember creates and maintains a bag in
the seine.
ii) In areas with current, pull the net downstream and then sweep toward the bank with
one or both poles, or post one pole on the bank and sweep the other end in a wide arc
from midstream to the same bank.
iii) You can work pools in short to long hauls and use seines of varying width depending on
the complexity and depth of the pool. Keep the seine depth constant at 2 meters.
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iv) Pull the bag completely to shore at a predesignated point.
d) Glide or Run Habitats (noticeable current)
i) Use two crewmembers, each tending a seine pole. Pull the seine diagonally across the
glide towards the bank. If necessary, a third crewmember creates and maintains a bag in
the seine.
ii) Pull the net quickly downstream along the glide moving diagonally toward the bank.
When you reach the bank with the outer edge of the seine, post the pole and sweep the
other end in a wide arc from midstream to the same bank.
iii) Because of decreased complexity and shallower depths, seine hauls in glides or runs are
typically longer and use wider nets. You can use a 1.25 m deep seine in shallow glides.
iv) Pull the bag completely to shore at a predesignated point.
i
e) Edge Habitats
Edge habitats may be shallow too deep with complex to uniform habitat, and may
include undercut banks.
Use two crewmembers, each tending a seine pole. Seine along the near shore area.
The near bank crewmember moves along the shore while jabbing along any undercut or
small structure. The other crewmember stays ahead of the shoreline pole to maintain a
"J" in the seine bag. At a predesignated point, post the near shore pole and sweep the
seine towards and up on the bank.
Depending on edge complexity and depth, seine width and haul length may vary. Use
wider seines and longer hauls in shallower, less complex habitats. As complexity, depths,
and flow increase, shorten the seine width and haul length accordingly. Seine depth may
vary depending on depth.
iv
f) Snag Habitats
i) Snag habitats often require creativity in terms of seine length and approach. You can use
a 1.25 meter deep seine to avoid snagging the net on structure, but use a 2 m deep seine
in deeper areas. Narrow seine widths and short hauls are preferred.
ii) Use two crewmembers, each tending a seine pole. Jab seining is often the most effective
method. Quickly jab a shortened seine (< 2 m wide) under the cover and near the river
bottom, and then quickly lift the seine to the water surface. You can use a third
crewmember to agitate the snag to move fish out toward the seine.
iii) For small snags along the bank, seining along the edge may work best. The near snag
crewmember moves along the snag, while jabbing along its length. The other
crewmember stays ahead of the shoreline pole to maintain a "J" in the seine bag. At a
predesignated point, quickly pull the seine to the surface.
9) To minimize mortality, process fish (i.e., identify, count, and prepare preserved voucher specimens or
photovoucher images) after each seine haul (rather than at the end of a subreach.
a) Record identifications, tallies, and voucher information on the fish collection form. You may use
separate lines on the fish collection form to record species information from separate seine
hauls.
b) Tally seine characteristics and haul length, habitat, and time on the seining information form.
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10) After you finish seining the wadeable area of the subreach, return to Step 11 of
16) Table 10.3 Procedure: Electrofishing (Non-Wadeable SitesjComplete the header section of the
fish gear form (Site ID and date).
11) At the end of all sampling, use the seining information form to determine the total number of hauls,
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the average haul length, the total time spent seining, and the proportion of the total fish sampling
reach sampled (recorded in the Fish Sampling Protocol section of the fish gear form) for each type of
seine. Record the totals in the Primary and Secondary Seine Wet section of the fish gear form.
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10.4 Processing Fish
Processing the fish at the end of each subreach is described in Table 10.6. However, if fish show
signs of stress (e.g., loss of righting response, gaping, gulping air, excessive mucus) in the middle
of a subreach, change the water in the live well or stop fishing and initiate processing. Always
process and release individuals of State or Federally listed threatened or endangered species or
large game fish immediately after collection. After processing fish, release them in a location
that prevents the likelihood of their recapture.
10.4.1 Identification and Tallying
Record species identifications, tallies, and other information for individuals collected on the Fish
Collection Form (Figure 10.2). Use multiple pages of the form as needed to record all species
collected. It is important to record page numbers correctly because page number is one of the
variables used to uniquely identify a species record. You can record separate collections of the
same species on multiple lines of the collection form (e.g., when you encounter a species in non-
adjacent subreaches, or collect a species with a secondary gear type). Do not process individuals
with total length < 25 mm (1 inch), as these are likely young of year individuals that cannot be
identified confidently to species. Only crew members designated as "taxonomic specialists" by
EPA regional coordinators can identify fish species. Tally fish by species and major size class (6
inch [15 cm] intervals), and examine them for the presence of DELT (Deformities, Eroded Fins,
Lesions and Tumors) anomalies. Use common names of species established by the American
Fisheries Society Common and Scientific Names of Fishes from the United States, Canada and
Mexico (Nelson, et al. 2004). Appendix D provides a list of species names reported from the
NRSA 2008/09.
If you believe a specimen is nonindigenous to the site, mark it as Introduced on the collection
form. If you suspect it represents a potential range extension for the species, prepare one or
more specimens (preserved if possible but photographs if not). Physical specimens are required
in order to publish reports of range extensions. Include specimens to document suspected range
extensions are included as part of the preserved Unknown/Range Extension voucher sample
(UNK/RNG; Section 10.4.2).
10.4.2 Unknown Specimens
If you cannot positively identify individuals to species in the field, record taxonomic information
of the collection form using scientific names rather than common names. If you can identify a
specimen only to family, record the scientific rather than the common family name (e.g.,
UNKNOWN PERCID A, not UNKNOWN PERCH A) on the fish collection form. If you can identify a
specimen to genus, record the scientific name rather than the common name (e.g., UNKNOWN
PERCINA A, not UNKNOWN DARTER) on the fish collection form. Using scientific rather than
common names for unknowns reduces ambiguity, since some common names may in fact refer
to multiple genera (e.g., "darter", "shiner", "sucker", "sunfish", etc.). If you identify an unknown
species to Genus, retain a small number (up to 20 individuals per putative species) as part of the
preserved UNK/RNG voucher sample (see SectionlO.4.6) or take good digital photographs LU
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(Section 10.4.3) for laboratory identification. If you are only able to identify an unknown to <
Family, retain as many of the individuals as possible for later identification. Use the UNK/RNG £°
Voucher label on the label sheet to label your jar of unknown to track from which sites the LU
unknowns originated. ^
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Table 10.6 Procedure: Processing Fish (Non-Wadeable Sites)
Fish Processing
1) Complete all header information accurately and completely on the fish collection form. It is
important to paginate the collection forms correctly (e.g., start with page 1, do not duplicate page
numbers, etc.), as page number is part of the unique record identifier for the fish count data.
2) Process individuals collected at the end of each subreach. You can record a single species on
multiple lines of the collection form (e.g., use separate lines for individuals collected in nonadjacent
subreaches, collect with a secondary gear type, or if you need additional space to record tally marks,
etc.).
a) Process species listed as threatened and endangered first as described in Step 4.
i) Photograph specimens for voucher purposes if conditions permit and stress to individuals
will be minimal. Mark as Photo in the Voucher section of the collection form.
ii) If individuals die due to sampling, prepare them as part of the local voucher sample and
preserve them in the field. Comply with the conditions of your collection permit in regards
to mortality of listed species).
iii) Return individuals to the river immediately after processing.
3) Only identify and process individuals > 25mm (1 inch) in total length (TL). Ideally handle specimens
only once. Although not required, you may note amphibians and reptiles captured on the fish
collection form.
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4) Identify each individual to the lowest possible taxonomic level:
a) If you can confidently identify the individual to species, record the common name on the first
blank line in the Common Name field of the fish collection form.
i) Common names should follow those recognized by the American Fisheries Society. Use of
alternative names is discouraged. Use names presented in Appendix D, which are based on
those used in the NRSA 2008/09.
(1) Record the complete common name. Avoid using shortened names (e.g., stoneroller,
carp, bass, etc.).
(2) If you use a non-standard name, you must assign a flag to the line and provide the
taxonomic reference for the name in the Comments section of the collection form.
b) If you cannot positively identify an individual to the species level:
i) Identify it to the lowest taxonomic level (i.e., family or genus). Record the putative name as
UNKNOWN plus the scientific name of the family or genus (e.g., UNKNOWN CATOSTOMID
A, UNKNOWN MOXOSTOMA A) in the Common Name column of the collection form.
ii) If you are permitted to retain the specimen, assign it the next available sequential tag
number (starting with 01) in the Voucher Tag Number column and see Step 9.
c) If you believe the individual is a hybrid:
i) Mark as Hybrid? on the collection form.
ii) If the hybrid has an accepted standard common name (e.g., Tiger muskellunge, Saugeye,
Wiper, etc), record that name. For other hybrids record the common name of both species
(e.g., Green sunfish x Bluegill, Cutthroat trout x Rainbow trout). Avoid using non-specific
terms such as Hybrid sunfish.
iii) If you are unsure of the identification and are permitted to retain the specimen, assign it
the next available sequential tag number (starting with 01) in the Voucher Tag Number
column and see Step 9.
5) If you know the species is not native to this location, mark as Introduced?
6) Visually estimate the total length of each individual (a measuring board is not necessary). Keep a
running tally in the appropriate Tally and Counts section (< 6 in., 6-12 in., 12-18 in., or > 18 in.) of the
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fish collection form.
a) If all individuals of a species appear to be the same size, provide a flag and comment for the line
if you believe the population is stunted.
7) Examine each individual for external anomalies. 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 you process all of the individuals of a species, record
the total number observed in iheAnom Count column of the collection form.
8) If an individual has died due to electrofishing or handling, include it in the running tally for the
species. After you process all of the individuals of a species, record the total number observed in the
Mortality/Count column of the collection form.
9) If you are retaining individuals of the species as part of the preserved Unknown/Range Extension
(Unk/Rng) voucher sample:
a) Mark as Unk/Rng in the Voucher section of the collection form.
b) Assign the species the next available voucher specimen tag, and record the number in the
Voucher Tag # column of the collection form.
i) If you take one or more photographs of the species instead of preserving specimens, assign
the next available voucher specimen tag number in the Voucher Tag # column of the
collection form. Include the specimen tag in all photos of the species. Mark Photo in the
Voucher section of the collection form.
(1) Ideally, take photos of all species collected at a site that are not being preserved.
c) Record the number of individuals retained for the preserved voucher sample in the Vouchers
Retained column of the collection form.
i) NOTE: Do not keep separate tallies of voucher and non-voucher specimens. Record all
individuals in the appropriate area of the Tally and Counts section. The retained voucher
specimens represent a subsample of the total count.
ii) Place the specimens in a jar which has been labeled with the site ID. You can have multiple
individuals of the same species in the jar, but each species will have a separate voucher tag
number (i.e. one tag number per line on the collection form).
10) If you are retaining specimens as part of a preserved QA voucher sample for the site:
a) Mark as QA in the Voucher section of the fish collection form.
i) NOTE: This should be marked at least once for all species collected at the site (including
unknowns).
b) Use the sheet of labels and tags for the QA voucher sample (the jar label has a preprinted
sample ID number). Assign the species the next available voucher specimen tag number. Record
the specimen tag number in the Voucher Tag # column of the collection form.
i) If you take one or more photographs of the species instead of preserving specimens, assign
the next available voucher specimen tag number, and record the number in the Voucher
Tag# column of the collection form. Include the specimen tag in all photos of the species.
Mark Photo in the Voucher section of the collection form.
c) Record the number of individuals retained for the preserved voucher sample in the Vouchers
Retained column of the collection form.
i) NOTE: Do not keep separate tallies of voucher and non-voucher specimens. Record all
individuals in the appropriate area of the Tally and Counts section. The retained voucher
specimens represent a subsample of the total count.
ii) Place the specimens in a fine mesh bag (or separate jar) along with the voucher specimen
tag that matches the number recorded on the collection form. You can have multiple bags
(or jars) of the same species, but each bag (or jar) will have a separate voucher tag number
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(i.e., one tag per line on the collection form).
11) Repeat Steps 2 through 10 for each subreach sampled. Use additional fish collection form sheets as
needed, being careful to paginate each sheet correctly.
12) If you collect fish via seining, record he information on a separate line on the field form.
13) At the end of sampling, follow the appropriate procedure to prepare the preserved voucher samples
(UNK/RNG and/or QA) and/or select specimens for tissue samples.
a) For all voucher samples, use a sufficient volume of 10% buffered formalin—the volume of
formalin solution used must exceed the volume of specimens. Use additional jars if necessary.
Slit large individuals (TL > 200 mm [~8 in.]) along the right side in the lower abdominal cavity to
allow penetration of the formalin solution.
14) Complete a sample jar label for the UNK/RNG voucher sample. Attach it to the sample jar and cover
it with clear tape.
15) If you did not prepare a QA voucher sample, mark Wo Voucher Preserved on the back of the fish gear
form.
a) Otherwise complete a sample jar label for the QA voucher sample. Attach it to the sample jar
and cover it with clear tape.
b) Record QA voucher sample label information on the back of the fish gear form.
16) Record the file names of any photovouchers taken on the back of the fish gear form.
a) Use only one line per voucher tag, even if you took multiple photos (record the beginning and
end of the sequence in the Sequence column). Make sure the page and line numbers you record
match those on the collection form.
b) Name image files as: Site ID + Visit number + tag number + sequence (e.g.,
CTLS1001_Vl_tag01a).
17) If you did not collect any fish from the entire fish sampling reach, mark Fished - None Collected in
the Fish Sampling - Not Conducted or Suspended section of the fish gear form.
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10.4.3 Photovouchering
Use digital imagery for fish species that cannot be preserved as voucher specimens (e.g., rare,
threatened, and endangered species; very large bodied). Ideally, take photos of all species
collected a site (that are not preserved) to provide a minimal level of documentation of
occurrence. Take photographs of entire specimens and additional specific morphological
features that are appropriate and necessary for an independent taxonomist to accurately
identify the specimen. Additional detail for these guidelines is provided in Stauffer et al. (2001),
which 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 (or morphological features) should occupy as much of the field of view as
possible. Use a fish measuring board, ruler, or some other calibrated device to provide a
reference to scale. Provide an adequate background color for photographs (e.g., fish measuring
board). Include a card with site ID number, site name, and date in each photograph so that
photos can be identified if file names become corrupted. In addition, include the voucher
specimen tag that you assign to the species to provide a link to the line on the fish collection
form. For each photovoucher specimen, include at least a full body photo (preferably of the left
side of the fish), and other macro images of important morphological features (e.g., lateral line,
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ocular/oral orientation, fin rays, gill arches, mouth structures, etc.). It may also be necessary to
photograph males, females, or juveniles.
Save images in medium to high quality jpeg format. It is important that time and date stamps
are accurate, as this information can also be useful in tracking the origin of photographs.
Transfer images stored in the camera to a PC or external storage device (e.g., thumb drive or
flash memory card) at the first available opportunity. At this time, rename the original files to
include the site ID, visit number, voucher specimen tag number, and photo sequence (e.g.,
CTLS1001_Vl_tag01a.jpg). Record the file names on the back of the fish gear form (Figure 10.6).
Maintain a complete set of your photovoucher files in a safe location (e.g., an office computer
that is backed up regularly) for the duration of the sampling season. At this time, you will
transfer all images to the NRSA IM staff. You will also send copies of your image files as part of
your QA voucher samples.
10.4.4 Preparing Preserved Voucher Specimen Samples
There are two different types of samples for preserved voucher specimens. The UNK/RNG
voucher samples are used to identify specimens that cannot be confidently identified in the
field, and to provide physical specimens of suspected range extensions. After submitting the fish
collection form to the NRSA IM staff, you will receive an update form that lists only the records
for unknown species recorded on the fish collection form (including photovouchers) that were
marked as being part of the UNK/RNG voucher sample.
In addition to a UNK/RNG voucher sample (if needed), you will prepare an additional QA voucher
sample (Section 10.4.7). A QA voucher sample will be performed at a pre-designated set of sites
and includes preserved specimens (or photographs) of all species collected at a site (including
the unknowns). Use the voucher specimen tags and sample labels designated for QA voucher
samples. QA voucher samples are eventually sent to an independent taxonomist as a check on
the accuracy of each fish taxonomist.
10.4.5 Preserving Voucher Specimen Samples
Preserve UNK/RNG and QA voucher specimens in the field with a 10% buffered formalin
solution. The volume of formalin must be equal to or greater than the total volume of
specimens. Use additional jars if necessary to ensure proper preservation. For individuals having
a total length larger than 200 mm (~8 in.), make a slit along the right side of the fish in the lower
abdominal cavity to allow penetration of the preservative solution. Follow all the precautions for
handling formalin outlined in the MSDS. Formalin is a potential carcinogen. Handle 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.
Once you have completed preserving all jars of voucher specimens, complete the appropriate
jar label (Figure 10.5 for UNK/RNG samples, and Figure 10.7 for QA voucher samples). Attach
the completed label to the jar with clear shipping tape. If you have > 2 jars of either type of
sample, prepare a hand-made label for each additional jar (this is more important for the QA
voucher sample, which has a unique sample ID number).
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FISH VOUCHER - UNK/RNG
Site ID:
_/ l2Q-\_
FISH - BAG
TAG: 41
FISH - BAG
TAG: 37
FISH - BAG
TAG: 33
FISH - BAG
TAG: 29
FISH - BAG
TAG: 25
FISH - BAG
TAG: 21
FISH - BAG
TAG: 17
FISH - BAG
TAG: 13
FISH - BAG
TAG: 09
FISH - BAG
TAG: 05
FISH - BAG
TAG: 01
FISH - BAG
TAG: 42
FISH - BAG
TAG: 38
FISH -BAG
TAG: 34
FISH - BAG
TAG: 30
FISH - BAG
TAG: 26
,0
FISH - 5AG
TAG. 22
FISH - BAG
TAG: 18
se
FISH -BAG
TAG: 14
FISH - BAG
TAG: 10
FISH -BAG
TAG: 06
FISH - BAG
TAG: 02
FISH VOUCHER - UNK/RNG
Site ID:
_l /201_
FISH - BAG
TAG: 43
FISH - BAG
TAG: 39
FISH- BAG x\v
TAG: 35
FISH t AG
TAo. 31
\3~FISH - BAG
TAG: 27
FISH - BAG
TAG: 23
FISH - BAG
TAG: 19
FISH -BAG
TAG: 15
FISH - BAG
TAG: 11
FISH - BAG
TAG: 07
FISH -BAG
TAG: 03
FISH - BAG
TAG: 44
FISH -BAG
TAG: 40
FISH - BAG
TAG: 36
FISH - BAG
TAG: 32
FISH -BAG
TAG: 28
RSH - BAG
TAG: 24
FISH -BAG
TAG: 20
RSH - BAG
TAG: 16
FISH -BAG
TAG: 12
FISH -BAG
TAG: 08
FISH -BAG
TAG: 04
Figure 10.5 Unknown/Range Extension Voucher Sample Labels and Voucher Specimen Tags
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Site ID:
NRSA 2013/14 FISH GEAR AND SAMPLING INFORMATION (Back)
Date: /
Reviewed by Hnnul]:.
QA VOUCHER SAMPLE INFORMATION (VERT)
NO VOUCHERS PRESERVED Q
Sample ID
it of Jars Preserved
O
PHOTO VOUCHER FILE INFORMATION
Page
Photo Fils Name (SltelD_V(Vlsll#LTagS)
(e.g.: SSCT001_V1_Tag01)
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10.4.6 Processing Unknown/Range Extension (UNK/RNG) Voucher Samples
Table 10.7 outlines the procedure for determining the identification of unknown specimens
from each UNK/RNG sample. A more detailed procedure for conducting the laboratory
identifications is presented in the NRSA laboratory operations manual (USEPA 2013b). Identify
unknown using whatever resources are necessary (magnification, literature, reference
collections/specimens, including dissected anatomical features or in house colleagues).
Following positive laboratory identification, use the update form for the sample (Figure 10.8) to
reconcile the unknown records to reflect the actual species identifications and numbers. It is
important to update counts and identifications by voucher tag do not combine multiple
samples of the same unknown before updating.
If all specimens for an unknown record are a single species, simply record the final identification
(as common name from the standard list [Appendix D]) in the Revised Name column, and enter
the original count in the Revised Count column. If you determine that a single unknown record is
actually >1 species, you will record the new species on blank lines. For example, if a sample of
20 specimens of UNKNOWN COITUS A is later identified as 15 individuals of one species and 5
individuals of another correct the total count for the unknown assigning 75% of the original total
count to the first species and 25% of the original total count to the second species. Record the
information (revised common name and revised count) for the first species on the same line as
the original unknown record. Record the information for additional species from this original
unknown as new data records, but retain the page, line number, and voucher specimen tag
number of the original unknown record. Record the name and count in the Revised Name and
Revised Count columns.
If you use a non-standard name, enter the page, line number, tag number, and taxonomic
reference for the name in the Comments section of the update form. Submit your completed
update forms to the NRSA IM staff as soon as possible after completing the laboratory
identifications. Retain the preserved UNK/RE voucher samples from each site- contact your
regional EPA coordinator if you cannot store the samples at your facility.
If your attempts at identification do not yield a positive identification for 100% of the fish you
retained, contact the Field Logistics Coordinator for further guidance (Chris Turner,
cturner@glec.com. 715-829-3737). There are provisions under which fish can be identified by a
contracted lab and the results returned to you.
10.4.7 Processing QA Voucher Samples
Prepare the QA voucher sample as outlined in Table 10.8. Prepare the QA voucher sample
separately from the UNK/RNG voucher sample. Processing involves ensuring that the sample
jar(s) and photovoucher files include specimens of ALL species (including unknowns and
common species) collected from the site. Each unique species (including unknowns) should have
a unique QA voucher specimen tag assigned (Figure 10.7). Record information about the
preserved QA voucher sample on the back side of the fish gear form (Figure 10.6).
S Retain all of your QA voucher samples (including digital image files) until given direction by EPA
-i regarding where to send them. When you are ready to ship the samples, complete a sample
^ tracking form as described in Appendix C. QA voucher samples may require shipping as
I i i
$ "dangerous goods," and packing and documentation requirements will differ depending on
** whether the samples contain formalin or ethanol, and on the particular shipping service used.
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QA FISH VOUCHER (VERT)
Site ID:
_/ /201_
990011
FISH - BAG
TAG:
41
990011
FISH - BAG
TAG:
37
990011
FISH - BAG
TAG:
33
990011
FISH - BAG
TAG:
29
990011
FISH - BAG
TAG:
25
990011
FISH - BAG
TAG:
21
990011
FISH - BAG
TAG:
17
990011
FISH - BAG
*~fto:
13
990011
FISH - BAG
TAG:
09
990011
FISH - BAG
TAG:
05
990011
FISH - BAG
TAG:
01
990011
FISH - BAG
TAG:
42
990011
FISH - BAG
TAG:
38
990011
FISH - BAG
TAG:
34
990011
FISH - BAG
TAG:
30
990011
FISH -BAG
TAG:
26
990011
FISH - BAC
OTAG:
22
9s>ini
FISH - BAG
TAG:
18
990011
FISH - BAG
TAG:
14
990011
FISH - BAG
TAG:
10
990011
FISH -BAG
TAG:
06
990011
FISH - BAG
TAG:
02
990011
QA FISH VOUCHER (VERT)
Site ID:
_/ /201_
990011
FISH - BAG
TAG:
43
99001 1
FISH - BAG
TAG:
39
990011
FISH - BAG
T,AG:
?5
990011
FISH - BAU
TAG:
31
900011
FISH - BAG
TAG:
27
990011
FISH - BAG
TAG:
23
99001 1
FISH - BAG
TAG:
19
990011
FISH - BAG
TAG:
15
990011
FISH - BAG
TAG:
11
990011
FISH -BAG
TAG:
07
990011
FISH - BAG
TAG:
03
990011
FISH -BAG
TAG:
44
990011
FISH -BAG
TAG:
40
990011
FISH -BAG
TAG;
36
990011
FISH - BAG
TAG!
32
990011
FISH - BAG
TAG:
23
990011
FISH -BAG
TAG:
24
990011
FISH -BAG
TAG:
20
990011
FISH - BAG
TAG:
16
990011
FISH -BAG
TAG:
12
990011
FISH -BAG
TAG:
08
990011
FISH -BAG
TAG:
04
990011
Figure 10.7 QA Voucher Sample Labels and Voucher Specimen Tags
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Table 10.7 Procedure: Processing Unknown/Range Extension (UNK/RNG) Voucher Samples
Processing UNK/RNG Voucher Samples
1) Following fixation for 5 to 7 days, decant and properly discard the formalin solution. 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.
a) Formalin must be disposed of properly. Contact your regional EPA coordinator if your laboratory
does not have the capability of handling waste formalin.
2) Replace the formalin with tap water and soak specimens over a 4-5 day period. Soaking may require
periodic water changes and should continue until the odor of formalin is barely detectable.
3) Decant the tap water. Use 45%-50% isopropyl alcohol or 70% ethanol as a final preservative for
specimens.
4) You will receive a Fish Update Identification and Count Form for each UNK/RNG sample from the
NARS IM staff. This form lists all records from the original collection form that were marked as being
part of the UNK/RNG voucher sample. Identify unknown fish to species in the laboratory, using the
procedure described in the NRSA laboratory operations manual, which is briefly described below.
a) Process unknowns by tag number do not combine multiple bags (or jars) of the same unknown
before determining the final identifications. Corrections and updates need to be linked back to
the original page and line number, and voucher specimen tag you recorded on the collection
form.
5) Record the final identification(s) and count for each unknown record in the Revised Name and
Revised Count columns of the update form.
a) Use common names from the standard list (Appendix D) as revised names.
b) If you must use a non-standard name, provide the page, line number, specimen tag number, and
the taxonomic reference in the Comments section of the update form.
6) If an unknown turns out to include > 1 species, correct the final counts based on the proportion of
each species found in the original unknown bag.
a) Record the revised name and count for one species on the line of the original unknown.
b) Record the revised name and count for the second species as a new record on the update form.
i) Record the page, line number, and specimen tag number from the original unknown record
on the next available blank line of the update form.
ii) Mark New Record. Leave the Original Name and Original Count Columns blank.
iii) Record the revised name and count in the Revised Name and Revised Count columns.
7) After reconciling all of the unknowns, submit the completed update forms to the NARS IM staff in
Corvallis. Retain the preserved UNK/RNG voucher samples. Contact your regional EPA coordinator if
you cannot store the samples at your facility.
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Table 10.8 Procedure: Processing QA Voucher Samples
1) Ensure that all species collected at a site are represented by either preserved voucher specimens or
photovouchers. There should be a unique QA voucher specimen tag assigned to every species
recorded on the fish collection form.
2) Before submitting the QA Voucher sample, ensure that all specimens have been positively identified.
If your attempts at identification do not yield a positive identification for 100% of the fish you
retained, contact the Field Logistics Coordinator for further guidance (Chris Turner,
cturner@glec.com, 715-829-3737).
3) After preparing the preserved QA voucher sample, check that the sample ID number recorded on the
fish gear form matches the preprinted label attached to each sample jar, and that the number of jars
recorded on the fish gear form is correct.
4) Retain the QA voucher samples in appropriate storage space for formalin until you receive
information regarding where to send them from the NRSA Project Manager or EPA Regional
Coordinator..
5) If you are storing the preserved QA voucher samples for an extended period, you may need to
replace the formalin fixative with ethanol.
a) Following fixation for 5 to 7 days, decant and properly discard the formalin solution. Formalin is
a potential carcinogen —handle 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.
b) Formalin must be disposed of properly. Contact your regional EPA coordinator if your laboratory
does not have the capability of handling waste formalin.
6) Replace the formalin with tap water and soak specimens over a 4-5 day period. Soaking may require
periodic water changes and should continue until the odor of formalin is barely detectable.
7) Decant the tap water. Use 45%-50% isopropyl alcohol or 70% ethanol as a final preservative for
specimens.
8) When ready to ship all of the QA voucher samples, complete a sample tracking form as described in
Section 3 and Appendix C.
9) Package the preserved samples properly for either formalin or ethanol and prepare all required
documentation and safety measures for the shipment.
10) Include a CD with the photovoucher files for each QA voucher sample in the shipment. Use the file
names that are recorded on the fish gear form.
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11 FISH TISSUE PLUG SAMPLING METHOD
11.1 Method Summary
Because many fish spend their entire life in a particular water body they can be important
indicators of water quality, especially for toxic pollutants (e.g., pesticides and trace elements).
Toxic pollutants, which may be present in the water column or sediments at concentrations
below our analytical detection limits, can be found in fish tissue due to bioaccumulation.
Typical fish tissue collection methods require the fish to be sacrificed, whether it be a whole fish
or a skin-on fillet tissue sample. This can be problematic when there is a need to collect large
trophy-sized fish for contaminant analysis or when a large sample size is necessary for statistical
analysis. The following describes an alternative method for the collection of fish tissue samples
which uses a tissue plug instead of a skin-on fillet. Two fish tissue plugs for mercury analysis will
be collected from two fish of the same species (one plug per fish) from the target list (below) at
every site. These fish are collected during the fish assemblage sample collection effort (Section
0). A plug tissue sample is collected by inserting a biopsy punch into a de-scaled thicker area of
dorsal muscle section of a live fish. After collection, antibiotic salve is placed over the wound
and the fish is released.
11.2 Equipment and Supplies
Table 11.1 lists the equipment and supplies necessary for field crews to collect fish tissue plug
samples. This list is comparable to the checklist presented in Appendix A, which provides
information to ensure that field crews bring all of the required equipment to the site. Record
the fish tissue plug sampling data on the Fish Gear and Voucher/Tissue Sample Information
Form (Figure 10.6).
Table 11.1 Equipment and Supplies: Fish Tissue Plug Sample
For fish tissue plug samples
Fish measuring board
Fish weigh scale
Plastic bags
Sterile 20 mL glass scintillation vial
Coolers with ice
Cooler with dry ice
Nitrile gloves
8 millimeter disposable biopsy punch (Acuderm brand Acu-Punch or
equivalent)
Sterile disposable scalpel
Sterile forceps
Laboratory pipette bulb.
Antibiotic salve.
Fish collection gear (electrofisher, nets, livewell, etc.)
Dip net
Field Operations Manual and laminated Quick Reference Guide
For recording measurements
Fish tissue plug sample labels
Fish Gear and Sampling Form
Soft (#2) lead pencils for recording data on field forms
Fine-tipped indelible markers for filling out sample labels
Clear tape strips for covering labels
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11.3 Sample Collection Procedures
The fish tissue plug indicator samples will be collected using the same gear and procedures used
to collect the fish assemblage. Collection of individual specimens for fish tissue occurs in the
sample reach during the fish assemblage sampling. Samples should be taken from the species
listed in the target list found in Table 11.2. 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) to obtain a sample from the
species that are available. Recommended and alternate target species are given in Table 11.2.
The procedures for collecting and processing fish plug samples are presented in Table 11.3.
Table 11.2 Recommended Target and Alternate Species for Fish Tissue Plug Collection
Family name Common name
Scientific name
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Cyprinidae
Centrarchidae
Northern pikeminnow
Bluegill
Rock bass
Redbreast sunfish
Length Guideline
(Estimated
Minimum)
Centrarchidae
Ictaluridae
Percidae
Moronidae
Esocidae
Spotted bass
Largemouth bass
Smallmouth bass
Black crappie
White crappie
Channel catfish
Blue catfish
Flathead catfish
Sauger
Walleye
Yellow perch
White bass
Northern pike
Chain pickerel
Brown trout
Cutthroat trout
Rainbow trout
Brook trout
Micropterus punctulatus
Micropterus salmoides
Micropterus dolomieu
Pomoxis nigromaculatus
Pomoxis annularis
Ictalurus punctatus
Ictalurus furcatus
Pylodictis olivaris
Sander canadensis
Sander vitreus
Perca flavescens
Moron e chrysops
Esox lucius
Esox niger
Salmo trutta
Oncorhynchus clarkii
Oncorhynchus mykiss
Salvelinusfontinalis
~280 mm
~280 mm
~300 mm
~330 mm
~330 mm
~300 mm
~300 mm
~300mm
~380 mm
~380 mm
~330 mm
~330 mm
~430 mm
~430 mm
~300 mm
~300 mm
~300 mm
~330 mm
Ptychocheilus oregonensis ~300 mm
Lepomis macrochirus ~200 mm
Ambloplites rupestris ~200 mm
Lepomis auritus ~200 mm
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Table 11.3 Procedure: Fish Tissue Plug Samples
Fish Tissue Plug Methods
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 two individuals of the same target species from each site. The fish should be of
adequate size to sample (refer to Table 11.2 for minimum species length guidelines). Select
fish based on the following criteria:
• is on the target list,
• both the same species
• both 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, and
• are of similar size, so that the smaller individual is no less than 75% of the total
length of the larger individual.
4. Remove one fish retained for analysis from the clean holding container(s) (e.g., livewell)
using clean nitrile gloves.
5. Measure the fish to determine total body length. Measure total length of the 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).
6. Weigh the fish in grams using the fish weigh scale.
7. Note any anomalies (e.g., lesions, cuts, sores, tumors, fin erosion) observed on the fish.
8. Record site ID, date, sample ID, species, and specimen length and weight on the back of the
Fish Gear and Sampling Form in the Fish Tissue Plug section (Figure 10.6). Make sure the
sample ID numbers and specimen numbers/lengths that are recorded on the collection form
match those on the sample tracking form and labels where applicable.
9. repare a Sample Identification Label for the sample, ensuring that the label information
matches the information recorded on the Fish Tissue Plug section of the Fish Gear and
Sampling Form. Affix label to a sterile 20 milliliter scintillation vial and cover with clear tape.
10. On a meaty portion of the left side dorsal area of the fish between the dorsal fin and the
lateral line, clear a small area of scales with a sterile disposable scalpel
11. Wearing clean nitrile gloves, insert the 8 millimeter biopsy punch into the dorsal muscle of
the fish through the scale-free area. The punch is inserted with a slight twisting motion
cutting the skin and muscle tissue. Once full depth of the punch is achieved a slight bending
or tilting of the punch is needed to break off the end of the sample. Remove biopsy punch
taking care to ensure sample remains in the punch. Note: The full depth of the punch
should be filled with muscle tissue, which should result in collecting a minimum of 0.25 to
0.35 grams of fish tissue for mercury analysis.
12. Apply a generous amount of antibiotic salve to the plug area and gently return the fish to the
water.
13. Using a laboratory pipette bulb placed on the end of the biopsy punch, give a quick squeeze,
blowing the tissue sample into a sterile 20 milliliter scintillation vial.
14. Repeat steps 2-13 for the second fish, collecting a second fish plug sample. Place the second
plug in the same scintillation vial as the first. The two plugs should provide at least 0.5 grams
of tissue.
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Fish Tissue Plug Methods
15. .Place the sample immediately on dry ice for shipment.
16. Dispose of gloves, scalpel and biopsy punch.
17. Keep the samples frozen on dry ice or in a freezer at <-20°C until shipment
18. Frozen samples will subsequently be packed on dry ice and shipped to the batched sample
laboratory via priority overnight delivery service within 1 week.
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12 WHOLE FISH SAMPLING METHOD
12.1 Method Summary
Fish are important integrators of toxic contaminants that are bioavailable in the water column and in
sediment. EPA monitors the occurrence of toxic chemicals in fish fillet samples to assess the potential
health impacts to people who consume fish. Results from the 2008/09 NRSA provided the first
statistically representative national data for fish contamination in U.S. rivers. Collecting whole fish tissue
samples and submitting them to the laboratory for filleting and homogenization during the 2013/14
NRSA allows consistency with fish tissue methods of the previous NRSA, and provides tissue amounts
that are sufficient to allow the analysis of multiple chemical contaminants of concern. It will also provide
the first opportunity for temporal analysis of probability-based national fish contamination trends in U.S.
rivers. Additionally, collecting fish at locations sampled during the previous NRSA will reduce the
variability in data for trends analysis.
Whole fish tissue sampling procedures are described in detail in Table 2.3. The objective is to collect one
whole fish tissue sample from each of the 453 designated target sites. The focus is on fish species that
commonly occur throughout the region of interest and that are sufficiently abundant within a sampling
reach. Each whole fish tissue sample will consist of five adult fish of the same species that are similar in
size (the smallest individual in the sample is no less than 75% of the total length of the largest
individual). Collection occurs in the sampling reach. Whole fish samples are shipped to the laboratory
designated for fish sample preparation. The laboratory fillets the fish and homogenizes the fillet tissue
for analysis of mercury and other contaminants (e.g., perfluorinated compounds).
12.2 Equipment and Supplies
Table 12.1 lists the equipment and supplies necessary for field crews to collect whole fish tissue
samples. This list is comparable to the checklist presented in Appendix A, which provides information to
ensure that field crews bring all of the required equipment to the site. Record the fish tissue sampling
data on the Whole Fish Tissue Collection Form (Figure 12.1).
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Table 12.1 Equipment and Supplies: Whole Fish Tissue Sample Collection
Q
o
For collecting whole
fish tissue sample
For storing and
preserving whole fish
tissue 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
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
whole fish tissue
sample
Whole Fish Tissue Collection Form
Clipboard
Clear tape strips
Sample Identification Labels
#2 pencils
Fine tipped indelible markers
For shipping the whole
fish tissue samples
Preaddressed FedEx airbill
Coolers
Tracking Form
Chain-of-custody labels
Packing/strapping tape
12.3 Sampling Procedures
The whole fish tissue samples will be collected using the same gear and procedures used to collect the
fish assemblage samples. Collection of individual specimens for whole fish samples occurs in the sample
reach during the fish assemblage sampling. Ideally, each fish sample will contain 5 fish of the same
species that are similar in size. Depending on the size of the fish, fewer than 5 fish may be acceptable or
more than 5 fish will be necessary to meet the 500-gram tissue requirement for chemical analysis and
archived tissue (refer to Frequently Asked Questions in the sampling kits). If sufficient fish are not
collected during the fish assemblage sampling, sample for up to one additional hour. If no fish can be
collected, call the Field Logistics Coordinator at the end of the day and record "no sample collected" on
the whole fish tissue collection form, along with the reason in the comments section of the 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 fish sample with adequate tissue for analysis) to obtain a whole fish sample
from the species that are available. Recommended target species are given in Table 12.2. The
procedures for collecting and processing whole fish tissue samples are presented in Table 12.3.
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Table 12.2 Recommended Target Species for Whole Fish Tissue Collection
Family name
Common name
Scientific name
Length Guideline
(Estimated
Minimum)
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Centrarchidae
Ictaluridae
Percidae
Moronidae
Spotted bass
Largemouth bass
Smallmouth bass
Black crappie
White crappie
Channel catfish
Blue catfish
Flathead catfish
Sauger
Walleye
Yellow perch
White bass
Northern pike
Chain pickerel
Brown trout
Cutthroat trout
Rainbow trout
Brook trout
Micropterus punctulatus
Micropterus salmoides
Micropterus dolomieu
Pomoxis nigromaculatus
Pomoxis annularis
Ictalurus punctatus
Ictalurus furcatus
Pylodictis olivaris
Sander canadensis
Sander vitreus
Perca flavescens
Morone chrysops
Esox lucius
Esox niger
Sal mo trutta
Oncorhynchus clarkii
Oncorhynchus mykiss
Salvelin us fan tin alis
~280 mm
~280 mm
~300 mm
~330 mm
~330mm
~300 mm
~300 mm
~300 mm
~380 mm
~380 mm
~330mm
~330 mm
~430 mm
~430 mm
~300mm
~300 mm
~300 mm
~330 mm
Table 12.3 Procedure: Whole Fish Tissue Samples
Whole Fish Tissue Method
1. Put on clean nitrile gloves before handling the fish. Do not handle any food, drink, sunscreen,
or insect repellant until after the whole fish 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. Collect one target species sample from each designated site. The sample should consist of 5
fish of adequate size to provide a total of 500 grams of edible tissue for analysis (refer to
4.
5. Table 12.2 for minimum species length guidelines). Select fish for each sample 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,
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Whole Fish Tissue Method
but no more than one week apart (Note: Individual fish may have to be frozen until all
fish to be included in the sample are available for delivery to the designated
laboratory).
Accurate taxonomic identification is essential in assuring and defining the organisms that
have been collected and submitted for analysis. Do not mix species in a single whole fish
sample.
6. 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).
7. Record site ID, date, sample ID, species (common name), and specimen length on the Whole
Fish Tissue Collection Form (Figure 12.1) in black ink. Fill in site type ("Wadeable" or
"Boatable") at the top of the form. Address the two sample criteria in the space above the fish
specimen data to confirm compliance. All samples must meet these two criteria (i.e., fish are
all the same species and fish lengths are all within 75% of the largest specimen length). Make
sure the sample ID numbers and specimen numbers/length s that are recorded on the
collection form match the corresponding information on each individual specimen label.
8. Remove each fish selected 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).
9. Wrap each fish in extra heavy-duty aluminum foil with the dull side in (foil provided by EPA as
solvent-rinsed, oven-baked sheets).
10. Prepare a Sample Identification Label for each sample, ensuring that the label information
matches the information recorded on the Whole Fish Tissue Collection Form.
11. 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).
12. Place all the wrapped fish in the sample from each site in a large plastic bag and seal with
another cable tie.
13. 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.
14. If possible, keep all (five) specimens designated for a particular sample in the same shipping
container (ice chest) for transport.
15. 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
• 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).
16. Ship fish tissue samples to the designated laboratory for fish sample preparation on Monday
through Thursday (no Saturday delivery to the laboratory).
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NRSA 2013/14 WHOLE FISH TISSUE COLLECTION
O WADEABLE Q BOATABLE
WHOLE FISH TISSUE FILLET SAMPLE (FTIS) SAMPLE ID:
NO SAMPLE COLLECTED Q
O FISH ARE ALL THE SAME SPECIES
O f ISH ALL WITHIN 75% OF LARGEST SPECIMEN
.01
.02
.03
.04
.05
.06"
.07*
.08*
.09*
.10*
Common Name
Total
Length (mm)
O
O
O
O
•Additional specimens for smaBer fish species to ensure sufficient tissue is available for chemical analysis of fiBet tissue
04/08/2013 2013 Fish Tissue Collection
Figure 12.1 Whole Fish Tissue Collection Form
2227506532
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13 FINAL SITE ACTIVITIES
13.1 Overview of Final Site Activities
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, as well as 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 13.1.
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 13.1 Final Site Activities Summary
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13.2 General Site Assessment
Complete the Site Assessment Form (Figure 13.2) 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.
13.2.1 Elevation at Transect K
Record elevation at Transect K using your GPS device. To record this information, record the elevation
holding the GPS at approximately 3 feet above the surface of the water. Ensure that the numbers are
properly recorded for Transect K on the Assessment Form.
13.2.2 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 13.2). Include those that were observed while
on the site, while driving or walking through the site 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.
13.2.3 Site Characteristics
Record observations regarding the general characteristics of the site on the Site Assessment Form
(Figure 13.2). 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 in the days prior to sampling.
13.2.4 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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LJJ
H
>
NRSA 2013/14 ASSESSMENT (Front)
Date: /
R«v!ttt*dbv(.nltJBl):_
Elevation at transect K:
Oft Om
WATERSHED ACTIVITIES AND DISTURBANCES OBSERVED
(Intensity: Blank=Not observed, L=Low. M=Moderate, H=Heavy|
BLAMK FIELD INDICATES ABSENCE
o
Residential
Q O O R
O O O "
O 0 O
O O O P'P»s. Oalnt
O O
O O OR
O O O Br
O 0 O Sewage Treatment
Recreational
O O O HikingTiads
O 0 0 Parks.Campgrounds
O 0 O Pr>mrtwePflrfcs,CBmpin[
O 0 O TiHhfUKit
O 0 0 Siirfon Films. Scums,
or Slicks
Agricultural
O O O Cropland
000 Pasture
O 0 0 Livestock Use
O 0 O Orchards
O 0 O Poultry
O 0 O Feedlot
000 WaterWiHidrawal
Industrial
Q Q Q tnduslnaiPlants
O 0 O Minss/Ouames
O 0 0 Oil/Gas WeJte
O O O Power Plants
O 0 OLoggmg
Q 0 0 EvtdenceoFFire
O 0 O Od"s
O 0 O Ccmmsicill
Stream Management
O 0 OL»™.9
O 0 O Chwni
O 0 0 AnglingPressure
O O O Dredsong
O 0 O Chonnstemion
Q 0 Q Water LevetFluctuatHins
O © O Fish Slocking
O O O Dams
SITE CHARACTERISTICS (200m radius)
WATERBODY CHARACTER
PRISTINE: Q5 Q4 Q3 Ql Q1 Highly Disturbed
APPEALING: Q5 O4 O3 O2 O1 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 'X' O Forest O Agriculture O Range O Urban O Suburban/Town
If Forest, Dominant Age Class O o - 25 yrs O25-75yts O>75yrs
WEATHER
CONDITIONS AND LOCAL CONTACTS
OBSERVATIONS (e.g. accessibility, boating, fishing, swimming, health concerns):
04/08/2013 2013 Assessment
0351596807
Figure 13.2 Site Assessment Form
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13.3 Processing the Fecal Indicator, Chlorophyll-a, and Periphyton Samples
13.3.1 Equipment and Supplies (Fecal Indicator Filtering)
Table 13.1 provides the equipment and supplies needed for field crews to collect the fecal indicator
sample.
Table 13.1 Equipment and Supplies: Fecal Indicator Sample
For processing samples
For recording measurements
Nitrile gloves
sterile screw-cap 50-mL PP tube
Filtration apparatus with collection flask
Sterile filter holder, Nalgene 145/147
Vacuum pump (electric pump may be used if available)
Sterile phosphate buffered saline (PBS)
Osmotics 47 mm polycarbonate sterile filters
Sterile disposable forceps
Petri dishes (60 x 15, disposable)
2 sterile microcentrifuge tubes containing sterile glass beads
1 additional sterile microcentrifuge tube if collecting filter blank
Dry ice
Cooler
Field Operations Manual and laminated Quick Reference Guide
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 (2 vial labels and 1 bag label)
Filter blank label if collecting filter blank
Clear tape strips for covering labels
13.3.2 Procedures for Processing the Fecal Indicator Sample
The fecal indicator sample must be filtered before the chlorophyll-o 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 13.2. The sample must be filtered and frozen within 6 hours of collection.
Table 13.2 Procedure: Processing Fecal Indicator Sample
Filtering for the Enterococci (fecal indicator) Sample
1. Put on nitrile gloves.
2. Set up sample filtration apparatus on flat surface and attach vacuum pump. Set-out 50-mL sterile PP 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 2 polycarbonate filters from filter box to base of opened Petri dish. Close filter box
and set aside.
5. Remove the pre-loaded 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.
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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 PP 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. Pourapprox. 10-mLof 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). Filter should be inserted open end down
(pointed side up) into the tube. Replace and tighten the screw cap, insert tube(s) into ziplock 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 50-mL sub-sample volume to be filtered.
°rocessing procedure—fecal indicator filter blank
Enterococci filter blanks will be prepared at all revisit sites during the first visit. 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 1 polycarbonate filter from filter box to base of opened Petri dish. Close filter box and
set aside.
3. Remove the pre-loaded 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-mLof the chilled phosphate buffered saline (PBS) in the sterile graduated PP 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 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). Filter should be inserted open end down
(pointed side up) into the tube. Replace and tighten the screw cap, insert tube(s) into ziplock 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 this sample to the lab
with the standard samples.
10. Indicate that you have collected a filter blank by filling in the "Blank Collected" button on the Sample
Collection Form.
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13.3.3 Equipment and Supplies (Chlorophyll-a from Water Sample Filtering)
Table 13.3 provides the equipment and supplies needed to process the chlorophyll-o water sample.
Table 13.3 Equipment and Supplies: Chlorophyll-a Processing
For filtering chlorophyll-a sample
Whatman GF/F 0.7 urn glass fiber filter
Filtration apparatus with graduated filter holder and collection flask
Vacuum pump (electric pump may be used if available)
50-mL screw-top centrifuge tube
Aluminum foil square
250 mL graduated cylinder
Dl water
Nitrile gloves
Forceps
Dry ice
For recording measurements
Sample Collection Form
Sample labels
#2 pencils
Fine-tipped indelible markers
Clear tape strips
13.3.4 Procedures for Processing the Chlorophyll-a Water Sample
The procedures for processing chlorophyll-o water samples are presented in Table 13.4. Whenever
possible, sample processing should be done in subdued light, out of direct sunlight.
Table 13.4 Procedure: Chlorophyll-a Sample Processing
Filtering for the Chlorophyll a Water Sample
1.
2.
Put on nitrile gloves.
Use clean forceps to place a Whatman GF/F 0.7 u.m glass fiber filter in the graduated filter holder apparatus
with the gridded side of the filter facing down.
3. Retrieve the 2 liter chlorophyll sample bottle from the cooler and shake the bottle to homogenize the
sample. While filtering sample, keep the bottle in the cooler on ice.
4. Measure 250 mLof water with a graduated cylinder and pour into the filter holder, replace the cap, and use
the vacuum pump to draw the sample through the filter (do not exceed 7 inches of Hg). 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.
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. Rinse the upper portion of the filtration apparatus and graduated cylinder thoroughly with Dl water to
include any remaining cells adhering to the sides and pump through the filter. Monitor the level of water in
the lower chamber to ensure that it does not contact the filter or flow into the pump. Remove the bottom
portion of the apparatus and pour off the water from the bottom as often as needed.
7. Remove filter funnel from base without disturbing filter.
8. Remove the filter from the holder with clean forceps. Avoid touching the colored portion of the filter. Fold
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the filter in half, with the colored side folded in on itself.
9. Place the folded filter into a 50-mL screw-top centrifuge tube and cap. Tighten the cap as tightly as possible.
The cap will seal tightly after an additional % turn past the point at which initial resistance is met. Failure to
tighten the lid completely could allow water to infiltrate into the sample and may compromise its integrity.
10. Record the sample volume filtered on a chlorophyll label and attach it to the centrifuge tube (do not cover
the volume markings on the tube). Ensure that all written information is complete and legible. Cover with a
strip of clear tape.
11. Wrap the tube in aluminum foil and place in a self-sealing plastic bag. Place this bag immediately on dry ice
to freeze.
13.3.5 Equipment and Supplies (Periphyton Sample)
Table 13.5 lists the equipment and supplies needed to process the periphyton sample.
Table 13.5 Equipment and Supplies: Periphyton Samples
For preparing
periphyton samples
For data recording
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 collection flask and graduated filter holder
Vacuum pump (electric pump may be used)
25 or 50-mL graduated cylinder
Pipette and pipette bulb (2 mL)
3 50 mL screw-top centrifuge tubes
60-mL syringe with tip removed
Aluminum foil squares
Forceps
deionized water in wash bottle
plastic electrical tape
dry ice
wet ice
coolers
formalin
Sample Collection Form
Sample labels
Pencils
Fine-tipped indelible markers
Clear tape strips
13.3.6 Procedures for Processing the Periphyton Samples
Three different types of laboratory samples are prepared from the composite index samples: an
ID/enumeration sample (to determine taxonomic composition and relative abundances), chlorophyll a
sample, and a biomass sample (for ash-free dry mass [AFDM]). 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.
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13.3.6.1 ID/Enumeration Sample
Prepare the ID/Enumeration sample as a 50-mL aliquot from the composite index sample, following the
procedure presented in Table 13.6. Preserve each sample with 2 ml of formalin. Record the sample ID
number from the container label and the total volume of the periphyton sample in the appropriate
fields on the Sample Collection Form as shown in Figure 4.2. Store the preserved samples upright in a
container containing absorbent material.
Table 13.6 Procedure: ID/Enumeration Samples of Periphyton
Periphyton ID Sample Processing Procedure
1. Prepare a sample label (with pre-printed sample ID number sample type "PERI"). 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 Sample Collection Form.
3. Rinse a 60-mL syringe with deionized water.
4. Thoroughly mix the bottle containing the composite sample.
5. Immediately after mixing, withdraw 50 mLof the mixed sample into the syringe and place the
contents of syringe into the labeled 50-mL centrifuge tube.
6. Use a syringe or bulb pipette to add 2 mL of 10% formalin to the tube. Cap the tube tightly and seal
with plastic electrical tape. Tighten the cap as tightly as possible. The cap will seal tightly after an
additional % turn past the point at which initial resistance is met.
7. Shake gently to distribute preservative.
8. 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.
13.3.6.2
Periphyton Chlorophyll a Sample
Prepare the periphyton chlorophyll a 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 periphyton chlorophyll a
samples is presented in Table 13.7. Chlorophyll a 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. If using the same filtration chamber that was used for Enterococci and index
site chlorophyll-o samples, rinse it with deionized water prior to filtering the periphyton chlorophyll-o
sample. If you are reusing a filtration chamber from a previous site, you should rinse it with Dl water
each day before use at the base site and then seal in a plastic bag until use at the stream (be sure to use
a new chamber at each site for the Enterococci sample as it needs to be filtered in a sterile chamber).
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 13.7 Procedure: Preparing Chlorophyll Samples of Periphyton
Periphyton Chlorophyll a Sample Processing Procedure
Using clean forceps, place a Whatman GF/F 0.7 urn glass fiber filter on the filter holder gridded
side down. If needed, use a small amount of deionized water from a wash bottle to help settle
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the filter properly. Attach the filter funnel to the filter holder and filter chamber, and then attach
the vacuum pump to the filter flask.
2. Rinse the sides of the filter funnel and the filter with a small volume of deionized water to
prevent contamination from the previously filtered sample.
3. Rinse a 25-mLor 50-mL graduated cylinder three times with small volumes of deionized water
and discard.
4. Mix the composite sample bottle thoroughly.
5. Measure 25 mL (±1 mL) of sample into the graduated cylinder.
NOTE: For a 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 vacuum 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. Monitor the level of water in the lower chamber to ensure that it does not contact the filter or
flow into the pump. Remove the bottom portion of the apparatus and pour off the water from
the bottom as often as needed.
8. Remove the filter chamber from the filter holder being careful not to disturb the filter. 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.
9. Tighten the cap as tightly as possible. The cap will seal tightly after an additional % turn past the
point at which initial resistance is met.
10. Prepare a sample label (with pre-printed sample ID number sample type "PCHL") including the
volume filtered, and attach it to the centrifuge tube. Cover the label completely with a strip of
clear tape.
11. Place the centrifuge tube into a self-sealing plastic bag.
12. Record the sample ID number of the label and the total volume of the composite index sample
on the Sample Collection Form. Record the volume filtered in the "Periphyton 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.
13. Place the centrifuge tube containing the filter on dry ice.
13.3.6.3
Periphyton Biomass Sample (AFDM)
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 13.8. Using the same filtration chamber that was used for Enterococci and
chlorophyll-a samples, rinse it with deionized water prior to filtering the periphyton biomass sample. If
you are reusing a filtration chamber from a previous site, you should rinse it with Dl water each day
before use at the base site and then seal in a plastic bag until use at the stream (be sure to use a new
chamber at each site for the Enterococci sample as it needs to be filtered in a sterile chamber). 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
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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 13.8 Procedure: Preparing Ash-Free Dry Mass (AFDM) Samples of Periphyton
Periphyton AFDM Sample Processing Procedures
1. Using clean forceps, place a Whatman 47 mm 1.2 micron GF/C glass fiber filters on the filter holder
gridded side down. If needed, 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 filter flask.
2. Rinse the sides of the filter funnel and the filter with a small volume of deionized water to prevent
contamination from the previously filtered sample.
3. Rinse a 25-mL or 50-mL graduated cylinder three times with small volumes of deionized water and
discard.
4. Mix the composite sample bottle thoroughly.
5. Measure 25 mL (±1 mL) of sample into the graduated cylinder.
NOTE: For a 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 vacuum 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. Monitor the level of water in the lower chamber to ensure that it does not contact the filter or flow
into the pump. Remove the bottom portion of the apparatus and pour off the water from the
bottom as often as needed.
8. Remove the filter chamber from the filter holder being careful not to disturb the filter. 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.
9. Tighten the cap as tightly as possible. The cap will seal tightly after an additional % turn past the
point at which initial resistance is met.
10. Prepare a sample label (with pre-printed sample ID number sample type "PBIO"), 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.
11. Record the sample ID number of the label and the total volume of the composite index sample on
the Sample Collection Form. Record the volume filtered in the "Periphyton 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.
12. Place the centrifuge tube containing the filter on dry ice.
13.4 Data Forms and Sample Inspection
After the Site Assessment Form is completed, the Field Crew Leader reviews all of the data forms and
sample labels for accuracy, completeness, and legibility. The other crew 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.
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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 is 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.
13.5 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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14 FIELD QUALITY CONTROL
Standardized training and data forms provide the foundation to help assure that data quality standards
for field sampling are met. This section for field sampling and data collection are the primary guidelines
for all cooperators and field crews. In addition, repeat sampling and field evaluation and assistance visits
will address specific aspects of the data quality standards for the National Rivers and Streams
Assessment.
14.1 Revisit Sampling Overview
1
Visit 1
I
Revisit Sites (4 per State)
1
Space revisits minimum of two
weeks to one m onth apart
1 1
Collect all
samples
In situ measures
Water chemistry
Chlorophyll-a
Periphyton
Benthos
Enterococci
Fish
Fish plugs
Fish tissue
(at select sites)
Physical habitat
Filter Blank
Enterococci
BEFORE filtering
other samples
Revisits = Measurement
Variation + Index period
variation
i
Visit 2
i-1
Collect all
Samples
In situ measures
Water chemistry
Chlorophyll-a
Periphyton
Benthos
Enterococci
Fish
Fish plugs
Physical habitat
Figure 14.1 Summary of the Revisit Sampling Design
Revisit 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 revisit sampling design is provided in
Figure 14.1.
14.2 Revisit Sampling Sites
A total of 200 (approximately 10% ) of the target sites visited will be revisited during the same field
season by the same field crew that initially sampled the site. Revisit samples and measurements are
taken from the same reach as the first visit. Each state has four revisit sites; two wadeable and two
non-wadeable sites. For each state these sites are:
Wadeable Revisit sites:
• The two wadeable revisit sites are re samples from the NRSA 2008/09 (1-4th order). They are in
the base sites in NRSA09, and have _RVT in the panel column. For most States, these are the
first two sites in that category, but this is not always the case.
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Non-Wadeable Revisit sites:
• The two non-wadeable revisit sites are resamples from the NRSA 2008/09 (5th and above
order). They are in the base sites in NRSA09, and have _RVT in the panel column. For most
States, these are the first two sites in that category, but this is not always the case.
If a site selected for revisit sampling is dropped, then an alternate assigned to replace it should be
revisited. The alternate site is the next site on the sampling list, in the same category (i.e. Repeat NRSA
1-4th order). If a non-wadeable site is sampled with wadeable methods, the next non-wadeable site
should be selected as the revisit 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 is, not less than 2 weeks and not more than one
month. Label the samples Visit 2 to indicate that they are samples from revisits sites. We will not be
collecting replicate data on whole fish tissue. Whole fish tissue samples will only be collected on the first
visit.
At each revisit site, a filter blank will be collected for Enterococci during the first sampling visit (Visit 1).
The crews will filter a small amount (10 ml) of sterile buffer through 1 filter, 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 first field visit
(Visit 1) (Figure 14.1). Detailed description of the filter blanks is found in Table 13.2.
14.3 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 quality assurance/quality control (QA/QC) check for the uniform evaluation of the
data collection methods, and an opportunity to conduct procedural reviews as required minimizing data
loss due to improper technique or interpretation of field procedures and guidance. Through uniform
training of field crews 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 crew collecting and contributing data under this program (EPA will
make a concerted effort to evaluate every crew, but will rely on the data review and validation process
to identify unacceptable data that will not be included in the final database).
14.3.1 Specifications for QC Assurance Field Assistance Visits
Field evaluation and assistance personnel are trained in the specific data collection methods detailed in
O 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
O 14.1 summarizes the plan, the checklist, and corrective action procedures.
=i It is anticipated that evaluation and assistance visits will be conducted with each Field Crew early in the
13 sampling and data collection process, and that corrective actions will be conducted in real time. If the
Q Field Crew misses or incorrectly performs a procedure, the Evaluator will note this on the checklist and
LU immediately point this out so the mistake can be corrected on the spot. The role of the Evaluator is to
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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.
Table 14.1 General Information Noted During Field Evaluation
Field
Evaluation
Plan
Field Logistics Coordinator will arrange the field evaluation visit with each Field
Crew, ideally within the first two weeks of sampling.
The Evaluator will observe the performance of a crew through one complete set of
sampling activities.
If the Crew 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 Crew 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 Crew has followed NRSA protocols for
locating the site.
The Evaluator observes the complete set of sampling activities, confirming that all
protocols are followed.
The Evaluator will record responses or concerns, if any, on the Field Evaluation and
Assistance Check List.
Corrective
Action
Procedures
If the Evaluator's findings indicate that the Field Crew is not performing the
procedures correctly, safely, or thoroughly, the Evaluator must continue working
with this Field Crew until certain of the Crew's ability to conduct the sampling
properly so that data quality is not adversely affected.
If the Evaluator finds major deficiencies in the Field Crew operations the Evaluator
must contact a NRSA QA Project Coordinator.
14.4 Reporting
When the sampling operation has been completed, the Evaluator will review the results of the
evaluation with the Field Crew 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 Crew 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 Crew members must read and sign
off on the evaluation.
If the Evaluator's findings indicate that the Field Crew is not performing the procedures correctly, safely,
or thoroughly, the Evaluator must continue working with this Field Crew until certain of the Crew's
ability to conduct the sampling properly so that data quality is not adversely affected. If the Evaluator
finds major deficiencies in the Field Crew operations (e.g., major misinterpretation of protocols,
equipment or performance problems) the Evaluator must contact the following QA official:
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Sarah Lehmann, EPA National Rivers and Streams Assessment Project QA Officer
The QA Officer will contact the Project Manager to determine the appropriate course of action. Data
records from sampling sites previously visited by this Field Crew will be checked to determine whether
any sampling sites must be redone.
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