National Rivers and Streams Assessment 2023/24
Version 1.0, January 2023
Field Operations Manual
Wadeable
vvEPA
United States Environmental Protection Agency
Office of Water
Washington, DC
EPA-841-B-22-006
National Rivers and Streams
Assessment 2023/24
Field Operations
Manual
Wadeable
Version 1.0
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National Rivers and Streams Assessment 2023/24 Field Operations Manual
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National Rivers and Streams Assessment 2023/24 Field Operations Manual
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NOTICE
The complete documentation of overall National Rivers and Streams Assessment (NRSA) project
management, design, methods, and standards is contained in four companion documents,
including:
National Rivers and Streams Assessment 2023-24: Quality Assurance Project Plan EPA-841-B-22-
004
National Rivers and Streams Assessment 2023-24: Site Evaluation Guidelines EPA-841-B-22-005
National Rivers and Streams Assessment 2023-24: Wadeable Field Operations Manual EPA-841-
B-22-006
National Rivers and Streams Assessment 2023-24: Non-Wadeable Field Operations Manual EPA-
841-B-22-007
National Rivers and Streams Assessment 2023-24: Laboratory Operations Manual EPA-841-B-22-
008
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, algal toxins, fish
assemblage, fish tissue plugs, Enterococci, antimicrobial resistance, and physical habitat. These
methods are based on the guidelines developed and followed in the National Rivers and Streams
Assessment 2008-2009 (USEPA 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 NRSA
2023/24. 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 documents. This document and associated QAPP ensure compliance
with the EPA policies Assuring the Competency of EPA Laboratories (2004) and Policy to Assure
Competency of Laboratories, Field Sampling, and Other Organizations Generating Environmental
Measurement Data under Agency-Funded Acguisitions (2011).
The suggested citation for this document is:
USEPA. 2022. National Rivers and Streams Assessment 2023/24: Field Operations Manual -
Wadeable. EPA-841-B-22-006. U.S. Environmental Protection Agency, Office of Water
Washington, DC.
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TABLE OF CONTENTS
NOTICE II
TABLE OF CONTENTS IV
LIST OF FIGURES VII
LIST OF TABLES VII
ACRONYMS/ABBREVIATIONS IX
DISTRIBUTION LIST XI
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 3
1.5 Supplemental Material to the Field Operations Manual 4
1.6 Recording Data and Other Information 5
2 INTRODUCTION TO WADEABLE SAMPLING 8
2.1 Daily Operations 8
2.2 Bas e S ite Acti viti es 10
2.2.1 Pre-departure Activities 10
2.2.2 Post Sampling Activities 12
2.3 Safety and Health 15
2.3.1 General Considerations 15
2.3.2 Safety Equipment 16
2.3.3 Safety Guidelines for Field Operations 17
2.4 Forms (NRSA App) 18
2.4.1 Field Forms 18
2.4.2 Tracking Form and Packing Slips 19
2.4.3 Equipment and Supplies 20
3 INITIAL SITE PROCEDURES 22
3.1 Site Verification Activities 22
3.1.1 Locating the X-Site 22
3.1.2 Determining the Sampling Status of a Stream 23
3.1.3 Elevation at Transect A 25
3.1.4 Sampling During or After Rain Events 25
3.1.5 Site Photographs 26
3.2 Laying outthe sampling reach 26
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
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Z 4 IN SITU MEASUREMENTS OF DISSOLVED OXYGEN, PH, TEMPERATURE, AND CONDUCTIVITY 32
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5 4.1 Summary of Method 32
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u 4.2 Equipment andSupplies 32
q 4.2.1 Multi-Probe Sonde 32
4.3 Sampling Procedure 33
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£ 5 COLLECTION OF ANTIMICROBIAL RESISTANCE SAMPLES 33
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5.1 Summary of Method 33
5.2 Equipment andSupplies 33
5.3 Sampling Procedure 34
5.4 Antimicrobial Resistance Field Blanks 35
6 COLLECTION OF WATER CHEMISTRY AND CHLOROPHYLL-^ SAMPLES 35
6.1 Summary of Method 35
6.2 Equipment andSupplies 36
6.3 Water Chemistry and Chlorophyll-^ Sampling Procedure 36
7 ALGAL TOXINS (MICROCYSTES AND CYLINDROSPERMOPSIN) 37
7.1 Summary of Method 38
7.2 Equipment andSupplies 38
7.3 Sampling Procedure 38
8 BENTHIC MACROINVERTEBRATES 40
8.1 Summary of Method 40
8.2 Equipment andSupplies 40
8.3 Sampling Procedure 41
8.4 Sample Processing in Field 45
9 PERIPHYTON 47
9.1 Summary of Method 47
9.2 Equipment andSupplies 47
9.3 Sampling Procedure 47
9.4 Sample Processing in the Field 49
10 PHYSICAL HABITAT CHARACTERIZATION 50
10.1 Equipment andSupplies 50
10.2 Summary of Methods Approach 50
10.3 Components of the Habitat Characterization 51
10.4 Work Flow for the Physical Habitat Components 52
10.4.1 Channel/Riparian Cross-Sections 52
10.4.2 Thalweg Profile and Large Woody Debris Tally 52
10.4.3 Channel Constraint and Torrent Evidence 53
10.5 Habitat Sampling within the Reach 53
10.5.1 Channel and Riparian Measurements at Cross-Section Transects 55
10.5.2 Thalweg Profile and Large Woody Debris Measurements 70
10.6 Cross-section Transects on Side Channels 76
10.7 Slope and Bearing 77
10.7.1 Measurement of Slope using Level and Stadia Rod 77
10.7.2 Alternate Methods for Obtaining Slope 80
10.7.3 Method for Obtaining Bearing 82
10.8 Channel Constraint, Debris Torrents, and Recent Floods 83
10.8.1 Channel Constraint 83
10.8.2 Debris Torrents and Recent Major Floods 85
10.9 Elevation at Transect K 86
11 FECAL INDICATOR (ENTEROCOCCI) 87
11.1 Summary of Method 87
11.2 Equipment andSupplies 87
11.3 Sampling Procedure 87
11.4 Sample Processing in the Field 88
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12 FISH ASSEMBLAGE 89
12.1 Summary of Method 89
12.2 Equipment andSupplies 89
12.3 Sampling Procedures 90
12.3.1 Irruptive Species 91
12.3.2 Small Wadeable Streams 93
12.3.3 Large Wadeable Streams 96
12.4 Seining 100
12.5 Processing Fish 103
12.5.1 Identification and Tallying 103
12.5.2 Unknown Specimens 104
12.5.3 Photovouchering 109
12.5.4 Preparing Preserved Voucher Specimen Samples 109
12.5.5 Preserving Voucher Specimen Samples 110
12.5.6 Processing Unknown/Range Extension (UNK/RNG) Voucher Samples 112
12.5.7 Processing QA Voucher Samples 112
13 FISH TISSUE PLUG SAMPLING METHODS 116
13.1 Method Summary 116
13.2 Equipment and Supplies 116
13.3 Sample Collection Procedures 117
14 FINAL SITE ACTIVITIES 121
14.1 Overview of Final Site Activities 121
14.2 General Site Assessment 122
14.2.1 Elevation at Transect K 122
14.2.2 Watershed Activities and Disturbances Observed 122
14.2.3 Site Characteristics 122
14.2.4 General Assessment 122
14.3 Processing the Fecal Indicator (Enterococci), Chlorophyll-^, and Periphyton Samples 123
14.3.1 Equipment and Supplies (Fecal Indicator Filtering) 123
14.3.2 Procedures for Processing the Fecal Indicator (Enterococci) Sample 123
14.3.3 Equipment and Supplies (Chlorophyll-a from Water Sample Filtering) 125
14.3.4 Procedures for Processing the Chlorophyll-a Water Sample 125
14.3.5 Equipment and Supplies (Periphyton Sample) 126
14.3.6 Procedures for Processing the Periphyton Samples 127
14.4 Data Forms and Sample Inspection 131
14.5 Launch Site Cleanup 131
15 FIELD QUALITY CONTROL 132
15.1 Revisit Sampling Overview 132
15.2 Revisit Sampling Sites 132
15.3 Field Evaluation and Assistance Visits 133
15.3.1 Specifications for QC Assurance Field Assistance Visits 134
^ 15.4 Reporting 134
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g 16 REFERENCES 136
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g APPENDIX A LIST OF EQUIPMENT AND SUPPLIES A-l
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u- APPENDIX B SAMPLE FORMS B-l
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^ APPENDIX C SHIPPING AND TRACKING GUIDELINES C-l
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H APPENDIX D COMMON & SCIENTIFIC NAMES OF FISHES OF THE UNITED STATES D-l
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APPENDIX E EXAMPLE ELECTROFISHING SETTINGS E-l
LIST OF FIGURES
Figure 1.1 Example Sample Labels for Sample Tracking and Identification 7
Figure 2.1 Field Sampling Scenario for Wadeable Sites 9
Figure 2.2 Overview of Base Site Activities 10
Figure 2.3 Example Sample Packing Slips to Accompany Sample Shipments 19
Figure 2.4 Electronic Request Form 20
Figure 3.1 Sampling Reach Features for a Wadeable Site 28
Figure 8.1 Benthic Macroinvertebrate Collection at Wadeable Sites 41
Figure 8.2 Transect Sample Design for Collecting Benthic Macroinvertebrates at Wadeable Sites 42
Figure 10.1 Reach Layout for Physical Habitat Measurements for streams 2.5m or greater (plan view).... 54
Figure 10.2 Substrate Sampling Cross-Section 55
Figure 10.3 Riparian Zone and Instream Fish Cover Plots for a Stream Cross-Section Transect 58
Figure 10.4 Determining Bank Angle Under Different Types of Bank Conditions 61
Figure 10.5 Schematic Showing Relationship Between Bankfull Channel and Incision 63
Figure 10.6 Determining Bankfull and Incision Heights 64
Figure 10.7 Schematic of Modified Convex Spherical Canopy Densiometer 65
Figure 10.8 Proximity Classes for Human Influences in Wadeable Streams 68
Figure 10.9 Large Woody Debris Influence Zones (modified from Robison and Beschta, 1990) 75
Figure 10.10 Large Woody Debris Section of Physical Habitat Form in the NRSA App - Wadeable 75
Figure 10.11 Riparian and Instream Fish Cover Plots for a Stream with Minor and Major Side Channels .. 76
Figure 10.12 Measurements of Bearing Between Transects 78
Figure 10.13 Channel Slope Measurement using a Clinometer 81
Figure 10.14 Types of Multiple Channel Patterns 85
Figure 12.1 Reach Layouts for Fish Sampling at Wadeable Sites 92
Figure 12.2 Unknown/Range Extension Voucher Sample Labels and Tags Ill
Figure 12.3 QA Voucher Sample Labels and Tags 113
Figure 14.1 Final Site Activities Summary 121
Figure 15.1 Summary of the Repeat Sampling Design 132
LIST OF TABLES
Table 1.1 Summary Table of Indicators for all NRSA 2023/24 Sites 3
Table 1.2 Guidelines for Recording Field Measurements and Tracking Information 6
Table 2.1 Stock Solutions, Uses, and Methods for Preparation 12
Table 2.2 Post-sampling Equipment Care 14
Table 2.3 General Health and Safety Considerations 16
Table 2.4 General Safety Guidelines for Field Operations 18
Table 3.1 Equipment and Supplies: for Site Verification 23
Table 3.2 Procedure: Site Verification 24
Table 3.3 Guidelines to Determine the Influence of Rain Events
Table 3.4 Procedure: Laying Out the Sampling Reach at Wadeable Sites
Table 3.5 Procedure: Sliding the Sampling Reach
Table 3.6 Reach Layout Modifications for Interrupted Streams
Table 3.7 Procedure: Modifications for Braided Rivers and Streams
Table 4.1 Equipment and Supplies: DO, pH, Temperature, and Conductivity
Table 4.2 Procedure: Temperature, pH, Conductivity and Dissolved Oxygen
Table 5.1 Equipment and Supplies: Antimicrobial Resistance Samples
Table 5.2 Procedure: Antimicrobial Resistance Sample Collection (Wadeable Sites)
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Table 5.3 Procedure: Antimicrobial Resistance Field Blank Collection (Visit 1 to Revisit Sites) 35
Table 6.1 Equipment and Supplies: Water Chemistry Sample Collection and Preservation 36
Table 6.2 Procedure: Water Chemistry and Chlorophyll-o Sample Collection (Wadeable Sites) 36
Table 7.1 Equipment and Supplies: Microcystin 38
Table 7.2 Procedure: Algal Toxin (Microcystin and Cylindrospermopsin) Collection (Wadeable Sites) 38
Table 8.1 Equipment and Supplies: Benthic Macroinvertebrate Collection at Wadeable Sites 40
Table 8.2 Procedure: Benthic Macroinvertebrates (Wadeable Sites) 43
Table 8.3 Procedure: Compositing Samples for Benthic Macroinvertebrates (Wadeable Sites) 45
Table 9.1 Equipment and Supplies: Periphyton (Wadeable Sites) 47
Table 9.2 Procedure: Collecting Composite Index Samples of Periphyton (Wadeable Sites) 48
Table 10.1 Equipment and Supplies: Physical Habitat 50
Table 10.2 Summary of Components of Physical Habitat Characterization at Wadeable Sites 51
Table 10.3 Procedure: Substrate Measurement 56
Table 10.4 Procedure: Estimating Instream Fish Cover 59
Table 10.5 Procedure: Measuring Bank Characteristics 59
Table 10.6 Procedure: Canopy Cover Measurements 66
Table 10.7 Procedure: Characterizing Riparian Vegetation Structure 67
Table 10.8 Procedure: Estimating Human Influence 69
Table 10.9 Procedure: Thalweg Profile 71
Table 10.10 Channel Unit Categories 73
Table 10.11 Procedure: Tallying Large Woody Debris 74
Table 10.12 Procedure: Obtaining Slope Data 79
Table 10.13 Modified Procedure: Obtaining Slope Data (without Surveyor's Level) 80
Table 10.14 Procedure: Obtaining Bearing Data 82
Table 10.15 Procedure: Assessing Channel Constraint 83
Table 11.1 Equipment and Supplies: Fecal Indicator Sampling (Wadeable Sites) 87
Table 11.2 Procedure: Fecal Indicator (Enterococci) Sample Collection (Wadeable Sites) 87
Table 12.1 Equipment and Supplies: Fish Collection (Wadeable Sites) 89
Table 12.2 Summary of Wadeable Fishing Protocols 91
Table 12.3 Procedure: Electrofishing (Small Wadeable Streams) 93
Table 12.4 Procedure: Electrofishing (Large Wadeable Sites) 96
Table 12.5 Procedure: Seining (Wadeable Sites) 100
Table 12.6 Procedure: Processing Fish (Wadeable Sites) 105
Table 12.7 Procedure: Processing Unknown/Range Extension (UNK/RNG) Voucher Samples 114
Table 12.8 Procedure: Processing QA Voucher Samples 115
Table 13.1 Equipment and Supplies: Fish Tissue Plug Sample 116
Table 13.2 Recommended Target and Alternate Species for Fish Tissue Plug Collection 118
Table 13.3 Procedure: Fish Tissue Plug Samples 119
Table 14.1 Equipment and Supplies: Fecal Indicator (Enterococci) Sample Processing 123
Table 14.2 Procedure: Fecal Indicator (Enterococci) Sample Processing 123
Table 14.3 Equipment and Supplies: Chlorophyll-o Processing 125
Table 14.4 Procedure: Chlorophyll-o Sample Processing 126
Table 14.5 Equipment and Supplies: Periphyton Samples 126
Table 14.6 Procedure: ID/Enumeration Samples of Periphyton 127
Table 14.7 Procedure: Preparing Metagenomic Sample of Periphyton 128
Table 14.8 Procedure: Preparing Chlorophyll Samples of Periphyton 129
Table 14.9 Procedure: Preparing Periphyton Biomass Sample 130
Table 15.1 General Information Noted During Field Evaluation 134
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ACRONYMS/ABBREVIATIONS
AFDM Ash-Free Dry mass
AMR Antimicrobial resistance
ANC Acid-neutralizing capacity
Ca Calcium
CI Chloride
CPR Cardiopulmonary Resuscitation
CWA Clean Water Act
DELT Deformities, Eroded Fins, Lesions and Tumors
DO Dissolved Oxygen
DOC Dissolved Organic Carbon
Dl Di-ionized Tap Water
EPA Environmental Protection Agency
FOM Field Operations Manual
GIS Geographic Information System
GPP Generator Powered Pulsator
GPS Global Positioning Device
IBI Index of Biotic Integrity
IM Information Management
K Potassium
LIMS Laboratory Information Management System
LOM Lab Operations Manual
LWD Large Woody Debris
Mg Magnesium
MMI Multimetric Index
MSDS Methods Safety Data Sheets
Na Sodium
NAD North American Datum
NARS National Aquatic Resources Survey
NHD National Hydrology Database
NHs Ammonia
NH4 Ammonium
NIST National Institute of Standards and Technology
NO2 Nitrite
NO3 Nitrate
NRSA National Rivers and Streams Assessment
O/E "Observed" over "Expected"
OSHA Occupational Safety and Health Administration
OW Office of Water
PESD-AL Pacific Ecological System Division - Analytical laboratory
PETG polyethylene terephthalate glycol
PFD Personal Flotation Device 1/1
PP Polypropylene O
PPT Parts per thousand ^
Psig Pounds per square inch - gauge >
LU
QA Quality Assurance
CO
QAPP Quality Assurance Project Plan co
<
QA/QC Quality Assurance/Quality Control ^
QCS Quality Control Solution ^
QRG Quick Reference Guide z
SEG Site Evaluation Guideline §
Si02 Silica <
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so4
Sulfate
SOP
Standard Operating Procedures
Std
Standard
TL
Total Length
TOC
Total Organic Carbon
TN
Total Nitrogen
TP
Total Phosphorus
TSS
Total Suspended Solids
UNK/RNG
Unknown Range Extension
USGS
United States Geological Survey
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DISTRIBUTION LIST
This FOM and associated manuals or guidelines will be distributed to the following U.S.
Environmental Protection Agency (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.
National Monitoring Coordinators
Richard Mitchell
NRSA Project Leader
mitchell.richardPepa.gov
202-564-0064
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Sarah Lehman
NRSA Project QA
Officer
lehmann.sarahfSeoa.gov
202-566-1379
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Brian Hasty
NRSA Logistics Lead
hastv.brianfSeoa.gov
202-566-2236
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Lareina Guenzel NRSA
ESA Lead
Lareina.guenzelPepa.gov
202-566-0455
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Bernice L. Smith
WRAPDQA
Coordinator
smith.bernicelPepa.gov
202-566-1244
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Cynthia N. Johnson
QA Officer
Johnson.CvnthiaNPepa.gov
202-566-1679
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Amanda Nahlik, EPA
ORD Technical Advisor
Nahlik.Amanda@epa.gov
541-754-4581
U.S. EPA Office of Research and Development
200 S.W. 35th St. Corvallis, OR 97330
Michelle Gover
NARS Information
Management
Coordinator
gover. michellefSeoa. gov
541-754-4793
541-754-4799 (fax)
General Dynamics Information Technology
200 S.W. 35th Street
Corvallis, OR 9733
Chris Turner
Contract Field Logistics
Coordinator
cturner(5)glec.com
715-829-3737
Great Lakes Environmental Center, Inc.
739 Hastings Street
Traverse City, Ml 49686
Regional Monitoring Coordinators
Tom Faber
Region 1
Faber.tomfSeoa.gov
617-918-8672
U.S. EPA - Region 1
11 Technology Drive, North Chelmsford, MA 01863-2431
Emily Nering
Region 2
nering.emilvfaJeoa.gov
732-321-6764
U.S. EPA-Region II
2890 Woodbridge Ave, Edison, NJ 08837-3679
Leah Ettema
Region 3
Ettema. leahPepa.gov
215-814-5675
U.S. EPA-Region III
1650 Arch Street, Philadelphia, PA 19103-2029
Elizabeth Belk
Region 4
belk.elizabethfSeoa.gov
404-562-9377
U.S. EPA - Region IV
61 Forsyth Street S.W., Atlanta, GA 30303-8960
Mari Nord
Region 5
nord. mari Pepa.gov
312-353-3017
U.S. EPA-Region V
77 West Jackson Blvd, Chicago, IL 60604-3507
Rob Cook
Region 6
cook, robe rt Pepa.gov
214-665-7141
U.S. EPA-Region VI
1445 Ross Ave -Ste 1200, Dallas, TX 75202-2733
Laura Webb
Region 7
webb.laurafSeoa.gov
913-551-7435
U.S. EPA-Region VII
300 Minnesota Ave, Kansas City, KS 66101
Liz Rogers/Shera
Reems
Region 8
rogers.lizfSeoa.gov
/reems.sherafSeoa.gov
303-312-6974/303-312-6888
U.S. EPA-Region VIII
1595 Wynkoop Street, Denver, CO 80202-1129
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Matthew Bolt
Region 9
bolt. matthewfaJeoa.gov
415-972-3578
U.S. EPA-Region IX
75 Hawthorne Street, San Francisco, CA 94105
Lillian Herger
Region 10
Herger.lillian(5)eDa.gov
206-553-1074
U.S. EPA-Region X,
1200 Sixth Avenue, Seattle, WA 98101
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1 BACKGROUND
This manual describes field protocols and daily operations for crews to use in the Wadeable
NRSA 2023/24 method. 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 NRSA 2008/09, NRSA 2013/14, and NRSA 2018/19.
• 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. 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
The survey design consists of two separate designs to address the dual objectives of: (1)
estimating current status and (2) estimating change in status for all flowing waters:
• Resample design applied to NRSA 2018/19 sites.
• New site design for NRSA 2023/24.
The Resample survey design is a group of sites that were targeted for sampling in NRSA 18/19.
These sites all begin with the NRS23_18 prefix. The major objective for this design is change
estimation. This results in 790 resample base sites which are part of the 2023/24 design.
Allocation of sites to NARS aggregated ecoregions is proportional to the number sampled in the
prior surveys. The NRSA 2023/24 survey design includes 1028 new sites which have not been
included in previous NRSA surveys. Allocation of a number of sites to states is proportional to
stream length and is stratified by state. Unequal probability categories are 27 combinations of
NARS nine aggregated ecoregions and three waterbody reach categories (SS - small streams, LS
- large streams, and RV - Rivers). A minimum of 20 sites was guaranteed in each state and the
maximum number of sites was set at 75 for an individual state. The sample frame was derived
from the high-resolution National Hydrography Dataset (NHD), in particular NHDPIus V2.
Additional details on the NRSA survey design are found in the National Rivers and Streams
Assessment Survey Design: 2023/24 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, including major rivers and small streams. Sites
must have > 50% of the reach length with standing water, and 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 < 0.5 ppt for this study).
• Run-of-the-river ponds and reservoirs with greater than seven-day residence time.
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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-22-005 and the NRSA Web site
http://www.epa.gov/national-aquatic-resource-survevs/nrsa) for more detailed information on
the target population and exclusion criteria.
1.3 Replacing Sites
All base sites must be evaluated for sampling. If a stream or river site is determined to be
nontarget or otherwise unsampleable, it must be replaced by another site within the same state
and same panel. The panels for NRSA 2023/24 are listed below. Note, the FT suffix found in
some panels in the site lists refers to "Fish Tissue" sites which may not be applicable to the
evaluation process though have been left in should EPA obtain funding for the collection of fish
for fish tissue purposes. The evaluation process is the same regardless of the FT suffix and those
suffixes have been omitted from the list below.
• NRS23_18RVT2RV: River sites that were selected for sampling in NRSA 2018/19 and
have been selected to be sampled twice in 2023/24. This is referred to as a "revisit site"
as it will be evaluated and sampled twice during the same year, with at least two weeks
in between each visit.
• NRS23_18BaseRV: River sites that were selected for sampling in NRSA 2018/19 and have
been selected to be sampled once in 2023/24
• NRS23_180verRV: River sites to be used as replacements for dropped 18/19 river base
sites or as additional sites for state level surveys in some cases.
• NRS23_18RVT2LS: Large stream sites that were selected for sampling in NRSA 2018/19
and have been selected to be sampled twice in 2023/24. This is referred to as a "revisit
site" as it will be evaluated and sampled twice during the same year, with at least two
weeks in between each visit.
• NRS23_18BaseLS: Large stream sites that were selected for sampling in NRSA 2018/19
and have been selected to be sampled once in 2023/24
• NRS23_180verLS: Large stream sites to be used as replacements for dropped 18/19
large stream base sites or as additional sites for state level surveys in some cases.
• NRS23_18BaseSS: Small stream sites that were selected for sampling in NRSA 2018/19
and have been selected to be sampled once in 2023/24
• NRS23_180verSS: Small stream sites to be used as replacements for dropped 18/19
small stream base sites or as additional sites for state level surveys in some cases.
• NRS23_23BaseRV: River sites new to NRSA 2023/24, designated to be sampled once in
2023/24.
• NRS23_230verRV: River sites to be used as replacements for dropped 23/24 river base
sites or as additional sites for state level surveys in some cases. May also be used to
replace 2018/19 river base sites if no 18/19 river oversample sites remain.
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• NRS23_23BaseLS: Large stream sites new to NRSA 2023/24, designated to be sampled
once in 2023/24.
• NRS23_230verLS: Large stream sites to be used as replacements for dropped 23/24
large stream base sites or as additional sites for state level surveys in some cases. May
also be used to replace 2018/19 large stream base sites if no 18/19 large stream
oversample sites remain.
• NRS23_23BaseSS: Small stream sites new to NRSA 2023/24, designated to be sampled
once in 2023/24.
• NRS23_230verSS: Small stream sites to be used as replacements for dropped 23/24
small stream base sites or as additional sites for state level surveys in some cases. May
also be used to replace 2018/19 small stream base sites if no 18/19 small stream
oversample sites remain.
Please refer to the Site Evaluation Guidelines (EPA- 841-B-22-005J for more detailed
information.
1.4 Selection of NRSA Indicators
As part of the indicator selection process, EPA worked with state, tribal, 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 2023/24
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, NRSA
2013/14, and NRSA 2018/19. 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 2023/24 are briefly described in Table 1.1.
Table 1.1 Summary Table of Indicators for all NRSA 2023/24 Sites
Indicator
Core or Supplemental
Indicator
Specs/Location in Sampling Reach
In Situ measurements (pH, DO,
temperature, conductivity)
Core Indicator
Measurements taken at X site at mid-
channel; readings are taken at 0.5 m
depth, or mid-depth if water depth is less
than 1 meter.
Water chemistry (TP, TN, NH3-
N, NO3-NO2, NO3, basic anions
and cations, silica, alkalinity
[Acid-neutralizing capacity
(ANC)], DOC, TOC, TSS,
conductivity, pH, turbidity, true
color)
Core Indicator
Collected at the X site at mid-channel;
from a depth of 0.5 m, or mid-depth if
water depth is less than 1 meter.
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Indicator
Core or Supplemental
Indicator
Specs/Location in Sampling Reach
Chlorophyll-o
Core Indicator
Collected as part of water chemistry and
periphyton samples
Microcystin and
cylindrospermopsin
Supplemental Indicator
Collected at the X site at mid-channel;
from a depth of 0.5 m, or mid-depth if
water depth is less than 1 meter.
Periphyton composite and
periphyton metagenomic
Supplemental Indicator
Collected from 11 locations
systematically placed at each site and
combined into a single composite sample
Benthic macroinvertebrate
assemblage (Littoral)
Core Indicator
Collected from 11 locations
systematically placed at each site and
combined into a single composite sample
Fish Assemblage
Core Indicator
Sampled throughout the sampling reach
at specified locations
Physical habitat assessment
Core Indicator
Measurements collected throughout the
sampling reach at specified locations
Fecal indicator (Enterococci)
Supplemental Indicator
Collected at the last transect one meter
off the bank at 0.3 m depth
Antimicrobial resistance (genes
and bacteria)
Supplemental Indicator
Collected at the X site at mid-channel;
from a depth of 0.5 m, or mid-depth if
water depth is less than 1 meter.
Fish Tissue Plug
Supplemental Indicator
Tissue sample collected from target
species which are collected throughout
the sampling reach as part offish
assemblage sampling
1.5 Supplemental Material to the Field Operations Manual
The FOM describes wadeable 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.
Field Crews will collect field data in the NRSA App on an iPad. Within the App, users will find a
number of information (/') buttons that contain tables and figures summarizing field activities
and protocols from the FOM. These informational buttons replace the need for a Quick
Reference Guide (QRG) as has been used in previous survey years. In addition to the
informational buttons within the App, crews will also have access to this FOM in electronic
(Adobe® PDF) format on the iPad. Field Crews are required to know how to access and search
the FOM in the field for reference and for possible protocol clarification.
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Quality Assurance (QA) 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 digital
copy of the integrated 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 QA procedures, refer to the National Rivers
and Streams Assessment: Quality Assurance Project Plan (EPA-841-B-22-004).
Related NRSA documents include the following: National Rivers and Streams Assessment:
Quality Assurance Project Plan (EPA 841-B-22-004), National Rivers and Streams Assessment:
Site Evaluation Guidelines (EPA 841-B-22-005), and National Rivers and Streams Assessment:
Laboratory Methods Manual (EPA-841-B-22-008). These documents are available at:
http://www.epa.gov/national-aquatic-resource-survevs/nrsa.
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 1.1).
Field and sample information must be recorded accurately and consistently. 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. Guidelines for recording field measurements
are presented in Table 1.2. At the end of each sampling day, the Field Crew Leader is
responsible for reviewing each field form for completeness and accuracy. Field Crews will find a
number of data validation routines within each electronic form that will help find missing or
possibly incorrect data. These routines can be accessed at any time by tapping the data review
button at the top of each form.
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Table 1.2 Guidelines for Recording Field Measurements and Tracking Information
ACTIVITY GUIDELINES
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Field Measurements
Data Recording
• Record observations and measurement values only using the official NRSA App
(provided on EPA owned Apple iPads for all regional, state, and tribal crews).
• If you make an error when recording data and changes are required, it is best to enter
the new value and resubmit that electronic form.
• Use the correct crew ID assigned during field training.
• Use the units and formats specified on individual data forms for recording data.
• For any sample or data where additional explanation is needed, use the provided
comment bubble adjacent to the data
Sample Collection
Sample Labels and
Tags
• Use a writing instrument that leaves clear, dark text to record information (e.g., a No. 2
pencil on paper tags or a water or smear proof fine-point indelible marker on adhesive
labels). Use the sample-type appropriate adhesive labels with preprinted Sample ID
numbers for each sample. Be sure to fill in any requested information about the
sample on the sample label and affix it to the outside of the sample container. Cover
completed labels with clear tape.
• Place a waterproof paper tag inside each benthic macroinvertebrate collection jar and
fish voucher jar with the required information written with a No. 2 lead pencil.
Sample Collection
Information
• Record that each sample has been collected on the appropriate data form. Be sure to
cross-check the Sample ID number from labels and tags with the Sample IDs populated
in the Tracking Form.
QA and Tracking
Before Leaving Site:
Review of Data
Forms and
Comparison of
Sample Labels and
Data Forms
• Review all data forms for accuracy and completeness.
• Review all sample labels for accuracy, completeness, and legibility.
• Verify that the information recorded on the sample labels and tags is consistent with
all Sample IDs listed on the Tracking Form.
Before Shipping
Samples: Review of
Sample Labels and
Tracking Forms
• Complete all tracking forms required for all samples being shipped. Review tracking
forms for consistency and correctness.
• Compare labels on samples with the Sample IDs recorded on the Tracking Form for
accuracy and completeness before shipping samples.
Review of Data
Forms
• The Field Crew Leader should review the completed forms in the NRSA App as soon
as is practicable to ensure they are complete, and all data forms are consistent and
correct
• Confirm that the forms have been reviewed by selecting the reviewed bubble in the
App for each electronic data form.
• If any revisions are made, re-submit the updated form(s) as soon as possible to
update the IM Database.
• After each submission, a data summary email will be sent to the email address which
submitted the data. This data summary contains a list of the data forms and their
most recent submissions date/time as well as a list of the most critical data points
collected at the site. The Field Crew Leader should review this data summary to
ensure that the data forms were successfully received and that critical values are
present and correct.
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WATER CHEMISTRY (CHEM)
Site ID: NRS23
Date:
/
T1
/202 Visit#: 01 02
999000
PERIPHYTON BIOMASS (PBIO)
Site ID: NRS23
Date:
T1
/
/202_
Volume Filtered:
Visit#: 01 02
mL
999002
WCHL, PBIO, PCHL - OUTER BAG
Site ID: NRS23
Date:
/
1202
Visit#: 01 02
999001,999002,999003
T1
PERIPHYTON METAGENOMIC (PDNA)
Site ID: NRS23
Date:
/ 202
Visit#: 01 02
"
999005
BENTHIC MACROINVERTEBRATES (BERW)
(Wadeable Sites)
Site ID: NRS23
Date:
/
1202 Visit #: 01 02
Jar 1 of
999007
BACTERIA CULTURE (BCUL)
Site ID: NRS23
Date: / 1202 Visit#: 01 02
T2
/202
Jar 1 of
999009
FISH TISSUE PLUG (FPLG)
Site ID: NRS23
Date: /
T3
_/202_ Visit#: 01 02
999011
Anchor ID: 999000
Site ID: NRS23
_Visit#: OlO 2
CHLOROPHYLL-a (WCHL)
Site ID: NRS23
Date:
T1
/ 202
Visit#: 01 02
Volume Filtered: mL
999001
PERIPHYTON CHLOROPHYLL (PCHL)
Site ID: NRS23
ALGAL TOXIN (MICX)
Site ID: NRS23
Date:
1202 Visit #: 01 02
999006
BENTHIC MACROINVERTEBRATES (BETB)
(Boatable Sites)
Site ID: NRS23
Date:
/
/202 Visit#: 01 02
Figure 1.1 Example Sample Labels for Sample Tracking and Identification.
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2 INTRODUCTION TO 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 wadeable site will typically consist of four people. Any additional crew
members may either help collect samples, or may remain on the bank to provide logistical
support. Typically, in wadeable sites, two crew members will work on the "habitat" crew, and
two or three will work on the "fish" crew.
A daily field sampling scenario showing how the workload 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 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. For example, at sites where fish collections are expected to take
longer than physical habitat assessments, crews may choose to task Group A with in situ
measurements and water collections, but these tasks need to be completed before any tasks
near or directly upstream of the water collection are to begin.
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Whole Crew
Lay out sampling reach (from X- site to Transect A)
Begin sampling activities at Transect A
Conduct physical habitat
characterizations
Including slope & bearing
Collect benthic samples
Collect periphyton samples
Collect fecal indicator sample at last transect
Measure stream discharge
Return to staging area
Filter Enterococci sample
Must be filtered and frozen within 6 hours of collection
Filter chlorophyll-a sample and periphyton samples
Preserve benthic and periphyton samples
Lay out sampling reach (from X- site to Transect K)
Return to Transect F (X-site)
Measure in situ temperature
pH, DO, & conductivity
Collect antimicrobial resistance
samples directly from the water
(0.3 meters below surface)
Collect water chemistry, chlorophyll-a,
and algal toxin samples
(use 3 L beaker as collection vessel)
Travel to Transect A
Conduct fish assessment
Collect fish tissue plug samples if possible
Collect fish vouchers at designated sites
Return to staging area
Clean and organize equipment for loading Inventory supplies for next sampling event
Request additional supplies if needed
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Check and prepare all samples for transport or shipment
Inspect and clean boats, motors, and trailers to prevent transfer of
nuisance species and contaminants
Review App data forms for completeness
Submit data via the App / Ship samples / Submit App Tracking
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Figure 2.1 Field Sampling Scenario for Wadeable Sites
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2.2 Base Site Activities
Field Crews conduct several 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.
PREDEPARTURE ACTIVITIES
Crew Leader
- Prepare daily itinerary
Crew Members
- Instrument checks & calibration
- Equipment & supplies preparation
Whole Crew
- Site Verification
POST SAMPLING ACTIVITIES
Crew Leader
- Review App forms & labels
- Submit data and tracking via the NRSA App
- Order site kits or replacement items
via Request Form
- Obtain ice and other consumable supplies
as needed
Crew Members
- Filter, preserve & inspect samples
- Clean boats with 1% bleach solution and perform safety
Checks (boat, trailer, & equipment)
- Package & ship samples
- Charge or replace batteries
- Refuel vehicle and boat
Figure 2.2 Overview of Base Site Activities
2.2.1 Pre-departure Activities
Pre-departure activities include the development of 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
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.
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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.
• 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) when handling chemicals.
Site kits of consumable supplies for each sampling site will be delivered based on the supply
requests each crew submits prior to and during the sampling season. Crews will submit an
electronic request form to order site kits, labels, etc. Site kit requests must be submitted at least
two weeks before sampling is to take place. If your schedule (and therefore your supply needs)
changes, report the change as soon as possible to the Contract Field Logistics Coordinator (Chris
Turner, cturner@glec.com, 715-829-3737). The site kit will include sample labels, sample jars,
bottles, shipping materials, 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 river/stream. Store at least (J
one extra site kit in the vehicle in the event replacement items are needed immediately. zi
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Table 2.1 Stock Solutions, Uses, and Methods for Preparation
Solution
Use
Preparation
Bleach (1%)
Clean nets, other gear, and boat.
Add 40 mL bleach to 4L distilled water.
Bleach (10%)
Clean periphyton sampling
equipment.
Add 40 mL bleach to 400 mL distilled water.
10% Buffered
Formalint
Preservation of periphyton ID sample
and fixing Fish Vouchers
Formaldehyde (37-40%) 100 ml
Distilled water 900 ml
NaH2P04 4.0 g
Na2HPC>4 (anhydrous) 6.5 g
Mix to dissolve
95% Ethanol
Preservative for benthic invertebrate
samples and fish vouchers.
No preparation needed (use stock solution as
is).
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 must review the data summary email received
after each data submission. 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 and completeness. Ensure that comments use no "shorthand" or abbreviations. The
data forms must be thoroughly reviewed by the Field Crew Leader. Upon completing the review,
the Field Crew Leader should select the data reviewed button at the top of each form in the App
which will run a series of data validation routines to help identify missing or potentially incorrect
data. 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 populated on
w the Tracking Form in the App.
^ 2.2.2.2 Inspect and Prepare Samples
<
1/1 All samples need to be inspected and appropriately preserved and packaged for transport.
^ Check that all samples are labeled, and all labels are filled in completely. Check that each label is
55 covered with clear plastic tape (with the exception of the Enterococci labels). 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
i— immediate shipment.
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q 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
z Eurasian watermilfoil (Myriophyllum spicatum), zebra mussels (Dreissena polymorpha), New
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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.protectvourwaters.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 trailer, and waders, and clean off any plant and animal material. Inspect, clean, and
handpick plant and animal remains from vehicle, boat, motor, trailer and waders. Before moving
to the next site, if a commercial car wash facility is available, wash the vehicle, boat, and trailer
and rinse thoroughly (hot water pressurized rinse with no soap). Rinse equipment and boat with
1% -10% bleach solution or other specialized disinfectant 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. Requests must be made two weeks before
needed. Note that not all supplies can be replenished by EPA through the Logistics Contractor,
so crews will need to supply some items themselves.
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Table 2.2 Post-sampling Equipment Care
Equipment Care after Sampling
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1. Clean for biological contaminants.
Prior to departing site, drain all water from live wells and buckets used to hold and process fish.
Inspect 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 is provided below:
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 minute.
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 all containers 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.
3. Inventory equipment and supply needs and relay orders to the IM Team via the supply request
form.
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 and no soap).
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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 (Tlotemersch, et al. (editors)
2000). Additional sources of information regarding safety-related training include the American
Red Cross (1979), the National Institute for Occupational Safety and Health (1981), U.S. Coast
Guard (1987) and Ohio EPA (1990).
Field Crew members should become familiar with the hazards involved with sampling
equipment and establish appropriate safety practices prior to using them. 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). A personal floatation device (PFD)
must be worn by crew members at all times on the water. All boats with motors must have fire
extinguishers, boat horns, PFDs 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 Field 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 ej
clothing instead of cotton. If it is necessary for a crew member to reach into the water to pick up zi
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
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45° tilt switch that interrupts the current. Do not make any modifications to the electrofishing <
unit that would hinder safety. 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 Field Crew Leader has authority, each crew g
member has the responsibility to question and modify an operation or decline participation if it g
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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
¦Cell (or satellite) phone and VHF 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
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 PFD 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,
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o rickettsia, fungi, or parasites.
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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
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3 available in the field. Cellular or satellite telephones and/or portable radios should be provided
O to Field Crews working in remote areas in case of an emergency. Supplies (e.g., clean water,
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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]; USEPA [1986]).
During the course of field research activities, Field Crews may observe violations of
environmental regulations, discover improperly disposed hazardous materials, or observe or be
involved with an accidental spill or release of hazardous materials. In such cases it is important
that the proper actions be taken and that field personnel do not expose themselves to
something harmful. The following guidelines should be applied:
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. i
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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.
¦ 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 venomous 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, Lyme disease,
and other illnesses.
¦ Field personnel should be familiar with the symptoms of hypothermia and know what to do in
case symptoms occur. Hypothermia can kill a person at temperatures much above freezing (up to
10°C or 50°F) if he or she is exposed to wind or becomes wet.
¦ Field personnel should be familiar with the symptoms of heat/sun stroke and be prepared to
move a suffering individual into cooler surroundings and hydrate immediately.
¦ Handle and dispose of chemical wastes properly. Do not dispose any chemicals in the field.
2.4 Forms (NRSAApp)
w Forms are the key to data collection and tracking for the NRSA 2023/24. Field Crews must use
^ electronic forms to submit their field data. If the NRSA App or iPad isn't available, collect
^ information on the provided paper data forms but data must be transferred to the App to be
^ submitted.
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2.4.1 Field Forms
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^ Field forms are the primary documents where crews 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.
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O • Electronic Field Forms: This form of data collection can be collected through an Apple
t iOS portable electronic device (tablets). The field forms will be optimized for tablet
o devices. The App will be preloaded on all EPA-owned iPads prior to distribution to Field
oc Crews or can be downloaded from the Apple App Store and installed on the Field Crews'
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iPads. It is important for a Field Crew to familiarize themselves with the App prior to
field sampling.
2.4.2 Tracking Form and Packing Slips
The Tracking Form in the App describes the status and location of all samples collected during
NRSA 2023/24. Field Crew Leaders will submit the pertinent portion(s) of this form electronically
through App submissions) whenever samples are shipped. See APPENDIX C: SHIPPING
GUIDELINES for more information.
Packing slips (Figure 2.3) are postcard sized slips pre-populated with sample IDs that match the
sample labels. Packing slips are included in the site kit with sample labels. Packing slips are to
accompany samples sent to any of the national labs.
For Lab Staff only:
Lab Sample ID:
Tl: NRSA23/24 Daily Water Chemistry Sample Tracking
Site ID: NRS23_ Visit#: Q 1 O 2
Date Collected:
I !
Sample ID Lab Comments (For lab staff only)
CHEM
999000
WCHL
999001
PBIO
999002
PCHL
999003
Rec'd by: Date Rec'd:
T2: NRSA23/24 Daily Bacteria Sample Tracking
Site ID: NRS23_ Visit #:Q102
Date Collected:
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Lab Staff:
Samples In Box? Sample ID Lab Comments (For lab staff only)
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BCUL
999009
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BDNA
999010
(13
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Please cross out sample id(s) not included in shipment.
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AMR Field
Blank included
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T3: NRSA23/24 Frozen Batched Tracking
Site ID: NRS23_ Visit #: O 1 O 2
Date Collected:
I I.
T4: NRSA23/24 Non-Chilled Batched Tracking
Site ID: NRS23_ visit #: Q 1 O 2
Date Collected:
I .
Lab Staff:
Samples In Box? Sample ID Lab Comments ( For lab staff only)
~
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ENTE 999013
PDNA
999005
MICX
999006
f|Slg 999011
Please cross out sample id(s) not included in shipment.
Lab Staff:
Samples In Box? Sample ID Lab Comments (For lab staff only)
For Lab use only
BERW
999007
BETB
999008
PERI
999004
Please cross out sample id(s) not included in shipment. 9901360974
Figure 2.3 Example Sample Packing Slips to Accompany Sample Shipments
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2.4.3 Equipment and Supplies
2.4.3.1 Request Form
Field Crews will submit requests for site kits or other supplies via an electronic request form
(Figure 2.4). This form will be submitted to the NARS IM Coordinator who will ensure that the
request reaches the appropriate entity. Crews should submit the Request Form at least two
weeks prior to their desired sampling date. In addition to typing in specific requests, crews may
select one or more of the pre-populated items listed below:
• Site kit: contains all consumable supplies for one site, sample labels, packing slips, FedEx
shipping labels to PESD-AL and EPA-Cincinnati, coolers (for sending immediate samples
to PESD-AL and EPA-Cincinnati), and cooler liners.
• Frozen batched box: contains box, two-piece dry ice liner, Class 9 hazardous label, FedEx
label to GLEC.
• Non-chilled batched cooler: contains cooler, cooler liner, FedEx label to GLEC
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NRSA23/24 Request Form
Request Date: / /
Requester:
Phone:
Email:
REQUESTER EMAIL
Ship to:
Name:
Company:
Address:
City:
SHIP TO EMAIL, IF DIFFERENT THAN REQUESTER
Supplies Needed (Mark all that apply):
O Site Kit (2 coolers with all bottles and consumables for 1 site, including label packet and
packing slips, T1 & T2 shipping labels, and 2 cooler liners)
How many?
Need by:
/
/
Comments:
O Frozen Batched Dry Ice box - up to 4 sites max (includes box; 2-piece dry ice liner, T3 shipping label,
and Class 9 hazardous label)
How many? Need by: / / Comments:
O Non-chilled batch cooler (includes 1 cooler, T4/T5 shipping label and cooler liner)
How many? Need by: / / Comments:
O Replacement Label Racket(s)
How many? Need by:
O Foil squares (aluminum) - pack of 25
How many? Need by: /
O 1L Benthic/Fish Voucher Bottles (HDPE, white, wide-mouth)
How many? Need by: / / Comments:
O Sodium Thiosulfate Tablets - vial of 25
How many? Need by: /
O Tape strips - packs of 25
How many? Need by:
/
/
Don't see your item listed above? List items separately below. Refer to equipment lists in FOMfor correct terminology
How many?
How many?
How many?
How many?
Need by:
Need by:
Need by:
Need by:
12/07/2022 NRSA23 Request Form
7571010607
Figure 2.4 Electronic Request Form
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2.4.3.2 Base Kit
The Base Kit is comprised of the subset of durable equipment and supplies needed for NRSA
2023/24 sampling that is provided by EPA 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 EPA in the Base Kit.
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 EPA through the Field Logistics Contractor. The site
kit will contain all the sample bottles and labels necessary for sampling a single site. A new site
kit is provided (upon request) for each site sampled. See APPENDIX A: EQUIPMENT & SUPPLIES
for the consumable items that will be provided by EPA.
2.4.3.4 Field Crew Supplied Items
The Field Crew will also supply particular items for the field sampling day. These items might
include supplies from a previous NRSA, typical field equipment (like a GPS), or a boat. 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 that you are at the correct site, and then
determine if the site meets the criteria for sampling and data collection activities (See Site
Evaluation Guidelines EPA-841-B-22-005). 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.
All field data will be recorded and submitted through the NRSA App. As a Crew begins to sample
a site, they access the App and select the correct state, site ID, and visit number (visit 1 for all
sampling events except the second visit to a revisit site). The final selection before data entry
may begin is the selection of wadeable versus boatable sampling methods (see following
section). The choice of wadeable or boatable will allow the user to access the correct set of field
forms for the given sampling method.
The first form that is typically completed is the Verification Form. Upon opening this form, enter
the sampling date by using the "today" button or by entering the date using the specified
format. If a site is sampled over multiple days, use the date the water chemistry sample was
collected as the official sampling date. Typically many key site activities will take place on this
date, which is why it will be used as the official sampling date. The date entered on the
Verification Form will populate to other forms as well, but those dates can be changed manually
if needed. Enter or verify the site name (if known) and enter the crew ID that was assigned to
the sampling crew prior to the field season.
3.1 Site Verification Activities
3.1.1 Locating the X-Site
River and stream sampling points were chosen using the medium resolution National
Hydrography Dataset (NHD), in particular NHD-Plus HR, 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 evaluation
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 evaluation 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 in the App in the Measured Latitude and Longitude fields. Also record
the location of the coordinates (X-site in the case of wadeable sites), the number of satellites
your GPS is using and all methods used to verify the site. If the provided coordinates are located
on dry land near a stream, move the coordinates to the nearest blue line in NHD-Plus HR during
the desktop reconnaissance. Note this movement on the site evaluation spreadsheet and in the
comments section of the Verification Form.
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Table 3.1 Equipment and Supplies: for Site Verification
For locating and
verifying site
Sampling permit and landowner access (if required)
Field Operations Manual (PDF Format loaded onto iPad)
Site dossier, including access information, site spreadsheet with map
coordinates, street and/or topographic maps with "X-site" marked
NRSA 2023/24 Community Fact Sheet
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)
For recording
measu rements
Verification Form in NRSA App
Fine-tipped indelible markers to write on flagging
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 access to the site. By using the information below, the Field
Crew will ultimately determine if the site will be sampled or not. In the App, select Yes on No to
the question "Did you sample this site". The selection will open either a list of sampling methods
or a list of specific reasons why the site was not sampled.
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 20 channel-widths upstream and downstream of
the X-site - for small systems, 150 m is the minimum reach length that can be sampled, so the
upstream and downstream lengths would be 75 meters each. 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 in the App. 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
in the App. If the site is non-sampleable or inaccessible, no further sampling activities are
conducted. Replace the site with the appropriate replacement site (Section 1.3). i/i
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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 = NAD83). Record these on the Verification Form.
3. Use all available means to ensure 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 20 channel-widths
upstream and downstream of the X-site (150 m is minimum sampling reach length, so in small
systems the upstream and downstream lengths would be 75 meters each). 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.
Boa table: 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
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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" in the site evaluation spreadsheet.
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.
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Other: The site could not be sampled on that particular day, but is still a target site. Examples might
include a recent precipitation event that has caused unrepresentative conditions.
No Access to Site Categories
Access Permission Denied: You are denied access to the site by the 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).
Other: explain in the comments section.
6. Do not sample non-target or "Non-sampleable" or "No Access" sites. Select 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.
7. If the reason for not sampling the site is a temporary condition, the site should be scheduled
for sampling at a later time. The subsequent sampling event, if successful, will use the same
visit number and the crew will simply update the previous information recorded on the
Verification Form and submit the form with the new data.
3.1.3 Elevation at Transect A
Record the 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 Verification Form in the App. You
will use this same method to record the elevation at Transect K at the end of the sampling day
and record that value on the Assessment 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 £2
event, do not sample the site that day. 3
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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. Most cell phone cameras
also have the ability to attach geographical location data to a particular picture. Alternatively,
start the sequence with one photograph of an 8.5 x 11 inch piece of paper with the site ID,
waterbody name, and date printed in large, thick letters. After the photo of the site ID
information, take at least two photographs at the X-site, one in the upstream direction and one
downstream. Take any additional photos you find interesting after these first three pictures.
Keep a log of your photographs and briefly describe each one. Photographs can be uploaded to
the NARS SharePoint site.
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
average 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 (wadeable sites).
When you arrive at the site, 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 in the App. If you did not have to slide the reach, the upstream and
downstream lengths will be equal.
Figure 3.1 illustrates the principal features of the established sampling reach for wadeable sites,
including the location of 11 cross-section transects used for collecting samples and physical
habitat measurements. The figure also shows the specific sampling stations on each transect for
collection of periphyton and benthic macroinvertebrate samples.
For each site, crews may prepare an optional map of the reach sampled to help explain specific
characteristics or sampling decisions. This can be drawn by hand or produced digitally with any
number of mapping programs. 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. The optional map can be uploaded to the NARS SharePoint site. If a map is uploaded,
check the "Crew scanned or submitted a site sketch" box on the Verification Form in the App.
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Table 3.4 Procedure: Laying Out the Sampling Reach at Wadeable Sites
1. Locate the X-site using the coordinates provided for the site.
2. Use a surveyor's rod, tape measure, or laser range finder to determine the wetted width of the
channel at five places of "typical" width within approximately five channel widths upstream
and downstream from the X-site. Average the five 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. Record both the average channel width and total intended reach length
on the Verification Form in the App.
4. 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).
5. 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 3.
6. 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. Record the upstream
and downstream reach lengths on the Verification Form in the App.
7. Starting at the X-site (or the new midpoint of the reach if it had to be adjusted as described in
Step 6), measure a distance of one-half the reach length down one side of the stream using a
GPS unit, laser rangefinder, or tape measure. Be careful not to "cut corners". Enter the channel
to make measurements only when necessary to avoid disturbing the stream channel prior to
sampling activities. This endpoint is the downstream end of the reach, and is flagged as Transect
A.
8. At Transect A, use the seconds display on a digital watch or other random number selection
method to select the initial sampling station for standard transect samples: l-3="Left", 4-
6="Center", 7-9=Right. Mark "L", "C", or "R" on the transect flagging; the three potential
collection points are roughly equivalent to 25%, 50%, and 75% of the channel width,
respectively. 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.
9. Measure 1/10 of the required reach length upstream from Transect A. Flag this spot as Transect
B. Assign the sampling station systematically after the first random selection following the
repeating pattern L, C, R as you move upstream (Figure 3.1).
10. Proceed upstream with the tape measure and flag the positions of nine additional transects
(labeled "C" through "K" as you move upstream) at intervals equal to 1/10 of the reach length.
Continue to assign the sampling stations systematically.
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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:
• Lack of landowner permission.
• 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. Record the upstream and downstream reach lengths on the Verification Form in
the App.
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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 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 6) will be collected at the X-site (even if the
reach has been adjusted by "sliding" it). If the X-site is dry and there is water elsewhere in the
sample reach, collect the sample from a location having water with a surface area >1 m2 and a
depth >10 cm.
Collect data for the physical habitat indicator along the entire sample reach from interrupted
streams, regardless of the amount of water present at each of 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. l2
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Table 3.6 Reach Layout Modifications for Interrupted Streams
3.3.2 Braided Rivers and Streams
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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 10). 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 x 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 on the Verification Form in the NRSA App about what you did. Also, add additional details
to the optional site map and submit it to EPA via the NARS SharePoint site. It is important to
remember that the purpose of the 40 x 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 two to three meander cycles (about six 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
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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 two to three meander cycles (about six pool riffle habitat sequences).
2. Make detailed notes on the Verification Form and optional site map about what you did.
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4 In Situ Measurements of Dissolved Oxygen, pH,
Temperature, and Conductivity
4.1 Summary of Method
Measure in situ DO, pH, water temperature, and conductivity using a calibrated multi-parameter
water quality meter (or sonde). Take the measurements mid-channel at the X-site. Take the
readings at 0.5 m depth. Measure the site depth accurately before taking the measurements. 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 in the App.
4.2 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
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
For recording measurements
Field Measurement Form in NRSA App
4.2.1 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 (e.g., ambient air saturated with water).
Note that DO should always be calibrated at the site and should never be calibrated at your base
location. 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 Standard Operating Procedures
(SOP). Ideally, a quality control solution (QCS) should be used that is similar in ionic strength to
the water samples you will be measuring.
Temperature Meter
Check the accuracy of the sensor against a thermometer that is traceable to the National
Institute of Standards and Technology (NIST) at least once per sampling season and record the
NIST thermometer reading and the sensor reading on the Field Measurement Form. These same
values can be recorded each time for the Field Measurement Form unless the accuracy check is
conducted again. The entire temperature range encountered in the NRSA should be
incorporated in the testing procedure and a record of test results kept on file.
Conductivity Meter
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Calibrate the conductivity meter prior to each sampling event. Calibrate the meter in
accordance with the manufacturer's instructions. Ideally, a QCS solution should be used that
incorporates the entire conductivity range encountered in the NRSA and a record of test results
kept on file.
4.3 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. Samples are taken mid-channel, at the X site, at a depth of 0.5 meters or at mid-depth if less
than 1 meter deep.
3. Lower the sonde in the water and measure DO, pH, temperature, and conductivity at 0.5 m
depth (or at mid-depth if less than 1 meter deep).
4. Record the measurements on the Field Measurement Form, noting whether the conductivity
value is corrected to 25°C.
5. Provide comments on any measurements that the crew feels needs further explanation or when
a measurement cannot be made.
5 Collection of Antimicrobial Resistance Samples
5.1 Summary of Method
Antimicrobial resistance is a naturally occurring phenomenon in the environment. However,
human activity can contaminate the environment with antibiotics, antifungals, organisms
resistant to these compounds, and genes that confer resistance traits, all of which can alter
natural levels of resistance. Rapid spread and emergence of antimicrobial resistance is
recognized as a global health problem, but scientists currently do not fully understand the risk of
resistance increasing in the environment on human, animal, or environmental health. Studying
antimicrobial resistance in surface waters is critical for expanding this knowledge since
waterways provide a way for resistance to spread throughout populations.
Collect two antimicrobial resistance samples in pre-sterilized bottles from the X-site at the
midpoint of the stream. One sample (Bacteria Culture) will be collected in a 2 L bottle, and the
other (Bacteria DNA) will be collected in a 1 L bottle. Both samples will be collected directly from
the stream without the use of any additional sampling equipment. Collect the samples by
submerging the mouth of each bottle to a depth of 0.3 meters and allowing the bottle to fill
completely. If the sampling location is shallow, submerge the mouth of the bottle to a lesser
depth and be careful not to contaminate the sample with bottom sediments or detritus. After
collection, store both samples on ice in a closed cooler.
5.2 Equipment and Supplies
Table 5.1 provides the equipment and supplies needed to collect antimicrobial resistance
samples at the X-site. Record the sample collection data on the Sample Collection Form in the
NRSA App.
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Table 5.1 Equipment and Supplies: Antimicrobial Resistance Samples
For collecting samples
Nitrile gloves
Pre-sterilized 2 L sample bottle (Bacteria Culture)
Pre-sterilized 1 L sample bottle (Bacteria DNA)
Plastic electrical tape
Cooler with ice
Field Operations Manual
For collecting field blanks (at Visit 1 to Revisit
sites)
Pre-sterilized 500 mL PETG bottle
Ultra-pure water (1000 mL bottle)
AMR Field Blank Label
For recording measurements
Sample Collection Form in NRSAApp
Sample label with pre-printed Sample ID
Clear tape strips
Fine tipped indelible markers for sample labels
5.3 Sampling Procedure
Table 5.2 presents step-by-step procedures for collecting antimicrobial resistance samples at
wadeable sites.
Table 5.2 Procedure: Antimicrobial Resistance Sample Collection (Wadeable Sites)
Collection of Antimicrobial Resistance Samples
1. Fill out the pertinent information (Site ID, visit, and date) on the antimicrobial resistance sample
labels.
2. Affix the BCUL label to the pre-sterilized 2 L sample bottle and cover with clear tape.
3. Affix the BDNA label to the pre-sterilized 1 L sample bottle and cover with clear tape.
4. Collect the antimicrobial resistance (bacteria culture and bacteria DNA) samples from the X-site in a
flowing portion of the stream near the middle of the transect. Use the same location as water
chemistry samples collection.
5. Collect the samples directly from the water column without the use of any other sampling devices.
Do not rinse the containers.
6. Put on clean, nitrile gloves.
7. Approach the sampling location slowly from downstream or downwind.
8. Remove the lid from the first sample bottle. Do not touch the inside of the sterile container, lip, or
lid. When opening sterile sampling containers, work rapidly. Open sterile sampling containers only
to admit the sample and close it immediately.
9. Lower the uncapped, inverted sample bottle to a depth of 1 foot (0.3 m) below the water surface,
avoiding surface scum, vegetation, and substrates. If the sampling location is shallow, submerge the
mouth of the bottle to a lesser depth and be careful not to contaminate the sample with bottom
sediments or detritus.
10. 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.
11. After removing the container from the water, discard a small portion of the sample to create 1 inch
of headspace in the bottle to allow for proper mixing before analysis.
12. Tightly close the lid and seal the cap with electrical tape before shipping.
13. Repeat steps 9-13 for the second sample bottle.
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14. Enter the depth of the sample collection (in meters) in the provided fields. This is the approximate
depth from the surface of the water to the mouth of the submerged container, not the depth of the
water at the sampling location.
Storage
Place both bottles in a cooler (on ice or water) and shut the lid. If a cooler is not available, place the
bottles in an opaque garbage bag and immerse it in the stream. Once the samples are placed on ice,
mark the "Chilled" boxes on the Sample Collection Form in the App.
If sample(s) are not collected, select the "No Sample Collected" box on the form and indicate the reason
why targeted sample(s) were not collected in the adjacent comment bubble(s).
5.4 Antimicrobial Resistance Field Blanks
Field blanks for the antimicrobial resistance samples will be collected at all revisit sites during
the first visit. The initial order each crew receives will contain the required equipment for the
crew's reported number of revisit sites (Table 5.1). Collect the field blank before collecting the
antimicrobial resistance samples. Table 5.3 presents step-by-step procedures for collecting
antimicrobial resistance field blanks at visit 1 to revisit sites.
Table 5.3 Procedure: Antimicrobial Resistance Field Blank Collection (Visit 1 to Revisit Sites)
Antimicrobial Resistance Field Blanks
1. Fill out the pertinent information (Site ID, visit, and date) on the antimicrobial resistance field blank
label.
2. Affix the label to the pre-sterilized 500 mL field blank bottle and cover it with clear tape.
3. Put on clean, nitrile gloves.
4. Open the provided 1000 mL bottle of ultra-pure water and pour 500 mL of water into the sterile
500 mL field blank bottle.
5. Tightly close the lid of the 500 mL sample bottle and seal the cap with electrical tape before
shipping.
6. The residual ultra-pure water and the 1000 mL bottle are to be discarded.
Storage
Place the 500 mL field blank bottle in the same cooler with the antimicrobial resistance samples (on ice
or water) and shut the lid. The field blank will be shipped to the lab along with the two stream samples.
On the Sample Collection Form in the NRSA App, check the "AMR Field Blank Collected" box to indicate
that a field blank was collected.
6 Collection of Water Chemistry and Chlorophyll-a Samples
6.1 Summary of Method
The water chemistry samples will be analyzed for total phosphorus (TP), total nitrogen (TN),
total ammonium (NH4), nitrate (N03), basic anions, cations, total suspended solids (TSS),
turbidity, acid neutralizing capacity (ANC), alkalinity, dissolved organic carbon (DOC), and total
organic carbon (TOC). Using a 3 L Nalgene beaker, collect a grab sample into one 4 L cube
container (for water chemistry) and one 2 L amber Nalgene bottle (for chlorophyll-a) from the X-
site at the midpoint of the stream. After collection, store all samples on ice in a closed cooler.
After you filter the chlorophyll-a sample, the filters must be kept frozen until ready to ship.
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6.2 Equipment and Supplies
Table 6.1 provides the equipment and supplies needed to collect water samples at the X-site.
Record the water sample collection and preservation data on the Sample Collection Form in the
NRSA App.
Table 6.1 Equipment and Supplies: Water Chemistry Sample Collection and Preservation
For collecting samples
Nitrile gloves
4 L cube container
2 L amber Nalgene bottle
3 L Nalgene beaker
Cooler with ice
Dry Ice
Plastic electrical tape
Dl water (for cleaning beaker and 2 L amber bottle between sites)
Field Operations Manual
For recording
Sample Collection Form in NRSA App
measu rements
Water Chemistry sample label with pre-printed Sample ID
Clear tape strips
Fine tipped indelible markers for sample labels
6.3 Water Chemistry and Chlorophyll-a Sampling Procedure
Table 6.2 presents step-by-step procedures for collecting water chemistry samples at wadeable
sites.
Table 6.2 Procedure: Water Chemistry and Chlorophyll-a Sample Collection (Wadeable Sites)
Sampling Procedure
Water Chemistry
1. Fill out the pertinent information (Site ID, visit number, 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 the X-site in a flowing portion near the middle of the stream. Be
sure to collect the water samples prior to any disturbance of the stream upstream of the X-
site.
3. Put on nitrile gloves. Make sure not to handle sunscreen or other chemical contaminants until
after the sample is collected or implement measures to reduce contamination by such
chemicals, if applied, such as washing, wearing long gloves, etc.
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 if needed
(the weight of the water alone while filling will often open the container sufficiently). NOTE: DO
NOT BLOW into the cube container or place your fingers inside the opening to expand it,
because this will cause contamination.
6. Fill the 3 L 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 two 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. Completely fill the cube container. Rinse the cube container lid with
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water. Eliminate any air space from the cube container by squeezing the closed container and
opening the cap slightly to allow air to escape, and cap it tightly. Make sure the cap is tightly
sealed and not on at an angle.
8. Seal the cap with plastic electrical tape before shipping.
Chlorophyll-a
9. Fill the 3 L 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 two more times.
10. 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.
Storage
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. Once
the water chemistry sample is placed on ice, mark the "Chilled" box on the Sample Collection Form in
the App.
Use the pertinent comment bubbles to note anything that could influence sample chemistry (heavy
rain, potential contaminants, etc.). If the sample was collected at the X-site as planned, select the X-site
bubble in the location field. If you had to move to another part of the reach to collect the sample,
select the "Other" bubble and place the letter of the nearest transect in the adjacent box. Record more
detailed reasons and/or information regarding the sampling location a in the provided comment
bubble(s).
If sample(s) are not collected, select the "No Sample Collected" box on the form and indicate the
reason why targeted sample(s) were not collected in the adjacent comment bubbles.
Record the Water Chemistry Sample ID on the Tracking Form in the NRSA App. This is also referred to
as the Anchor ID because it determines the numeric series from which all samples at the site will come.
By entering the water chemistry sample ID in the Tracking Form, the rest of the sample IDs will auto-
populate in the remainder of the Tracking Form. It is important to keep the labels and packing slips
organized so the sample IDs will match when shipping occurs.
7 ALGAL TOXINS (MICROCYSTES and
CYLINDROSPERMOPSIN)
Cyanobacteria naturally occur in surface waters. Under certain conditions, such as in warm
water containing an abundance of nutrients, they can rapidly form harmful algal blooms (HABs).
HABs can produce toxins known as cyanotoxins, which can be harmful to humans and animals.
Microcystin and cylindrospermopsin are two cyanotoxins known to occur in the surface waters
of the United States. Microcystins are the most widespread cyanobacterial toxins and primarily
affect the liver but can also affect the kidney and reproductive system.
Cylindrospermopsin is another commonly identified cyanotoxin found in U.S. waters. The
primary toxic effects of this toxin are damage to the liver and kidney.
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7.1 Summary of Method
The algal toxin (microcystin and cylindrospermopsin) sample is a grab sample taken from the X-
site. All Field Crews must collect a grab sample using the 3 L beaker to fill one 500 ml bottle.
Collect the sample after the in situ measurements and water chemistry samples are collected.
Store the sample on wet ice in a closed cooler.
7.2 Equipment and Supplies
Table 7.1 provides the equipment and supplies needed to collect the algal toxin sample at the X-
site. Record the water sample collection and preservation data on the Sample Collection Form in
the NRSA App.
Table 7.1 Equipment and Supplies: Microcystin
For collecting samples
Nitrile gloves
3 L Nalgene beaker
PETG bottle (500 mL, clear, square) - algal toxins (MICX)
Plastic electrical tape
Cooler with ice
Field Operations Manual
For recording
measu rements
Sample Collection Form in NRSA App
Algal toxin sample label with pre-printed Sample ID
Clear tape strips
Fine tipped indelible markers for labels
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7.3 Sampling Procedure
Table 7.2 presents step-by-step procedures for collecting the algal toxin (microcystin and
cylindrospermopsin) sample at wadeable sites.
Table 7.2 Procedure: Algal Toxin (Microcystin and Cylindrospermopsin) Collection (Wadeable Sites)
Microcystin Sample Collection
1. Fill out the pertinent information (Site ID, visit, and date) on the algal toxin label.
2. Affix the MICX label to the 500 mL PETG clear square Nalgene bottle. Completely cover the label
with clear tape.
3. Collect the algal toxin sample from the X-site in a flowing portion of the stream near the middle
of the transect.
4. Put on nitrile gloves. Make sure not to handle sunscreen or other chemical contaminants until
after the sample is collected or implement measures to reduce contamination by such
chemicals, if applied, such as washing, wearing long gloves, etc.
5. Rinse the 3 L Nalgene beaker three times with water and discard the rinse downstream.
6. Rinse the water sample collection container and lid three times with water, discard the rinse
downstream.
7. Fill the beaker with water and pour into the 500 ml Nalgene bottle to the 500 mL mark (or just
below the shoulder of the bottle), leaving headspace so that the bottle doesn't burst when
frozen.
8. Seal the cap with plastic electrical tape before shipping.
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Storage
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 500 ml bottle in an opaque garbage bag and immerse it in the stream.
Upon returning to your base site (hotel, lab, office, etc.), freeze the sample and keep it frozen until
shipping. Mark the "Frozen" box on the form to verify that the sample has been frozen.
If the sample is not collected, select the "No Sample Collected" box on the form and indicate the
reason why the targeted sample was not collected in the adjacent comment bubble.
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8 BENTHIC MACROINVERTEBRATES
8.1 Summary of Method
Collect a benthic macroinvertebrate composite sample using a D-frame net with 500 pirn mesh
openings. Take individual samples from the sampling stations at the 11 transects equally
distributed along the targeted reach (Figure 8.1). Multiple habitats will be encountered and
sampled using this approach. Habitats will include various types of bottom substrate as well as
woody debris, macrophytes, and leaf packs. Composite all sample material from all 11 sampling
locations and field preserve with ~95% ethanol.
8.2 Equipment and Supplies
Table 8.1 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 Benthic Collection Form in the NRSA App.
Table 8.1 Equipment and Supplies: Benthic Macroinvertebrate Collection at Wadeable Sites
For collecting
Modified kick net (D-frame with 500 nm
Small spatula, spoon, or scoop to
samples
mesh) and 52" handle
transfer sample
Watch with timer or stopwatch
Sample jars, 1 L HDPE plastic suitable
Sieve bucket with 500 nm mesh openings
for use with ethanol
(U.S. std No. 35)
95% ethanol, in a proper container
5 gallon bucket
Cooler (with absorbent material) for
Watchmakers' forceps
transporting ethanol & samples
Wash bottle, 1 L capacity labeled "STREAM
Plastic electrical tape
WATER"
Scissors
Funnel, with large bore spout
Field Operations Manual
For recording
Composite benthic sample labels with &
Soft (#2) lead pencils
measu rements
without preprinted ID Sample ID numbers
Fine-tip indelible markers
Blank labels on waterproof paper for inside of
Clear tape strips
jars
Benthic Collection Form in the App
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8.3 Sampling Procedure
Figure 8.1 summarizes how samples will be collected from wadeable sites. The transect sample
design for collecting benthic macroinvertebrates is shown in Figure 8.2. Collect a sample from 1
m downstream of each of the 11 cross-section transects at the assigned sampling station. The
process for selecting the sample stations is described in the Initial Site Procedures (Section 3). At
transects assigned a "Center" sampling point where the stream width is between one and two
net widths wide, pick either the "Left" or "Right" sampling point instead. If the stream is only
one net wide at a transect, place the net across the entire stream width and consider the
sampling point to be "Center". If a sampling point is located in water that is too deep or unsafe
to wade, select an alternate sampling point on the transect at random.
The procedure for collecting samples at each transect is described in Table 8.2. At each sampling
point, determine if the habitat is a "riffle/run" or a "pool/glide" (any area where there is not
sufficient current to extend the net is operationally defined as a pool/glide habitat). Record the
dominant substrate type (fine/sand, gravel, coarse substrate (coarse gravel or larger) or other
(e.g., bedrock, hardpan, wood, aquatic vegetation, etc.) and the habitat type (pool, glide, riffle,
or rapid) for each sample collected on the Benthic Collection Form in the NRSA App. As you
proceed upstream from transect to transect, combine all samples into a bucket.
Thoroughly rinse net and proceed
upstream to the next transect and ne
sampling location in the pattern L, C,
I
Composite the samples from all
transects to create a single sample fc
the site
Figure 8.1 Benthic Macroinvertebrate Collection at Wadeable Sites
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Combine ALL kick net samples collected from ALL transects
I
TRANSECT SAMPLES (1 per transect)
Sampling point at each transect selected systematically after random start
Sampling points proceed in L, C, R pattern upstream
Modified D-frame kick net
1 square foot quadrat sampled for 30 seconds
I
COMPOSITE SAMPLES FROM ALL TRANSECTS
• Sieve bucket or other bucket(s)
SIEVE SAMPLE
• 500 jjm sieve bucket
• Remove and wash large objects
COMPOSITE AND PRESERVE SAMPLE
1 liter bottle(s) (max of 4 bottles if possible)
Fill no more than 50% with sample
Preserve with ~95% ethanol for a final con-
centration of at least 70%
1 L
Figure 8.2 Transect Sample
Design for Collecting Benthic Macroinvertebrates at Wadeable Sites
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Table 8.2 Procedure: Benthic Macroinvertebrates (Wadeable Sites)
3. With the net opening facing upstream, quickly position the net securely on the stream bottom to
eliminate gaps under the frame. Avoid large rocks that prevent the net from seating properly on the
stream bottom.
NOTE: If there is too little water to collect the sample with the D-net, randomly pick up 10 rocks from the
riffle and pick and wash the organisms off them into a bucket which is half full of water.
4. Holding the net in position on the substrate, visually define a quadrat that is one net width wide and
long upstream of the net opening. The area within this quadrat is 1 ft2.
5. Check the quadrat for heavy organisms, such as mussels and snails. Remove these organisms by hand
and place them into the net. Pick up loose rocks or other larger substrate particles in the quadrat.
Use your hands to dislodge organisms and wash them into the net. Scrub all rocks that are golf ball
sized or larger and which are at least halfway into the quadrat. After scrubbing, place the substrate
particles outside of the quadrat.
6. Hold the D-net securely in position. Starting at the upstream end of the quadrat, vigorously kick the
remaining finer substrate within the quadrat for 30 seconds (use a stopwatch).
NOTE: For samples located within dense beds of long, filamentous aquatic vegetation (e.g., algae or
moss), kicking within the quadrat may not be sufficient to dislodge organisms in the vegetation. Usually
these types of vegetation are lying flat against the substrate due to current. Use a knife or scissors to
remove only the vegetation that lies within the quadrat (i.e., not entire strands that are rooted within
the quadrat) and place it into the net.
7. Pull the net up out of the water. Immerse the net in the stream several times to remove fine
sediments and to concentrate organisms at the end of the net. Avoid having any water or material
enter the mouth of the net during this operation.
8. Go to Step 13.
Pool/Glide Habitats:
9. Visually define a quadrat that is one net width wide and long at the sampling point. The area within
this quadrat is 1 ft2.
10. Check the quadrat for heavy organisms, such as mussels and snails. Remove these organisms by hand
and place them into the net. Pick up loose rocks or other larger substrate particles in the quadrat.
Use your hands to dislodge organisms and wash them into the net. Scrub all rocks that are golf ball
sized or larger and which are at least halfway into the quadrat. After scrubbing, place the substrate
particles outside of the quadrat.
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11. Vigorously kick the remaining finer substrate within the quadrat with your feet while dragging the net
repeatedly through the disturbed area just above the bottom. Keep moving the net all the time so
that the organisms trapped in the net will not escape. Continue kicking the substrate and moving the
net for 30 seconds.
NOTE: If there is too little water to use the kick net, stir up the substrate with your gloved hands and use
a sieve with 500 ixm mesh size to collect the organisms from the water in the same way the net is used
in larger pools.
12. After 30 seconds, remove the net from the water with a quick upstream motion to wash the
organisms to the bottom of the net.
All samples:
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13. Invert the net into a sieve bucket and transfer the sample. Remove as much gravel as possible so that
the organisms do not get damaged. Inspect the net for any residual organisms clinging to the net and
deposit them into the bucket. Use forceps if necessary to remove organisms from the net. Carefully
inspect any large objects (such as rocks, sticks, and leaves) in the bucket and wash any organisms
found off of the objects and into the bucket before discarding the object. Remove as much detritus as
possible without losing organisms.
NOTE: It is recommended that crews carry a sample bottle containing a small amount ofethanol with
them to enable them to immediately preserve larger predaceous invertebrates such as hellgrammites
and water beetles. Doing so will help reduce the chance that other specimens will be consumed or
damaged prior to the end of the field day.
14. Determine the predominant substrate size/type you sampled within the sampling quadrat. Mark the
sampled substrate type on the Benthic Collection Form under the pertinent heading and transect
row. The substrate types are:
• Fine/sand (F): not gritty (silt/clay/muck <0.06 mm diam.) to gritty, up to ladybug sized (2 mm)
• Gravel (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
comment bubbles on field form.
15. Identify the channel habitat type where the sampling quadrat was located. Indicate the channel
habitat type on the Benthic Collection Form under the pertinent heading and transect row. 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.
16. If any transects could not be sampled for benthos, leave the substrate and channel bubbles empty for
the transect and use the adjacent comment bubble to explain why the sample could not be collected
at the transect.
Thoroughly rinse the net before proceeding to the next sampling station. Proceed upstream to the next
transect (through Transect K, the upstream end of the reach) and repeat steps 1 -15. Assign the sampling
station systematically after the first random selection following the repeating pattern L, C, R as you move
upstream. Combine all kick net samples from riffle/run and pool/glide habitats into the bucket.
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Collecting Benthic Macroinvertebrate Sample
17. Record the number of transects that were sampled throughout the reach on the Benthic Collection
Form in the App.
8.4 Sample Processing in Field
Use a 500 |a,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 L jar and preserve with 95% ethanol. Do
not fill jars more than Vz full of material. Multiple jars may be required if detritus is heavy (Table
8.3). If there is a large amount of organic material in the sample, or there are adverse field
conditions (i.e., hot, humid weather), place sample in a 1 L jar with ethanol after each station.
Try to use no more than four 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 (as well as other pertinent sample
information) is 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 Benthic Collection Form in the NRSA App.
Place the samples in a cooler or other secure container for transporting and/or shipping to the
laboratory (see Appendix C).
Table 8.3 Procedure: Compositing Samples for Benthic Macroinvertebrates (Wadeable Sites)
Compositing Benthic Macroinvertebrate Sample
1. Pour the entire contents of the bucket into a sieve bucket with 500 nm 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 four jars per site.
4. Fill in a sample label with the Site ID, date of collection, visit, and jar number (i.e., Jar 1 of 2). Attach
the completed label to the jar and cover it with a strip of clear tape. Verify the sample ID number for
the composite sample matches the ID on the Tracking Form in the App.
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 K
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 Benthic Collection Form. Make sure the number you record matches the actual
number of jars used. Write "Jar N ofX" on each sample label using a waterproof marker ("N" is the
individual jar number, and "X" is the total number of jars for the sample).
6. Place a waterproof label inside each jar with the following information written with a number 2 lead
pencil:
Site ID Collectors initials
Type of sampler Number of stations sampled
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Compositing Benthic Macroinvertebrate Sample
Name of site
Date of collection Jar "N" of "X"
7. Completely fill the jar with 95% ethanol (no headspace). It is very important that sufficient ethanol
be used, or the organisms will not be properly preserved. Existing water in the jar should not dilute
the concentration of ethanol below 70%. Once preserved, mark the "Preserved" bubble on the data
form in the App.
NOTE: Composite samples can be transported back to the vehicle before adding ethanol if necessary. In this
case, fill the jar with stream water, which is then drained using the net (or sieve) across the opening to
prevent loss of organisms, and replace with ethanol.
8. Replace the cap on each jar. Slowly tip the jar to a horizontal position, then gently rotate the jar to
mix the preservative. Do not invert or shake the jar. After mixing, seal each jar with plastic tape.
9. Store labeled composite samples in a container with absorbent material that is suitable for use with
70% ethanol until transport or shipment to the laboratory.
10. If no benthic sample were collected, select the "No Sample Collected" box on the data form and
indicate the reason why targeted samples were not collected in the adjacent comment bubble.
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9 PERIPHYTON
9.1 Summary of Method
Collect periphyton from the 11 cross-section transects ("A" through "K") established within the
sampling reach. Collect periphyton samples at the same transect location (L, C, or R) as the
benthic macroinvertebrate samples (Section 8) directly after collecting the benthic
macroinvertebrate sample. Prepare one composite sample of periphyton for each reach. At the
completion of the day's sampling activities, but before leaving the site, prepare four types of
laboratory samples (an ID/enumeration sample to determine taxonomic composition and
relative abundances, a metagenomic sample, a chlorophyll-o sample, and a biomass sample (for
ash-free dry mass [AFDM])) from the composite periphyton sample.
9.2 Equipment and Supplies
Table 9.1 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 site.
Table 9.1 Equipment and Supplies: Periphyton (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 Dl water
500 mL graduated plastic bottle for the composite sample
60 mL plastic syringe with tip removed, and length of tubing (20 mL)
Timer or stopwatch
Cooler (small soft-sided preferred)
Wet ice
Field Operations Manual
For recording measurements
Sample Collection Form in NRSA App
Fine-tipped indelible markers for filling out sample labels
Sample labels (4 per site) with pre-printed Sample ID numbers
Clear tape strips for covering labels
For cleaning equipment
10% Bleach solution
9.3 Sampling Procedure
At each of the 11 transects, collect samples from the sampling station assigned during the layout
of the reach (L, C, or R). Collect the substrate selected for sampling from a depth no deeper than
0.5 m. If a sample cannot be collected because the location is too deep, pick another point along
the transect as close to the planned location as possible. The procedure for collecting samples
and preparing a composite sample is presented in Table 9.2. Collect one sample from each of
the transects and composite into one bottle to produce one composite sample for each site.
Record number of transects sampled and the total the volume of the composite sample on the
Sample Collection Form in the NRSA App.
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Table 9.2 Procedure: Collecting Composite Index Samples of Periphyton (Wadeable Sites)
Periphyton Composite Sample
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1. Starting with Transect "A", collect a single sample from the assigned sampling station using the
procedure below.
a) If coarse substrate (cobbles, woody materials, etc.) are present that can be removed from the
water:
i) 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 or over a plastic funnel which drains
into a 500 mL plastic bottle with volume graduations marked on it.
ii) 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.
iii) Fill a wash bottle with Dl 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.
iv) Put the bottle in a cooler on ice while you travel between transects and collect the subsequent
samples. (The sample needs to be kept cool and dark because a chlorophyll sample will be
filtered from the composite).
b) If large coarse substrate is present that is too large to remove from the water (bedrock, large
woody materials, boulders, etc.):
v) 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 clear tube attached to
the tip of the syringe in a scraping motion.
vi) While dislodging periphyton with the tube, simultaneously pull back to 25 mL on the syringe
plunger to draw the dislodged periphyton into the syringe. The 25 mL in the syringe combined
with the 20 mL in the tube equals the target volume of 45 mL.
vii) Empty the syringe and tube 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 Dl water to the bottle to raise the level to the next
graduation.
viii) Put the bottle in a cooler on ice while you travel between transects and collect the subsequent
samples. (The sample needs to be kept cool and dark because a chlorophyll sample will be
filtered from the composite.)
c) If no coarse sediments (cobbles or larger) are present:
ix) Use the area delimiter to confine a 12 cm2 area of soft sediments.
x) Vacuum the top 1 cm of sediments from within the delimited area into a de-tipped 60 mL
syringe with attached clear tube up to the 25 mL line of the syringe.
xi) 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 Dl water to the bottle to raise the level to the next graduation.
xii) Put the bottle in a cooler on ice while you travel between transects and collect the subsequent
samples. (The sample needs to be kept cool and dark because a chlorophyll sample will be
filtered from the composite.)
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2. Repeat Step 1 for transects "B" through "K". Place the sample collected at each sampling station into
the single 500 mL bottle to produce the composite index sample.
Storage
3. After samples have been collected from all 11 transects (or as many transects as possible), thoroughly
mix the 500 mL bottle regardless of substrate type. Record the total volume of the composite sample
in the periphyton section of the Sample Collection Form. The graduations on the collection bottle are
placed at 45 mL increments, so the total volume of the composite sample should be the number of
transects sampled times 45.
4. Record the number of transects sampled. If all 11 transects were not sampled, record the reason(s) for
any missed transects in the comment bubble adjacent to the "Number of Transects" field.
5. If sample(s) are not collected at all, select the "No Sample Collected" box on the data form in the NRSA
App and indicate the reason why targeted sample(s) were not collected in the adjacent comment
bubble(s).
Clean up
6. After preparing the four types of laboratory samples (see Section 14.3.6), thoroughly clean each of the
pieces of periphyton equipment (delimiter, brush, funnel, syringe, and composite bottle) with a 10%
Bleach solution and rinse with tap or Dl water.
9.4 Sample Processing in the Field
You will prepare four different types of laboratory samples from the composite sample: an
ID/enumeration sample (to determine taxonomic composition and relative abundances), a
metagenomic sample, a chlorophyll-o sample, and a biomass sample (for AFDM). All of the
methods for processing the four samples are found in the Final Site Activities (Section 14)
portion 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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10 PHYSICAL HABITAT CHARACTERIZATION
Field measurements for physical habitat are made at two scales of resolution along the mid-
channel length of the reach, and the results are later aggregated and expressed for the entire
reach. The protocol defines the length of each sampling reach proportional to stream channel
wetted width and then systematically places measurements to statistically represent the entire
reach.
10.1 Equipment and Supplies
Table 10.1 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
stream. 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 10.1 Equipment and Supplies: Physical Habitat
For making
Convex spherical canopy densiometer (Lemmon Model B), modified with taped "V"
measurements
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 (optional)
50 m or 100 m measuring tape with reel
Meter stick for bank angle measurements
Laser rangefinder (400 ft. distance range) and clear waterproof bag
Clinometer
Binoculars (optional)
Bearing compass
Surveyor's telescoping leveling rod
Sounding rod or wading staff
Level tripod
CST Berger SAL 20 Automatic Level
Field Operations Manual
For recording
In the NRSA App:
data
Physical Habitat Form
Slope Form
Channel Constraint Form
Torrent Evidence Form
Site Assessment Form
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10.2 Summary of Methods Approach
Physical habitat in streams includes all those physical attributes that influence or provide
sustenance to organisms within the stream. The physical habitat of a stream varies naturally,
thus expectations differ even in the absence of anthropogenic disturbance. The procedures are
employed on a reach length 40 times its mean wetted width at the time of sampling.
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Measurement points are systematically placed to statistically represent the entire reach. Stream
depth and wetted width are measured at very tightly spaced intervals, whereas channel cross-
section profiles, substrate, bank characteristics and riparian vegetation structure are measured
at larger intervals. Woody debris is tallied along the full length of the sampling reach. 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.
10.3 Components of the Habitat Characterization
There are six components of the physical habitat characterization (Table 10.2). Physical habitat
measurements are recorded using the NARS App. Similarly, slope and bearing measurements,
the degree of channel constraint, and evidence of debris torrents or recent major flooding are
recorded in the NRSA App.
Table 10.2 Summary of Components of Physical Habitat Characterization at Wadeable Sites
Component Description
Channel and
Riparian
Characterization
Wetted Width /
Bar Width
Thalweg Profile
Woody Debris
Tally
Assessment of
Channel
Constraint, Debris
Torrents, and
Major Floods
At 11 transects placed at equal intervals along reach:
• Measure channel cross-section dimensions, bank height, bank undercut
distance, bank angle, slope and compass bearing (backsight), and riparian
canopy density (with densiometer).
• Visually estimate3: substrate size class, embeddedness and water depth at
five equidistant points on cross-section; areal cover class and type (e.g.,
woody trees) of riparian vegetation in canopy, understory, and ground
cover; areal cover class offish concealment features, aquatic macrophytes
and filamentous algae.
• Observe and record3: presence and proximity of human disturbances.
At 10 cross-sections that are midway between the 11 transects above:
Visually estimate3 substrate size class at 5 equidistant points on each cross-
section
• Measure wetted width and bar width (if present) and evaluate substrate
particle size classes at 11 cross-section transects and midway between them
(21 width measurements and substrate notations along entire reach).
Measure maximum depth, classify habitat and check presence of backwaters,
side channels and loose, soft deposits of sediment particles at 10-15 equally
spaced intervals between each of 11 transects (100 or 150 individual
measurements along entire reach). The number of thalweg measurements is
specified by the stream's mean wetted width.
Between each of the channel cross-sections, tally large woody debris numbers
within and above the bankfull channel according to specified length and
diameter classes (10 separate tallies).
After completing thalweg and transect measurements and observations,
identify features causing channel constraint, estimate the percentage of the
channel margin that is constrained for the whole reach, and estimate the
bankfull and valley widths. Check for evidence of recent major floods and
debris torrent scour or deposition.
a Substrate size class is estimated for a total of 105 particles taken at 5 equally spaced points along each
of 21 cross-sections. Depth is measured and embeddedness estimated for the 55 particles located along
the 11 regular transects A through K. Cross-sections are defined by laying the surveyor's rod or tape to
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span the wetted channel. Woody debris is tallied over the distance between each cross-section and the
next cross-section upstream. Riparian vegetation and human disturbances are observed 5m upstream
and 5m downstream from the cross-section transect. They extend shoreward 10m from left and right
banks. Fish cover types, aquatic macrophytes, and algae are observed within the channel 5m upstream
and 5m downstream from the cross-section stations. These boundaries for visual observations are
estimated by eye.
10.4 Work Flow for the Physical Habitat Components
The six components (Table 10.2) of the habitat characterization are organized into four grouped
activities described below in the following sections.
10.4.1 Channel/Riparian Cross-Sections
At each of the 11 transects, one person proceeds with the channel cross-section dimension
measurements and substrate observations as described below; and also makes measurements
of bank angle (using rod and clinometer) and canopy cover (using densiometer). The second
person records those measurements in the specific sections of the Physical Habitat Form in the
NRSA App while making visual estimates of riparian vegetation structure, instream fish cover,
and human disturbance specified on that form.
Slope is measured by measuring the difference in elevation between each transect and bearing
is determined by backsighting to the previous transect. Supplementary points may need to be
located and flagged (using a different color) if the stream is extremely brushy, sinuous, or steep
to the point that you cannot sight for slope and bearing measures between two adjacent
transects.
The work flow for the thalweg profile and channel cross-section described above can be
modified by delaying the measurements for slope and bearing and/or the woody debris tally
until after reaching the upstream end of the reach. Backsighting and/or wood tallies can be
done on the way back downstream (note that in this case, the slope and bearing data form
would have to be completed in reverse order). Crews may also elect to return to Transect A and
record slope and bearing measurements on a second trip upstream through the reach.
10.4.2 Thalweg Profile and Large Woody Debris Tally
"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. Thalweg
spacing is calculated such that either 10 or 15 evenly spaced measurements are made between
each transect (see Section 10.5.2). Two people proceed upstream from the downstream end of
the sampling reach making observations and measurements at the calculated increment
spacing. One person is in the channel making width and depth measurements and determining
whether soft/small sediment deposits are present under his/her wading staff. The other person
records these measurements, classifies the channel habitat, records presence/absence of side
channels and off-channel habitats (e.g., backwater pools, sloughs, alcoves), and tallies large
woody debris. Each time the crew reaches a flag marking a new cross-section transect, they
start filling out a new portion of the Physical Habitat Form in the NRSA App by selecting the
appropriate Transect bubble at the top of the form. They interrupt the thalweg profile and
woody debris tallying activities to complete data collection at each cross-section transect as
they come to it. When the crew member in the water makes a width measurement at channel
locations midway between regular transects (i.e. at the fifth or seventh thalweg measurement in
each sub-reach), she or he also locates and estimates the size class of the substrate particles on
the left channel margin (0%) and at positions 25%, 50%, 75%, and 100% of the distance across
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the wetted channel. Procedures for this substrate tally are the same as for those at regular
cross-sections, but data are recorded in a different portion- of the field form.
10.4.3 Channel Constraint and Torrent Evidence
After completing observations and measurements along the thalweg and at all 11 transects, the
Field Crew completes the overall reach assessments of channel constraint and evidence of
debris torrents and major floods.
10.5 Habitat Sampling within the Reach
Measurements are made at two scales along the length of the reach; the results are later
aggregated for the entire reach using procedures described by Kaufmann et al. (1999). Figure
10.1 illustrates the locations within the reach where data for the different components of the
physical habitat characterization are collected. Most 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). Thalweg profile measurements will be
spaced evenly over the entire reach. In addition, they must be sufficiently close together that
they do not miss deep areas and major habitat units.
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~, Channel/Riparian
' Cross-section
^^^Transect
• V
Thalweg
profile
stations
% P Vi
....I..--***
Intermediate transects (width and
substrate measurements only
) Woody
^ Debris
-? Tally
B J (between
transects)
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of sampling reach ^
PRK/DVP 8ffl6
Figure 10.1 Reach Layout for Physical Habitat Measurements for streams 2.5m or greater (plan view)
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10.5.1 Channel and Riparian Measurements at Cross-Section Transects
10.5.1.1 Substrate Size an d Charm el Dim ensions
Substrate size and embeddedness are evaluated at 5 points at each of the 11 transects (refer to
Figure 10.2). Substrate size (but not embeddedness) is also evaluated at 10 additional cross-
sections located midway between each of the 11 regular transects (A-K) during the thalweg
profile. In the process of measuring substrate particle sizes at each transect, the water depth at
each substrate sample point is measured (at the 10 midway cross-sections, depth to the
substrate point is not recorded). If the wetted channel is split by a mid-channel bar (Section
10.5.2), the five substrate points are centered between the wetted width boundaries regardless
of the mid-channel bar in between. Consequently, substrate particles selected in some
cross-sections may be "high and dry." For cross-sections that are entirely dry, make
measurements across the unvegetated portion of the channel.
PRK/DVP 8/06
Figure 10.2 Substrate Sampling Cross-Section ^
O
I—
The substrate sampling points along the cross-section are located at 0, 25, 50, 75, and 100 m
percent of the measured wetted width, with the first and last points located at the water's edge 2j
just within the left and right banks. The procedure for obtaining substrate measurements is ^
described in Table 10.3 (including all particle size classifications). Record these measurements in ^
the Substrate Cross-sectional Information section of the Physical Habitat Form in the NRSA App, J
I—
To minimize bias in selecting a substrate particle for size classification, it is important to
concentrate on correct placement of the measuring stick along the cross-section, and to select "3
the particle right at the bottom of the stick (not, for example, a more noticeable large particle ^
that is just to the side of the stick). Classify the particle into one of the size classes listed on the <
data form based on the middle dimension of its length, width, and depth. This median sjj
dimension determines the sieve size through which the particle can pass. When you record the J
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size class as Other, describe the substrate type in the adjacent comment bubble on the field
form.
At substrate sampling locations on the 11 regular transects (A-K), examine particles larger than
sand for surface stains, markings, and algal coatings to estimate embeddedness. Embeddedness
is the fraction of a particle's volume that is surrounded by (embedded in) sand or finer
sediments on the stream bottom. For particles 16mm diameter and larger, (e.g., coarse gravel
[GC] and above), estimate the embeddedness of the volume of the entire individual particle at
the point of the wading rod. For fine gravel (GF), which are particles between 2 and 16mm
diameter, estimated the average embeddedness of all particles within a 10 cm diameter circle
around the substrate sampling point. By definition, the embeddedness of sand and fines (silt,
clay, and muck) is 100 percent, and the embeddedness of hardpan and bedrock is zero percent.
When these size classes are selected in the App, the correct embeddedness value will
automatically be entered.
Table 10.3 Procedure: Substrate Measurement
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1. Select the appropriate cross-section transect at the top pf the Physical Habitat Form in the NRSA
App. At the transect, extend the surveyor's rod or metric tape across the channel perpendicular to
the flow, with the "zero" end at the left bank (facing downstream).
NOTE: If a side channel is present, and contains 16 - 49% of the total flow, establish a secondary
cross-section transect. Mark the "IS THIS A SIDE CHANNEL" checkbox near the top of the form to
record data for the side channel. Unchecking the side channel box will display the primary transect
data once again. Collect all channel and riparian cross-section measurements from the side channel
as well as the primary channel. While collecting thalweg profile data (including mid-station substrate
size class) and large woody debris tallies, record the cumulative data on the primary transect only
(e.g., with the side channel check box unchecked).
2. Enter 0 as the left bank starting distance on the top row data form and record the distance to the
right bank (e.g., the wetted with at the transect) to the nearest 0.1 meters on the fifth row. Doing so
will prompt the App to divide the wetted channel width by 4 to locate substrate measurement
points on the cross-section. The App will enter these values in the three remaining fields of the
form. These calculated distances correspond 25% (LeftCenter), 50% (Center), 75% (RightCenter).
3. Place your sharp ended meter stick or calibrated pole at the Left location (0 m). Measure the depth
and record it on the field data form.
• Depth entries at the left and right banks may be 0 (zero) if the banks are gradual.
• If the bank is nearly vertical, let the base of the measuring stick fall to the bottom (i.e., the
depth at the bank will be > 0 cm), rather than holding it suspended at the water surface.
4. Pick up the substrate particle that is at the base of the meter stick (unless it is bedrock or boulder),
and visually estimate its particle size, according to the following table. Classify the particle according
to its median diameter (the middle dimension of its length, width, and depth). Record the size class
code on the data form by tapping the blue data field and selecting the correct size class.
Code
Size Class
Size Range (mm)
Description
RS
Bedrock (Smooth)
>4000
Smooth surface rock bigger than a car
RR
Bedrock (Rough)
>4000
Rough surface rock bigger than a car
RC
Concrete/Asphalt
Regardless of Size
Artificial materials
XB
Large Boulders
>1000 to 4000
Yard/meter stick to car size
SB
Small Boulders
>250 to 1000
Basketball to yard/meter stick size
CB
Cobbles
>64 to 250
Tennis ball to basketball size
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GC
Gravel (Coarse)
>16 to 64
Marble to tennis ball size
GF
Gravel (Fine)
> 2 to 16
Ladybug to marble size
SA
Sand
>0.06 to 2
Smaller than ladybug; gritty between fingers
FN
Fines
<0.06
Silt Clay Muck (not gritty between fingers)
HP
Hardpan
>4000
Firm, consolidated fine substrate
WD
Wood
Regardless of Size
Wood & other organic particles
OT
Other
Regardless of Size
Concrete, metal, tires, car bodies, etc.
(describe in comment bubble)
5.
Evaluate substrate embeddedness as follows at each of the five points. For particles larger than
sand, examine the surface for stains, markings, and algae. For particles 16mm diameter and
larger, (e.g., coarse gravel [GC] and above), estimate the embeddedness of the volume of the
entire individual particle at the point of the wading rod. For fine gravel (GF), which are particles
between 2 and 16mm diameter, estimate the average embeddedness of all particles within a 10
cm diameter circle around the substrate sampling point. By definition, sand and fines are
embedded 100%; bedrock and hardpan are embedded 0%. These values will automatically
populate the form. For sand and smaller particles, you will not be able to pick up an individual
particle, but a "pinch" of fine particles between your fingers. Determine and record the dominant
size of particles in the "pinch." If a fine flocculent coating coats the particle at the end of the
wading rod, we are interested in the diameter of the particle, not the flocculent coating of fine
material -— unless the coating is so thick that it obscures the larger particles. For example, If the
tip of the wading rod falls on a cobble, and there is a thin layer of fine sediment that can be
scoured away by waving your hand over the particle or by picking it up, note the presence of
these fine coatings in comments, but record the diameter and embeddedness of the larger
particle.
6.
Move to the next location on the transect, and repeat Steps 3 - 5 at each location. Repeat Steps 1 - 5
at each transect, including any additional side channel transects established if side channels are
present.
10.5.1.2 Instream Fish Cover, Algae, and Aquatic Macrophytes
Over a defined area upstream and downstream of the sampling transects (Figure 10.3), crews
shall estimate by eye and/or 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 instream fish cover is outlined in Table g
10.4. Data are recorded in the Fish Cover section of the Physical Habitat Form in the NRSA App. F
Estimate the areal cover of all of the fish cover and other listed features that are in the water n
and on the banks 5 m upstream and downstream of the cross-section (Figure 10.3). The areal ljj
cover classes of fish concealment and other features are the same as those described for ^
riparian vegetation (Section 10.5.1.5). <
The entry Filamentous algae refers to long streaming algae that often occur in slow moving ^
waters. Macrophytes are water loving plants, including mosses, in the stream that could provide
cover for fish or macroinvertebrates. If the stream channel contains live wetland grasses, ^
include these as aquatic macrophytes. WOODY DEBRIS are the larger pieces of wood that can ^
provide cover and influence stream morphology (i.e., those pieces that would be included in the ^3
large woody debris tally [Section 10.5.2]). Brush/woody debris refers to smaller wood pieces
that primarily affect cover but not morphology. Live Trees or Roots are living trees that are J
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within the channel - estimate the areal cover provided by the parts of these trees or roots that
are inundated. Overhanging vegetation includes tree branches, brush, twigs, or other small
debris that is not in the water but is close to the stream (within 1 m of the surface) and provides
potential cover. For ephemeral channels, estimate the proportional cover of these trees that is
inundated during bankfull flows. Boulders are typically basketball to car-sized particles.
Artificial structures include those designed for fish habitat enhancement, as well as in-channel
structures that have been discarded (e.g., concrete, asphalt, cars, or tires) or deliberately placed
for diversion, impoundment, channel stabilization, or other purposes.
10 m
10 m
RIPARIAN
PLOT
(Left Bank)
Instream Fish
Cover Plot
RIPARIAN
PLOT
(Right Bank)
10 m
PRK/DVP 8/06
Figure 10.3 Riparian Zone and Instream Fish Cover Plots for a Stream Cross-Section Transect
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Table 10.4 Procedure: Estimating Instream Fish Cover
Instream Fish Cover Measurements
1. Standing at a point along the transect where riparian observations can be made effectively, estimate
a 5 m distance upstream and downstream (10 m total width).
2. Examine the water and both banks within the 10 m segment of stream for the following features
and types offish cover '.filamentous algae, macrophytes, large woody debris, brush and small woody
debris, in-channel live trees or roots, overhanging vegetation, undercut banks, boulders, and artificial
structures.
3. For each cover type, estimate the areal cover. Record the appropriate cover class in the Fish Cover
section of the Physical Habitat Form in the App:
Chabsent: 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 (including any additional side channel
transects established when islands are present).
10.5.1.3 Bank Characteristics
The procedure for obtaining bank and channel dimension measurements is presented in Table
10.5. Data are recorded in the Bank Measurements section of the Physical Habitat Form in the
NRSA App. Bank angle and bank undercut distances are determined on the left and right banks
at each cross-section transect. Figure 10.4 illustrates how bank angle is determined for several
different situations. Measure bank angle on wadeable streams at the scale of approximately 0.5
m using a short (approx. 1 m long) pole. When measuring the angle, try to ensure that at least
half (0.5 m) of the pole length is in contact with the bank. Other features include the wetted
width of the channel, the width of exposed mid-channel bars of gravel or sand, estimated
incision height, and the estimated height and width of the channel at bankfull stage as described
in Figure 10.5. Bankfull height and incised height are both measured relative to the present
water surface (i.e., the level of the wetted edge of the stream). This is done by placing the base
of the small measuring rod at the bankfull elevation and sighting back to the survey rod placed
at the water's edge using the clinometer as a level (i.e., positioned so the slope reading is 0%.).
The height of the clinometer above the base of the smaller rod is subtracted from the elevation
sighted on the surveyor's rod. Bankfull flows are large enough to erode the stream bottom and
banks, but frequent enough (every one to two years) to prevent substantial growth of upland
terrestrial vegetation. Consequently, in many regions, it is these flows that have determined the
width and depth of the channel.
Table 10.5 Procedure: Measuring Bank Characteristics
1. To measure bank angle, lay a meter ruler or a short (approx. 1 m long) rod down against the left
bank (determined as you face downstream), with one end at the water's edge. At least 0.5 m of
the ruler or rod should be resting comfortably on the ground to determine bank angle. If the
ground adjacent to the water's edge is not indicative of the predominant angle of the 1 meter
shoreline, it may be necessary to move the end of the rod away from the water's edge to
correctly measure the predominant angle of the 1 meter shoreline. Lay the clinometer on the rod,
and read the bank angle in degrees from the external scale on the clinometer. Record the angle in
the field for the left bank in the Bank Measurements section of the Physical Habitat Form in the
App.
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• A vertical bank is 90°, overhanging banks have angles >90° approaching 180°, and more
gradually sloped banks have angles <90°. To measure bank angles >90°, turn the clinometer
(which only reads 0 to 90°) over and subtract the angle reading from 180°.
• If there is a large boulder or log present at the transect, measure bank angle at a nearby point
where conditions are more representative.
2. If the bank is undercut, measure the horizontal distance of the undercutting to the nearest 0.01
m. The undercut distance is the distance from the deepest point of the undercut out to the point
where a vertical plumb line from the bank would hit the water's surface. Record the distance on
the field data form. Measure submerged undercuts by thrusting the rod into the undercut and
reading the length of the rod that is hidden by the undercutting. If the bank is not undercut,
record 0 in the in the Undercut Distance field.
3. Repeat Steps 1 and 2 on the right bank.
4. Verify the wetted width value determined when locating substrate sampling points. The Wetted
Width field in the bank measurements section of the form will be populated based on the wetted
width entered earlier and is not editable here. Also determine the bankfull channel width and the
width of exposed midchannel bars (if present). Record these values in the Bank Measurements
section of the form. If no bars exist at the transect, enter 0 in the bar width field.
5. While still holding the surveyor's rod as a guide, and sighting with the clinometer as a level,
examine both banks to measure and record the height of bankfull flow above the present water
level. Look for evidence on one or both banks such as:
• An obvious slope break that differentiates the channel from a relatively flat floodplain terrace
higher than the channel.
• A transition from exposed stream sediments to terrestrial vegetation.
• Moss growth on rocks along the banks.
• Presence of drift material caught on overhanging vegetation.
• A transition from flood and scour tolerant vegetation to that which is relatively intolerant of
these conditions.
6. Hold the surveyor's rod vertical, with its base planted at the water's edge. Examine both banks,
then determine the channel incision as the height up from the water surface to elevation of the
first terrace of the valley floodplain (Note, this is at or above the bankfull channel height).
Whenever possible, use the clinometer as a level (positioned so it reads 0% slope) to measure this
height by transferring (backsighting) it onto the surveyor's rod. Record this value in the Incised
Height field of the bank measurement sections on the form.
7. Repeat Steps 1 through 6 at each cross-section transect, (including any additional side channel
transects established when islands are present).
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bank angle = clinometer reading
,_Abank an9|e = clinorneter reading
A
B
5 V' ~*i \
pole rests most this is as "comfortable"
as pole can get within q
1 m of wetted edge
. ;undercut bank
/ above water' - . [
; ~ surface <0 5m
< : ^ ¦
x , - . - ' bank angle = 180° minus
clinometer reading
; (e.g. 180° - 30° = 150°)
'/1 f p undercut distance
Figure 10.4 Determining Bank Angle Under Different Types of Bank Conditions
(A) typical, (B) incised channel, (C) undercut bank (less than 0.5 m), and (D) overhanging bank (greater
than 0.5 m).
Unfortunately, we have to depend upon evidence visible during the low flow sampling season. If
available, consult published rating curves relating expected bankfull channel dimensions to
stream drainage area within the region of interest. Graphs of these rating curves can help you
get a rough idea of where to look for field evidence to determine the level of bankfull flows.
Curves such as these are available from the USGS for streams in most regions of the U.S. (e.g.,
Dunne and Leopold 1978; Harrelson et al. 1994, Leopold 1994). To use them, you need to know
the contributing drainage area to your sample site. Interpret the expected bankfull levels from
these curves as a height above the streambed in a riffle, but remember that your field
measurement will be a height above the present water surface of the stream. Useful resources
to aid your determination of bankfull flow levels in streams in the United States are video
presentations produced by the USDA Forest Service for western streams (USDA Forest Service
1995) and eastern streams (USDA Forest Service 2002).
After consulting rating curves that show where to expect bankfull levels in a given size of
stream, estimate the bankfull flow level by looking at the following indicators:
First look at the stream and its valley to determine the active floodplain. This is a
depositional surface that frequently is flooded and experiences sediment deposition
under the current climate and hydrological regime.
Then look specifically for:
• An obvious break in the slope of the banks.
• A change from water loving and scour tolerant vegetation to more drought
tolerant vegetation.
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• 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 years 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, provide a comment associated with the height of drift material in the provided
comment bubble on the form.
We use the vertical distance (height) from the observed water surface up to the level of the first
major valley depositional surface (Figure 10.5) 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.
If the channel is not greatly incised, bankfull channel height and incision height will be the same
(i.e., the first valley depositional surface is the active floodplain). However, if the channel is
incised greatly, the bankfull level will be below the level of the first terrace of the valley
floodplain, making bankfull channel height less than incision height (Figure 10.6). Bankfull height
is never greater than incision height. You may need to look for evidence of recent flows (within
about one year) to distinguish bankfull and incision heights. In cases where the channel is
cutting a valley sideslope and has over-steepened and destabilized that slope, the bare
"cutbank" against the steep hillside at the edge of the valley is not necessarily an indication of
recent incision. In such a case, the opposite bank may be lower, with a more obvious terrace
above bankfull height; choose that bank for your measurement of incised height. Examine both
banks to more accurately determine incision height and bankfull height. Remember that incision
height is measured as the vertical distance to the first major depositional surface above bankfull
(whether or not it is an active floodplain or a terrace. If terrace heights differ on left and right
banks (both are above bankfull), choose the lower of the two terraces. In many cases your
sample reach may be in a "V" shaped valley or gorge formed over eons, and the slope of the
channel banks simply extends uphill indefinitely, not reaching a terrace before reaching the top
of a ridge. In such cases, record incision height values equal to bankfull values and make
appropriate comment that no terrace is evident. Similarly, when the stream has extremely
incised into an ancient terrace, (e.g., the Colorado River in the Grand Canyon), you may crudely
estimate the terrace height if it is the first one above bankfull level. If you cannot estimate the
terrace height, make appropriate comments describing the situation.
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A. Channel not Incised
First terrace on
valley bottom
above bankfull
level
No recent incision— bankfull
level at valley bottom
Downcutting over
geologic time
Active
floodplain at or near
valley bottom elevation
(Record this height)
Second
terrace
Valley Fill
B. Incised Channel
Downcutting over
geologic time
Valley Fill
Former active floodplain
no longer connected—
becomes new first terrace
above bankfull level
(Record this height),
Recent incision—
bankfull level below
first terrace of valley
bottom
Former second
terrace becomes
Former first third terrace
terrace becomes
second terrace
Figure 10.5 Schematic Showing Relationship Between Bankfull Channel and Incision
(A) Not recently incised, and (B) recently incised into valley bottom. Note level of bankfull stage relative to
elevation of first terrace (abandoned floodplain) on valley bottom. (Stick figure included for scale).
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A) Deeply Incised Channel
From Figure 7-9 (B)
Incision Height (Always
equal to or greater than
bankfull height)
Second Terrace
\
First Terrace
Fcfmerflist third terrace
ankfull
Height
(When
channel form
is not a good
indicator, use
evidence of
recent
flooding)
B) Small stream constrained in V-shaped valley
No incision:
No evidence of
downcutting,
vertical bank
angle; etc.)
Figure 10.6 Determining Bankfull and Incision Heights
(A) Deeply Incised Channels, and (B) Streams in Deep V Shaped Valleys (Stick figure included for scale).
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10.5.1.4 Canopy Cover Measurements
Canopy cover over the stream is determined at each of the 11 cross-section transects. A
spherical densiometer (model A convex type) is used (Lemmon 1957) and is provided in the
base kit to each crew. Mark the densiometer with a permanent marker or tape exactly as shown
in Figure 10.7 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).
Figure 10.7 Schematic of Modified Convex Spherical Canopy Densiometer.
From Mulvey et al. (1992). In this example, 10 of the 17 intersections show canopy cover, giving a
densiometer reading of 10. Note proper positioning with the bubble leveled and face reflected at the apex
of the "V".
The procedure for obtaining canopy cover data is presented in Table 10.6. 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 10.7. Concentrate on the 17 points of grid
intersection on the densiometer that lie within the taped "V". If the reflection of a tree or high
branch or leaf overlies any of the intersection points, that particular intersection is counted as
having cover. For each of the six measurement points, record the number of intersection points
(maximum=17) that have vegetation or other overhead objects covering them in the Canopy
Cover Measurements section of the Physical Habitat Form in the NRSA App.
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Table 10.6 Procedure: Canopy Cover Measurements
Canopy Cover Measurements with Densiometer
1. At each cross-section transect, stand in the stream at mid-channel and face upstream.
2. Hold the densiometer 0.3 m (1 ft) above the surface of the stream. Level the densiometer using the
bubble level. Move the densiometer in front of you so your face is just below the apex of the taped
"V".
3. Count the number of grid intersection points within the "V" that are covered by either a tree, a leaf,
or a high branch. Record the value (0 to 17) in the Center Up field of the canopy cover
measurements section of the Physical Habitat Form in the App.
4. Face toward the left bank (left as you face downstream). Repeat Steps 2 and 3, recording the value
in the Center Left field of the form.
5. Repeat Steps 2 and 3 facing downstream, and again while facing the right bank (right as you look
downstream). Record the values in the Center Down and Center Right fields of the form.
6. Move to the water's edge (either the left or right bank). Repeat Steps 2 and 3 again, this time facing
the bank. Record the value in the Left or Right fields of the form. Move to the opposite bank and
repeat.
7. Repeat Steps 1 through 6 at each cross-section transect (including any additional side channel
transects established when islands are present).
10.5.1.5 Visual Riparian Estimates
10.5.1.5.1 Riparian Vegetation Structure
The previous section described methods for quantifying the cover of canopy over the stream
channel. The following visual estimation procedures supplement those measurements with a
semi-quantitative evaluation of the type and amount of various types of riparian vegetation.
Riparian vegetation observations apply to the riparian area upstream five meters and
downstream five meters from each of the 11 cross-section transects. They include the visible
area from the stream back a distance of 10 m (~30 ft.) shoreward from both the left and right
banks, creating a 10 m x 10 m riparian plot on each side of the stream (Figure 10.3). The riparian
plot dimensions are estimated, not measured. On steeply sloping channel margins, the 10 m x
10 m plot boundaries are defined as if they were projected down from an aerial view. Table 10.7
presents the procedure for characterizing riparian vegetation structure and composition.
Riparian estimates are recorded in the Visual Riparian Estimates section of the Physical Habitat
Form. Conceptually divide the riparian vegetation into 3 layers: the Canopy layer (> 5 m high),
the Understory layer (0.5 to 5 m high), and the Ground cover layer (< 0.5 m high). Note that
several vegetation types (e.g., grasses or woody shrubs) can potentially occur in more than one
layer. Similarly note that some things other than vegetation are possible entries for the Ground
cover layer (e.g., barren ground).
Before estimating the areal coverage of the vegetation layers, record the type of woody
vegetation (broadleaf Deciduous, Coniferous, broadieaf 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 canopy or understory layer, record the type as None.
Estimate the areal cover separately in each of the three vegetation layers. Note that the areal
cover can be thought of as the amount of shadow cast by a particular layer alone when the sun
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is directly overhead. The maximum cover in each layer is 100%, so the sum of the areal covers for
the combined three layers could add up to 300%. The four areal cover classes are Absent (0%),
Sparse (<10%), Moderate (10 to 40%), Heavy (40 to 75%), and Very Heavy (>75%). These cover
classes and their corresponding codes are shown on the form for reference. When rating
vegetation cover types for a single vegetation layer, mixtures of two or more subdominant
classes might all be given Sparse (1), Moderate (2), or Heavy (3) ratings. One Very Heavy cover
class with no clear subdominant class might be rated 4 with all the remaining classes rated as
either Moderate (2), Sparse (1) or Absent (0). Note that within a given vegetation layer, two
cover types with 40-75% cover can both be rated 3, but no more than one cover type could
receive a rating of 4.
Table 10.7 Procedure: Characterizing Riparian Vegetation Structure
Riparian Vegetative Structure Measurements
1. Standing at a point along the transect where riparian observations can be made effectively, estimate
a 5 m distance upstream and downstream (10 m total length).
2. Facing the left bank (left as you face downstream), estimate a distance of 10 m back into the
riparian vegetation. On steeply sloping channel margins, estimate the distance into the riparian zone
as if it were projected down from an aerial view.
3. Within this 10 m x 10 m area, conceptually divide the riparian vegetation into 3 layers: a Canopy
Layer (> 5 m high), an Understory (0.5 to 5 m high), and a Ground Cover layer (<0.5 m high).
4. Within this 10 m x 10 m area, determine the dominant woody vegetation type for the CANOPY
LAYER (vegetation >5 m high) as either Deciduous, Coniferous, broadleaf Evergreen, Mixed, or None.
Consider the layer Mixed if more than 10% of the areal coverage is made up of the alternate
vegetation type. If the canopy layer contains no vegetation or 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 Physical Habitat Form.
5. Determine separately the areal cover class of large trees (>0.3 m [1 ft] diameter at breast height
[dbh]) and small trees (<0.3 m dbh) within the canopy layer. Estimate areal cover as the amount of
shadow that would be cast by a particular layer alone if the sun were directly overhead. Record the
appropriate cover class on the form (Chabsent: zero cover, l=sparse: <10%, 2=moderate\ 10-40%,
3=heavy: 40-75%, or 4=very heavy: >75%).
6. Look at the UNDERSTORY layer (vegetation between 0.5 and 5 m high). Determine the dominant
woody vegetation type for the understory layer as described in Step 4 for the canopy layer. If the
understory layer contains no vegetation or the dominant vegetation type in the understory is not
woody (e.g., herbaceous), record the vegetation type as "None".
7. Determine the areal cover class for woody shrubs and saplings (as well as trunks and branches of
trees) separately from non-woody vegetation within the understory, as described in Step 5 for the
canopy layer.
8. Look at the GROUND COVER layer (vegetation <0.5 m high). Determine the areal cover class for
woody shrubs and seedlings (including trunks and branches of trees), non-woody vegetation, and
the amount of bare ground or duff (dead organic material present as described in Step 5 for the
canopy layer. Include all non-vegetative materials such as buildings, pavement, standing water, etc.
In the "barren, bare dirt, or duff category.
9. Repeat Steps 1 through 8 for the right bank.
10. Repeat Steps 1 through 9 for all cross-section transects (including any additional side channel
transects established when islands are present).
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10.5.1.5.2 Human Influence
For the left and right banks at each of the 11 detailed Channel and Riparian Cross-sections,
evaluate the presence/absence and the proximity of 11 categories of human influences with the
procedure outlined in Table 10.8. Record data in the Human Influence section of the Physical
Habitat Form in the App. Relate your observations and proximity evaluations to the stream and
riparian area within 5 m upstream and 5 m downstream from the station (Figure 10.8). Four
proximity classes are used: In the stream or on the bank within 5 m upstream or downstream of
the cross-section transect (B), contained within the 10 m x 10 m riparian plot but not in the
stream or on the bank (C), present outside of the riparian plot (P), and absent (0). If a
disturbance is within more than one proximity class, record the one that is closest to the stream
(e.g., present in riparian plot "C" takes precedence over outside of riparian plot "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 Transects 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
another transect or its 10x10 meter riparian plot.
C—within riparian plot
R_nn KanU nr in ctroim
P—outside plot
(but do not sight through next
/ transect or pfot)
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Figure 10.8 Proximity Classes for Human Influences in Wadeable Streams
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Table 10.8 Procedure: Estimating Human Influence
Estimate Human Influence
Standing at a point along the transect where riparian observations can be made effectively, look
toward the left bank (left when facing downstream), and estimate a 5 m distance upstream and
downstream (10 m total width). 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 stream segment for the
following human influences: (1) walls, dikes, revetments, riprap, and dams; (2) buildings; (3)
pavement/cleared lots (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, hay fields, or evidence of livestock; (10) logging; and
(11) mining (including gravel mining). If human influences are observed that do not match one of the
listed categories, choose the category that best describes the influence and add further explanation
in the adjacent comment bubble.
For each type of influence, determine if it is present and what its proximity is to the stream and
riparian plot area. Consider human disturbance items as present if you can see them from the cross-
section transect. Do not include them if you have to sight through another transect or its 10 m x 10
m riparian plot.
For each type of influence, record the appropriate proximity class in the Human Influence part of the
Visual Riparian Estimates section of the Physical Habitat Form in the App. If a disturbance is within
more than one proximity class, record the one that is closest to the stream. Proximity classes (listed
in order of priority) are:
Present within the defined 10 m stream segment and located in the stream or on
the stream bank.
B (Bank)
C (Contained)
P (Present
0 (Absent)
Present within the 10x10 m riparian plot area.
Present, but outside the riparian plot area.
Not present within or adjacent to the 10 m stream segment or the riparian plot
area at the transect
5.
6.
Repeat Steps 1 through 4 for the right bank.
Repeat Steps 1 through 5 for each cross-section transect, (including any additional side channel
transects established when islands are present).
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10.5.2 Thalweg Profile and Large Woody Debris Measurements
10.5.2.1 Thalweg Profile
"Thalweg" refers to the flow path of the deepest water in a stream channel. The thalweg profile
is a longitudinal survey of maximum flow path depth. Data from the thalweg profile allows
calculation of indices of residual pool volume, stream size, channel complexity, and the relative
proportions of habitat types such as riffles and pools. Thalweg profile increments are spaced
evenly throughout the sampling reach to accurately represent the maximum stream depths
throughout the entire reach and must be spaced close enough to provide data on both major
and minor habitat units.
Follow these guidelines for choosing the increment between thalweg profile measurements:
Channel Width < 2.5 m:
• Minimum reach length of 150 m is used
• Thalweg increment = 1.0 m (resulting in 15 thalweg measurements per subreach)
• A total of 150 evenly spaced thalweg profile measurements will be made, 15 between
each channel cross-section
• Mid-subreach measurements are made at the 7th thalweg location
Channel Width 2.5 to 3.5 m:
• Minimum reach length of 150 m is used
• Thalweg increment = 1.5 m (resulting in 10 thalweg, measurements per subreach)
• A total of 100 evenly spaced thalweg profile measurements will be made, 10 between
each channel cross-section
• Mid-subreach measurements are made at the 5th thalweg location
Channel Width > 3.5 m:
• Reach length is 40 times channel width (maximum of 4 km)
• Thalweg increment = 0.01 x reach length (resulting in 10 thalweg measurements per
subreach)
• A total of 100 evenly spaced thalweg profile measurements will be made, 10 between
each channel cross-section
• Mid-subreach measurements are made at the 5th thalweg location
The procedure for obtaining thalweg profile measurements is presented in Table 10.9. Record
data in the Thalweg Profile and Large Woody Debris sections of the Physical Habitat Form in the
NRSA App Use the surveyor's rod and a metric ruler or calibrated rod or pole to make the
required depth and width measurements at each station, and to measure off the distance
between stations as you proceed upstream. You may need to make minor adjustments to align
each 10th measurement to be one increment short of the next transect. In streams with average
widths less than 2.5 m, make thalweg measurements at 1 meter increments. Because the
minimum reach length is set at 150 meters, there will be 15 measurements on a field data form:
Station 0 at the transect plus 14 additional stations between it and the next transect upstream.
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Use the five extra lines on the thalweg profile portion of the data form to record measurements
10-14.
Measure thalweg depths at all stations. Provide a comment for any missing measurements in
the comment bubbles on the data form. At points where a direct depth measurement cannot be
made, make your best estimate of the depth, record it on the form, and comment that it is an
estimated value. Where the thalweg points are too deep for wading, measure the depth by
extending the surveyor's rod or weighted line at an angle to reach the thalweg point. On the
field form enter a comment associated with the thalweg station (to indicate a nonstandard
technique), and calculate the thalweg depth based on the water level on the rod or line, and the
rod/line angle, as determined using the external scale on the clinometer (vertical = 90).
Table 10.9 Procedure: Thalweg Profile
1. Determine the increment distance between measurement stations based on the wetted width used
to determine the length of the reach. Using a laser rangefinder or surveyor's rod:
• For widths < 2.5 m, establish stations every 1 m (150 total).
• For widths > 2.5 and <3.5 m, establish stations every 1.5 m (100 total).
• For widths > 3.5 m, establish stations at increments equal to 0.01 times the reach length (100
total).
2. With Transect A selected on the Physical Habitat Form in the App, complete the Thalweg Profile
header information. Record the increment distance determined in Step 1 in the Increment field and
the total reach length (e.g., the total distance from Transect A to Transect K) in the Total Reach
Length field. It is not necessary to repeat these entries on the remainder of the Physical Habitat
transect forms, the App will populate the remainder of the fields based on the entries in Transect A.
3. Begin at the downstream end (station 0) of the first transect (Transect A).
4. Verify the wetted width at station 0. The Wetted Width field for station 0 will be populated based on
the wetted width entered earlier and is not editable here. You will also measure and record the
wetted width at either station 5 (if the stream width defining the reach length is e 2.5 m), or station
7 (if the stream width defining the reach length is < 2.5 m). Wetted width is measured across and
over mid-channel bars and boulders. Record the width on the field data form to the nearest 0.1 m.
For streams with interrupted flow, where no water is in the channel at the station or transect,
record zeros for wetted width.
NOTE: If a mid-channel bar is present at a station where wetted width is measured, measure the wetted
width across and including the bar, but also measure the bar width and record it on the field data form.
5. At station 5 or 7 (see above) classify the size of the bed surface particle at the tip of your depth
measuring rod at the left wetted margin (0%) and at positions 25%, 50%, 75%, and 100% of the
distance across the wetted width of the stream. This procedure is identical to the substrate size
evaluation procedure described for regular channel cross-sections (Transects A - K), except that for
these midway supplemental cross-sections, substrate size is entered in the Thalweg Profile section
of the Physical Habitat Form in the App.
6. At each thalweg profile station, use a calibrated pole or rod to locate the deepest point within the
deepest flow path (the thalweg), which may not always be found at mid-channel (and may not
always be the absolute deepest point in every channel cross-section). Measure the thalweg depth to
the nearest cm from the substrate surface to the water surface, and record it on the form. Read the
depth on the side of the rod to avoid inaccuracies due to the wave formed by the rod in moving
water.
NOTE: For streams with interrupted flow if there is no water at a transect, record zeros for depth.
NOTE: Obtain thalweg depths at all stations. If the thalweg is too deep to measure directly, stand in
shallower water and extend the surveyor's rod or pole at an angle to reach the thalweg. Determine the
angle by resting the clinometer on the upper surface of the rod and reading the angle on the external scale
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of the clinometer. Calculate the thalweg depth based on your measurements and record a comment in the
provided comment bubble to indicate a non-standard procedure was used. For deeper depths, use the
same procedure with a taut string as the measuring device. Tie a weight to one end of a length of string or
fishing line, and toss the weight into the deepest channel location. Draw the string up tight and measure
the length of the line that is under water. Measure the string angle with the clinometer exactly as done for
the surveyor's rod. If a direct measurement cannot be obtained, make the best estimate you can of the
thalweg depth, and use a comment to identify it as an estimated measurement.
7. At the point where the thalweg depth is determined, observe if unconsolidated, loose (soft)
deposits of small diameter (<16mm) sediments are present directly beneath your ruler, rod, or pole.
Soft/small sediments are defined here as fine gravel, sand, silt, clay or muck readily apparent by
"feeling" the bottom with the rod. Record presence or absence using the checkboxes in the
Soft/Small Sediment column on the form (an unchecked box indicates absence). Note: A thin coating
of fine sediment or silty algae coating the surface of cobbles should not be considered soft/small
sediment. However, fine sediment coatings should be identified in the comments section of the field
form when determining substrate size and type.
8. Determine the channel unit code for the station. Record this on the form by tapping the blue data
field and selecting from the codes provided. For dry and intermittent streams, where no water is in
the channel, record habitat type as dry channel (DR).
9. If the station cross-section intersects a mid-channel bar, indicate the presence of the bar by using
the checkboxes in the Bar Present column on the form (an unchecked box indicates absence).
However, a measurement of the bar width is only taken if the bar intersects a station at either the
endpoint or midpoint of a subreach (e.g., station 0 and station 5 or 7). If this is the case, measure
and record the bar width to the nearest 0.1 meter in the provided data field.
10. Record the presence or absence of a side channel at the station's cross-section using the
checkboxes in the Side Channel column on the form (an unchecked box indicates absence).
11. Record the presence or absence of quiescent off-channel aquatic habitats, including sloughs,
alcoves and backwater pools using the checkboxes in the Backwater column of the (an unchecked
box indicates absence).
12. Proceed upstream the specified increment to the next station, and repeat Steps 4 through 11.
13. Repeat Steps 4 through 12 until you reach the next transect (the last station will be one increment
prior to the next transect). At this point, select the next transect bubble at the top of the Physical
Habitat Form and complete Channel/Riparian measurements at the new transect (Section 10.4.1).
Then record Thalweg Profile and Woody Debris data and repeat Steps 4 through 12 for each of the
reach segments, until you reach the upstream end of the sampling reach (Transect K). At Transect K,
you will have completed 10 separate Thalweg Profiles, one for each subreach (A to B, B to C, etc.).
Q At every thalweg increment, determine by sight or feel whether deposits of soft/small
< sediments are present on the channel bottom. These particles are defined as substrate equal to
S or smaller than fine gravel (< 16 mm diameter). These soft/small sediments are different from
S3 Fines described when determining the substrate particle sizes at the cross-section transects
^ (Section 10.5.1.1). If the channel bottom is not visible, determine if soft/small sediment deposits
i are readily obvious by feeling the bottom with your boot, the surveyor's rod, or a calibrated rod
i_ or pole.
^ Measure wetted width at each transect (station 0), and midway between transects (station 5 for
i larger streams having 100 measurement points, or station 7for smaller streams having 150
^ measurement points). The wetted width boundary is the point at which substrate particles are
^ no longer surrounded by free water. Estimate substrate size for five locations evenly spaced
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across each midway cross-section at regular cross-sections (Figure 10.1), but at the
supplemental cross-sections, only the size class (not distance and depth) data are recorded.
While recording the width and depth measurements and the presence of soft/small sediments,
the second person evaluates and records the habitat class (Table 10.10) applicable to each of
the 100 (or 150) measurement points along the length of the reach. Make channel unit scale
habitat classifications at the thalweg of the cross-section. The habitat unit itself must meet a
minimum size criteria in addition to the qualitative criteria listed in Table 10.10. Before being
considered large enough to be identified as a channel unit scale habitat feature, the unit should
be at least as long as the channel is wide. For instance, if there is a small deep (pool like) area at
the thalweg within a large riffle area, do not record it as a pool unless it occupies an area about
as wide or long as the channel is wide. If a backwater pool dominates the channel, record PO
(Pool) as the dominant habitat unit class. If the backwater is a pool that does not dominate the
main channel, or if it is an off channel alcove or slough (large enough to offer refuge to small
fishes), select the checkbox to indicate presence of a backwater in the Backwater column of the
form, but classify the main channel habitat unit type according to characteristics of the main
channel. Sloughs are backwater areas having marsh like characteristics such as vegetation, and
alcoves (or side pools) are deeper areas off the main channel that are typically wide and shallow
(Helm 1985, Bain and Stevenson 1999).
Table 10.10 Channel Unit Categories
Channel Unit Habitat Classes1
Class (Code)
Description
Pool (PO)
Still water, low velocity, a smooth, glassy surface, usually deep compared to
other parts of the channel
Glide (GL)
Water moving slowly, with a smooth, unbroken surface. Low turbulence.
Riffle (Rl)
Water moving, with small ripples, waves and eddies waves not breaking,
surface tension not broken. Sound: babbling, gurgling.
Rapid (RA)
Water movement rapid and turbulent, surface with intermittent Whitewater
with breaking waves. Sound: continuous rushing, but not as loud as cascade.
Cascade (CA)
Water movement rapid and very turbulent over steep channel bottom. Much
of the water surface is broken in short, irregular plunges, mostly Whitewater.
Sound: roaring.
Falls (FA)
Free falling water over a vertical or near vertical drop into plunge, water
turbulent and white over high falls. Sound: from splash to roar.
Dry Channel (DR)
No water in the channel, or flow is submerged under the substrate (hyporheic
flow).
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1 Note that in order for a channel habitat unit to be distinguished, it must be at least as wide or long as
the channel is wide (except for off channel backwater pools, which are noted as present regardless of
size).
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10.5.2.2 Large Woody Debris Tally
Large Woody Debris (LWD) is defined here as woody material with a small end diameter of at
least 10 cm (4 in.) and a length of at least 1.5 m (5 ft.). This includes any portion of woody
material that meets those minimum size categories (e.g., include any LWD piece with at least 1.5
meters of its length having a diameter of 10 cm or greater). The procedure for tallying LWD is
presented in Table 10.11. The tally includes all pieces of LWD that are at least partially in the
baseflow channel (Zone 1), in the bankfull channel (Zone 2, flood channel up to bankfull stage),
or spanning above the bankfull channel (Zone 3), as shown in Figure 10.9. LWD in zones 1 and 2
will be tallied together (considered all or in-part in the bankfull channel). The bankfull channel is
defined as the channel that is filled by moderate sized flood events that typically recur every
one to two years. LWD in or above the bankfull channel is tallied over the entire length of the
reach, including the area between the channel cross-section transects. Pieces of LWD that are
not at least partially within Zones 1, 2, or 3 are not tallied.
For each tally (Pieces All/Part in Bankfull Channel and Pieces Bridge Above Bankfull Channel),
the field form (Figure 10.10) 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.
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 10.10) 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 >10 cm. Count each of the LWD
pieces as one tally entry and include the whole piece when assessing dimensions. If you
encounter massive, complex debris jams, estimate their length, width, and height. Estimate the
diameter and length of large "key" pieces and judge the average diameter and length of the
other pieces making up the jam. Record this information in the comments section of the form.
Table 10.11 Procedure: Tallying Large Woody Debris
Large Woody Debris Tally Form
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Note: Tally pieces of large woody debris (LWD) within each segment of stream while the thalweg profile
is being determined. Include all pieces in the tally whose large end is found within the subreach.
1. Scan the stream segment between the two cross-section transects where thalweg profile
measurements are being made.
2. Tally all LWD pieces within the segment that are at least partially within the bankfull channel.
Determine if a piece is LWD [any portion with a small end diameter >10 cm [4 in.], and a length >1.5 m
[5ft.]).
3. For each piece of LWD, determine the class based on the diameter of the large end (0.1 m to < 0.3 m,
0.3 m to <0.6 m, 0.6 m to <0.8 m, or >0.8 m), and the class based on the length of the piece (1.5m to
<5.0m, 5m to <15m, or >15m).
If the piece is not cylindrical, visually estimate what the diameter would be 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 >10 cm (4 in).
4. Tally each piece of LWD in the appropriate diameter and length class tally box in the Pieces All/Part in
Bankfull Channel columns in the Large Woody Debris section of the Physical Habitat Form in the App.
Tally all LWD pieces within the segment that are not actually within the bankfull channel, but are at
least partially spanning (bridging) the bankfull channel. For each piece, determine the class based on
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the diameter of the large end (0.1 m to < 0.3 m, 0.3 m to <0.6 m, 0.6 m to <0.8 m, or >0.8 m), and the
class based on the length of the piece (1.5 m to <5.0 m, 5 m to <15 m, or >15 m).
5. Tally each piece of LWD the appropriate diameter and length class tally box in the Pieces Bridge Above
Bankfull Channel columns in the Large Woody Debris section of the Physical Habitat Form in the App
Note that when entering data via the NRSA App, numbers can be typed directly into tally boxes, or the
"+" and buttons can be used to incrementally change the number.
If no LWD is present in the subreach for a given diameter/length class, enter 0 in the tally box. The NRSA
App contains a "Populate blank boxes with zeros" button which will automatically enter zeros into all
the blank LWD fields. This can be done after all LWD is tallied to quickly enter zeroes in the remaining
fields.
7, Steps 1 through 5 will be repeated at each subreach.
bankfull channel width
V I
ZONE 3
x-— water surface at
/ bankfull flow
ZONE 2
water surface at/
x— baseflow
Figure 10.9 Large Woody Debris Influence Zones (modified from Robison and Beschta, 1990).
LARGE WOODY DEBRIS (10cm small end dian
rieter; 1.5m length)
©
Populate blank boxes with zeros
DIAMET PIECES ALL/PART IN BANKFULL CHANNEL
1.5-5m 5-15m >15m
PIECES BRIDGE ABOV...
1.5-5m 5-15m
>15m
0.1 - <0.3m
—
F
F
F
F
+
—
F
F
F
F
+
0.3 - 0.6m
—
F
F
F
F
+
—
F
F
a
B
+
0.6 - 0.8m
—
~
H
~
H
+
—
~
H
~
H
+
> 0.8m
—
~
H
~
H
+
—
~
H
~
S
+
Figure 10.10 Large Woody Debris Section of Physical Habitat Form in the NRSA App - Wadeable
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10.6 Cross-section Transects on Side Channels
If the wetted channel at a transect is split by an island, and the estimated flow in the side
channel is less than 15% of the total flow, the bank and riparian measurements are made at
each side of the main channel only (the minor side channel is ignored other than to note its
presence on the Thalweg Profile section of the Physical Habitat Form), so one riparian plot is
established on the island as shown in Figure 10.11 (side A). If an island is present at a transect
that creates a major side channel containing 15% or more of the total flow, an additional cross-
section transect is established for the side channel as shown in Figure 10.11 (side B). Separate
substrate, bank and riparian measurements are made for side channel transects. Data from the
additional side channel transect are recorded by checking the "IS THIS A SIDE CHANNEL"
checkbox near the top of the Physical Habitat Form. Unchecking the side channel box will display
the primary transect data once again. Riparian plots established on the island for each transect
may overlap (and be < 10 m shoreward) if the island is less than 10 m wide at the transect.
Islands are distinguished from mid-channel bars by their relationship to bankfull flow: Islands
are not inundated at bankfull stage; bars are part of the main channel and are inundated at
bankfull flow. Collect all channel and riparian cross-section measurements from the side channel
as well as the primary channel. While collecting thalweg profile data (including mid-station
substrate size class) and large woody debris tallies, record the cumulative data on the primary
transect only (e.g., with the side channel check box unchecked).
l/l
>
A) Island and minor side channel
<15% total flow occurs in side channel
No side channel cross section transect
Note presence on field form
Riparian plot established on island
5 m 1 5 m
Instream Fish
Cover Plot
RIPARIAN ;
PLOT
(Right Bank)
PRK/DVP 806
B) Island and major side channel
>15% total flow occurs in side channel
Side channel cross-section must be assessed
Two riparian plots established on island [may overlap)
Data is recorded on additional PHab form
5 m ' 5 m
Instream Fish
Cover Plot
RIPARIAN
PLOT
(Right Bank)
Figure 10.11 Riparian and Instream Fish Cover Plots for a Stream with Minor and Major Side Channels
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10.7 Slope and Bearing
Measure slope and bearing by backsighting between transects (e.g., transect B and A, C and B,
etc.). To measure the slope between adjacent transects, follow the procedure presented in
Table 10.12 or the alternate procedures presented in Table 10.13. Measure bearing following
the procedure presented in Table 10.14. Record slope and bearing data on the Slope and
Bearing Form in the NRSA App.
10.7.1 Measurement of Slope using Level and Stadia Rod
Slope is typically measured by two people, one holding a surveyor's rod and the second sighting
through the surveyor's level. Be sure that the person is holding the marked pole or rod at the
surface of the water. The intent is to get a measure of the water surface slope, which may not
necessarily be the same as the bottom slope. The surveyor's level is leveled according to the
manufacturer's recommendations, which is generally to adjust the three leveling feet until the
bubble is centered. Level is checked in all planes to be measured. If the level does not "self
level" in all measured planes the user should check the instruction manual for suggested
options. Relative elevation readings are made at each transect and the difference between each
elevation reading is calculated and recorded as the change in elevation. NOTE: Multiple transect
elevations can often be made for each setup of the level, but every time the tripod and level
are moved, a second measurement of the last elevation from the last setup is required. You
cannot use elevations from previous setups because the relative height of the level has
changed.
To calculate sinuosity from bearing measurements, it does not matter whether or not you adjust
your compass bearings for magnetic declination, but it is important that you are consistent in
the use of magnetic or true bearings throughout all the measurements you make on a given
reach. Provide a comment on the Slope and Bearing Form about which type of bearings you are
taking, so the measurements can be used to describe reach aspect. Also, guard against recording
reciprocal bearings (erroneous bearings 180 degrees from what they should be). The best way
to do this is to know where the primary (cardinal) directions are in the field: (north [0 degrees],
east [90 degrees], south [180 degrees], and west [270 degrees]), and ensure that your bearings
"make sense."
As stated earlier, it may be necessary to set up intermediate (supplemental) bearing points
between a pair of cross-section transects if you do not have direct line-of-sight along (and
within) the channel between stations (Figure 10.12). This can happen if brush is too heavy, or if
there are sharp slope breaks or tight meander bends. If you would have to sight across land to
measure bearing between two transects, then you need to make one or more supplemental
measurements (i.e., do not "short-circuit" a meander bend). Mark these supplemental locations
with a different color of plastic flagging than used for the cross-section transects to avoid
confusion. Record these supplemental bearing measurements, along with the proportion of the
stream segment between transects included in each supplemental measurement, in the
appropriate sections of the Slope and Bearing Form in the NRSA App. Note that the main
bearing observations are always downstream of supplemental observations (i.e., to the
downstream transect). Similarly, first supplemental observations are always downstream of
second supplemental observations.
Because measurements of slope are a calculation of the elevation difference between transects,
you may sight over land for the purposes of slope only (Figure 10.12). You may need to use
supplemental points in your measurement of slope if visibility is severely limited, but
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supplemental points are not required for slope as they are for measurements of bearing. As a
result, there may be times when you record bearing data for a supplemental point (or two) on
the Slope and Bearing Form; but only record the total elevation change between the two
transects in the primary slope field.
Backsight with
compass and
record
main slope
and bearing
measurements
and % of reach
Supplemental slope
and bearing point
Backsight with
compass and record
supplemental slope
and bearing
Measurements and
% of reach
Backsight
with compass
and record
main slope
and bearing
measurements
and % of reach
PRX/DVP 646
Figure 10.12 Measurements of Bearing Between Transects
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Table 10.12 Procedure: Obtaining Slope Data
Slope Method with Surveyors Level
3.
4.
5.
6.
Instrument Setup:
1. Set up the instrument upstream of Transect A where a clear line of sight is possible to Transect A.
If visibility allows, the instrument may be placed as far upstream as possible, as long as Transect A
is still visible.
Note: In small streams with a clear line of site it may be possible to set the instrument up once and make
readings to several transects from a single set up. Simply record the readings for each transect and do not
skip transects.
2. Extend the tripod legs to approximately eye level and set the legs firmly into the ground; adjust
the legs so that they form a regular triangle and are firmly set with no wobble. Adjust the legs so
that the base plate is approximately level.
Hold the instrument on the tripod and start the centering screw. Ensure the adjustable feet are
roughly evenly adjusted. While the centering screw is still loose, slide the instrument on the base
plate until the bubble is approximately centered in the circular level. Tighten the centering screw.
Adjust the leveling foot screws until the bubble is exactly level in the center circle.
Self-Leveling instruments can now be swiveled gently on the base plate and should maintain level
as long as the tripod remains steady. Check to ensure the bubble indicates the instrument is level
across all planes at which measurements are to be made.
Adjust focus, brightness and parallax according to manufacturer's specifications. The instrument is
ready to make measurements.
Taking Measurements:
1. Determine a location along Transect A to hold a surveyor's rod that will be visible from the
location of the instrument:
Position the staff at Transect A, holding the bottom of the staff at the water level and the
staff as vertical as possible and the numbers facing the instrument.
Site the staff through the instrument and record the reading to the nearest 0.5 centimeter in
a field notebook or other workspace (this value is not entered on the field form).
Move the staff to Transect B and gently swivel the instrument to face the next reading. Hold
the staff as before, vertically, with the bottom at the water level and the numbers facing the
instrument.
Site the staff and record the reading to the nearest 0.5 centimeter (again in a field notebook
or elsewhere).
Subtract the elevation reading at Transect A from the reading at Transect B.
The difference in the readings is the elevation difference that is recorded on the Slope and
Bearing Form. Also be sure to select the "cm" bubble to indicate the units on the value
entered.
Repeat measurements between each transect.
2. Proceed to the next cross-section transect (or supplementary point), and repeat Steps a - f above.
For each transect pair where the above method is used, select "TR" (for surveyors level/transit) in
the Method field. If the above method cannot be used for one or more transect pair(s), see
Section 10.7.2 for alternative methods3.
NOTE: If you are sighting to a supplemental point (required for bearing measurements in some
streams), you may record the elevation difference in the appropriate Supplemental section(s) of the
Slope and Bearing Form. You may also calculate the total elevation change from one transect to the
next and entered only that value in the form. In this case, values would be entered for bearing in the
supplemental field(s), but not for slope.
a.
c.
g-
' Method codes are: CZ=clinometer, 77?=surveyor's level / transit, HL=hand level, l/l/T"=Water tube, L4=laser
level, Omf/?=method not listed (describe in comments section of form).
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10.7.2 Alternate Methods for Obtaining Slope
Because of ease of use, portability, and cost, hand-held clinometers were previously used to
determine slope. In NRSA, the Field Crews will have access to more sophisticated
instrumentation (e.g., surveyor's level), and have field personnel who are experienced in the use
of these instruments. Clinometers should only be used if the slope is greater than 2.75% or if the
surveyors level malfunctions. However, note that when properly used, a roofer's level
(hydrostatic level) can yield slope measurements more precise than even a laser level. The Slope
and Bearing Form in the NRSA App is designed to allow for different methods and/or different
units of measuring elevations or direct measurements of slope. Select the appropriate method
(instead of 77?; method codes are identified in Table 10.12) and mark the % BUBBLE (instead of
the CM bubble) if the method or instrument measures the percent slope rather than the
difference in elevation (Table 10.13).
Table 10.13 Modified Procedure: Obtaining Slope Data (without Surveyor's Level)
Modified Slope Method - Only Use if Surveyor Level is NOT Possible
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Use this procedure (Figure 10.13) if you are starting at the upstream Transect (K), after completing the
thalweg profile and other cross-section measurements at Transects A through K. It should only be used
if you cannot use the surveyor's level.
1. Stand in the channel at the upstream cross-section transect. Determine if you can see the next
transect downstream. If not, you may have to take supplementary slope measurement(s).
While sighting over land is not prohibited during any slope measurement, the clinometer
method is more difficult to use over larger distances, so supplementary measurements are
more likely to be needed.
a. If the next transect downstream still cannot be seen, you will have more than one
supplemental measurements (e.g., you will measure the second supplemental, then
the first supplemental, and finally the main).
2. Mark a surveyor's rod and a calibrated rod (or meter ruler) at the same height. If a shorter
pole or ruler is used, measure the height from the ground to the opening of the clinometer
when it is resting on top.
3. Have one person take the marked surveyor's rod to the downstream transect. Hold the rod
vertical with the bottom at the same level as the water surface. If no suitable location is
available at the stream margin, position the rod in the water and note the depth.
If you have determined in Step 1 that supplemental measurements are required for this
segment, walk downstream to the furthest point where you can stand in the center of the
channel and still see the center of the channel at the upstream transect. Mark this location
with a different color flagging than that marking the transects.
4. Place the base of the calibrated rod at the same relative height as the surveyor's rod (either at
the water surface or at the same depth in the water).
5. Place the clinometer on the calibrated rod at the height determined in Step 2. With the
clinometer, sight downstream to the flagged height on the surveyor's rod at the downstream
transect (or at the supplementary point).
If you are sighting to the next downstream transect, read and record the percent slope in
the Main field on the Slope and Bearing Form for the downstream transect (e.g., J < K),
which is at the bottom of the form (i.e., you are completing the form in reverse order).
Record the Proportion as 100%.
If you are sighting from a supplemental point, record the slope (%) and proportion (%) of
the stream segment that is included in the measurement in the appropriate Supplemental
field of the Slope and Bearing Form. The last sighting to a downstream transect (from either
the upstream transect or the nearest upstream supplemental point) is always recorded as
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the Main reading. This method requires the form to be filled out from the bottom up when
supplemental points are needed. If it was determined in Step 1 that two supplemental
points are needed, you will fill out the third row first.
6. Stand in the middle of the channel at the upstream transect (or supplemental point), and
backsight with your compass to the middle of the channel at the downstream transect (or
supplemental point). Record the bearing (degrees) in the same section of the Slope and
Bearing Form (Supplemental or Main) as you recorded the slope in Step 5 (see Table 10.14 for
details).
7. Proceed to the next cross-section transect (or to a supplementary point) and repeat Steps 3
through 6 above.
Short pole with
clinometer at
height h
Upstream
Transect
Both poles must be at water's
surface or at same depth
Surveyor rod
with flagging at
h8ight/7 6acKs^U°
Downstream
Transect
PRK/DVP 6/06
Figure 10.13 Channel Slope Measurement using a Clinometer
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10.7.3 Method for Obtaining Bearing
Table 10.14 presents the steps necessary to obtain bearing data with a compass in wadeable
streams.
Table 10.14 Procedure: Obtaining Bearing Data
Obtaining Bearing Data
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Use this procedure to backsight from one transect to another using a bearing sighting compass.
Readings (in degrees) are taken from the center of one transect to the center of the next transect
downstream (e.g., when using standard workflow and working upstream, you are sighting back to the
transect you just left).
It does not matter whether you use true or magnetic bearings as long as you are consistent for each
reading.
Take care not to accidentally take a reciprocal reading (one that is 180 degrees off).
1. Stand in the center of the channel at the upstream transect. If you cannot see the next transect
downstream without sighting across land, you will have to take supplementary bearing
measurements (i.e., do not "short-circuit" a meander bend).
2. Hold the compass in line with your body and sight down the "lubber line" or through the sight
window to the center of the next transect downstream (or supplemental point). It will help to have
another person at the center of the transect to which you are sighting. Remember that your line of
sight cannot "cross land."
3. While pointing the compass toward the middle of the transect or supplemental point, read the
bearing (in degrees)
• For many navigational compasses, you will rotate the bezel until the index marks are centered
over magnetic north needle. The base of the lubber line or index mark on the compass now
points to the compass heading on the bezel.
• For some direct reading compasses, the bearing will be displayed already.
4. Record the reading on the Slope and Bearing Form.
5. If you are sighting to the next downstream transect, read and record the bearing in the Main
section on the Slope and Bearing Form. Record the Proportion as 100%.
6. If you are sighting from a supplemental point, record the bearing and proportion (%) of the stream
segment that is included in the measurement in the appropriate Supplemental field of the Slope
and Bearing Form. The sighting to a downstream transect (from either the upstream transect or
the nearest upstream supplemental point) is always recorded as the Main reading.
a. The first measurement taken is from the supplemental point to the downstream
transect. This is recorded in the Main row.
b. The second measurement taken is from either the upstream transect to the first
supplemental, or from the second supplemental to the first supplemental. This is
recorded in the first supplemental row.
c. If two supplemental are needed, then the third measurement taken is from the
upstream transect to the second supplemental. This is recorded in the second
supplemental row.
d. It will be extremely rare to need more than two supplemental points, but if this were to
happen, record additional supplemental measurements in the comment bubble
provided.
7. Proceed to the next transect (or supplemental point), and repeat steps 1 through 6 above.
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10.8 Channel Constraint, Debris Torrents, and Recent Floods
10.8.1 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 10.15. Record data on the Channel
Constraint Form in the NRSA App. First, classify the stream reach channel pattern as
predominantly a single channel, an anastomosing channel, or a braided channel (Figure 10.14):
1. Single channels may have occasional in-channel bars or islands with side channels, but
feature a predominant single channel, or a dominant main channel with a subordinate
side channel.
2. Anastomosing channels have relatively long major and minor channels (but no
predominant channel) in a complex network, diverging and converging around many
vegetated islands. Complex channel pattern remains even during major floods.
3. Braided channels also have multiple branching and rejoining channels, (but no
predominant channel) separated by unvegetated bars. Channels are generally smaller,
shorter, and more numerous, often with no obvious dominant channel. During major
floods, a single continuous channel may develop
After classifying the channel pattern, determine whether the channel is constrained within a
narrow valley, constrained by local features within a broad valley, unconstrained and free to
move about within a broad floodplain, or free to move about, but within a relatively narrow
valley floor. Then examine the channel to ascertain the bank and valley features that constrain
the stream. Entry choices for the type of constraining features are bedrock, hillslopes,
terraces/alluvial fans, and human land use (e.g., a road, a dike, landfill, rip-rap, etc.). Estimate
the percent of the channel margin in contact with constraining features (for unconstrained
channels, this is 0%). To aid in this estimate, you may wish to refer to the individual transect
assessments of incision and constraint. Finally, estimate the "typical" bankfull channel width and
estimate the average width of the valley floor either with a topographic map or visually. If you
cannot directly estimate the valley width (e.g., it is further than you can see, or if your view is
blocked by vegetation), record the distance you can see and mark the appropriate bubble on the
field form.
Table 10.15 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.
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• 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 Physical Habitat Form.
NOTE: If the valley is wider than you can directly estimate, record the distance you can see and mark the
box on the form that indicates this fact.
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A) Anastomosing channel pattern
FLOW
Vegetated islands above bankfull flow. Multiple
channels remain during major flood events.
B) Braided channel pattern
FLOW
_| Unvegetated bars below bankfull flow. Multiple
channel pattern disappears during major flood events.
DVP
Figure 10.14 Types of Multiple Channel Patterns
10.8.2 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 Form in the NRSA App. 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.
10.9 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 in the NRSA App.
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11 FECAL INDICATOR {ENTEROCOCCI)
11.1 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 six 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 six
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.
11.2 Equipment and Supplies
Table 11.1 provides the equipment and supplies needed to collect the fecal indicator sample.
Record the sample data on the Sample Collection Form in the NRSA App.
Table 11.1 Equipment and Supplies: Fecal Indicator Sampling (Wadeable Sites)
For collecting samples
nitrile gloves
pre-sterilized, 250 ml sample bottle
sodium thiosulfate tablet
Wet ice
cooler
For recording measurements
Sample Collection Form in NRSA App
11.3 Sampling Procedure
The procedure for collecting the fecal indicator sample is presented in Table 11.2.
Table 11.2 Procedure: Fecal Indicator (Enterococci) Sample Collection (Wadeable Sites)
Enterococci Sample
1. Put on clean nitrile gloves.
2. Select a sampling location at Transect K that is approximately 1 m from the bank. Approach the
sampling location slowly from downstream or downwind.
3. Lower the uncapped, inverted 250 ml sample bottle to a depth of 1 foot (0.3 m) below the water
surface, avoiding surface scum, vegetation, and substrates. If the water depth 1 meter from the
shore is less than 0.3 meters, submerge the bottle to a lesser depth and be careful not to
contaminate the sample with bottom sediments or detritus.
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 along the transect at 1 m from the bank is too shallow to allow for the sample to be
collected without contamination from bottom sediments, take the sample at an alternate location
along the transect and provide a comment on the form describing the sampling location. If needed,
adjust the sampling depth to avoid contaminating the sample with bottom sediments or detritus.
6. After removing the container from the water, discard a small portion of the sample to allow for
proper mixing before filtering (down to the 250 mL mark on the bottle).
7. Add the sodium thiosulfate tablet, cap, and shake bottle 25 times.
8. On the Sample Collection Form in NRSA App, record the time of collection for the samples by using
the "Now" button or by entering the time in 24-hr format.
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9. Enter the depth of the sample collection (in meters) in the provided fields. This is the approximate
depth from the surface of the water to the mouth of the submerged container, not the depth of the
water at the sampling location.
Storage
10. Store the sample in a cooler on ice to chill (do not freeze immediately). Chill for at least 15 minutes
before filtering.
11. Sample must be filtered and all filters frozen within six hours of collection
11.4 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 14.3 for information regarding processing the
Enterococci samples.
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12 FISH ASSEMBLAGE
12.1 Summary of Method
The fish sampling method is designed to provide a representative sample of the fish community,
collecting all but the rarest fish taxa inhabiting the site. It is 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. 2008, 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.
For wadeable streams, collect fish using a bank or towed (e.g., a towed barge, small watercraft,
or float tube) electrofishing unit (2,500- to 5,000-V; 2.5-5.0 Generator Powered Pulsator [GPP]
unit or equivalent) or backpack electrofisher. Use a backpack electrofishing unit in smaller
streams, when conductivity is appropriate, or in larger streams that are inaccessible with a
towed unit. As a last option, use seining as an alternate method when electrofishing is precluded
by high or low conductivity or extreme levels of turbidity.
There are different protocols for collecting fish from wadeable streams of different sizes (see
Section 12.3). For all wadeable sites, conduct sampling in an upstream direction (i.e., from the
downstream end of the reach), allocating effort (button time) within subreaches (areas between
the cross-section transects). At smaller streams (mean channel width [CW] rounded to the
nearest meter and recorded on the Verification Form is less than or equal to 12 m), sample all
available habitats over the entire sampling reach (40 CW; see Section 3.2). At large wadeable
streams (mean channel width is greater than or equal to 13 m), the initial length of the fish
sampling reach is less than the entire sampling reach, and effort is focused on habitats along the
stream margins. At large wadeable streams, the minimum sample reach is 20 CW (five
subreaches). If you have not collected 500 individuals at the end of the 20 CW, sample
additional subreaches in their entirety until you obtain at least 500 individuals, or until you have
sampled the entire sampling reach (40 CW; 10 subreaches). Record information related to
sampling effort on the Fish Gear Form in the NRSA App. Record species identification and
enumeration data on the Fish Collection Form in the NRSA App.
12.2 Equipment and Supplies
Table 12.1 shows the checklist of equipment and supplies required to complete the fish
assessment. This checklist is similar to the one presented in Appendix A, which is used at the
base location to ensure that all of the required equipment is brought to the site.
Table 12.1 Equipment and Supplies: Fish Collection (Wadeable Sites)
For collecting
samples
Bank or towed electrofishing equipment
(e.g., towed barge or mounted on a small
watercraft or float tube), including
variable voltage pulsator unit, wiring
cables, generator, and electrodes
Backpack electrofishing unit with anode
and cathode
Linesman gloves, boots, and other
necessary safety equipment
1 Scalpel for slitting open large fish
before preservation.
1 container of 10% buffered formalin
Small mesh bags or several Leak proof
HDPE jars (various sizes from 250 mL - 4L)
for fish voucher specimens
2 non-conducting dip nets with % inch
mesh
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Dip nets
Extra electrofishing unit batteries
Scientific collection permit(s)
Digital camera with extra memory card &
battery
1 Laser rangefinder (optional)
Polarized sunglasses and hats
10 ft x 6 ft Minnow or Bag Seine with %
inch mesh (additional 4' depth seine may
also be used)
1 Minnow net for dipping small fish from
live well
2 measuring boards (3 cm size classes)
1 set Fish ID keys
Field Operations Manual
20 ft x 6 ft Minnow or Bag Seine with %
inch mesh (additional 4 ft depth seine
may also be used)
Buckets
For recording
measurements
Sheet of sample labels and voucher
specimen tags (for unknown/range
extension voucher samples)
Sheet of sample labels and voucher
specimen tags (for QA voucher samples)
Fish Gear Form in NRSA App
Fish Collection Form in NRSA App
Clear tape strips
Soft (#2) lead pencils for writing on
voucher inner labels
Fine-tip indelible markers
12.3 Sampling Procedures
The reach length sampled for fish varies based on the mean width of the stream used to
establish the length of the sampling reach (which has been rounded to the nearest meter as per
the directions in Section 3.2) and on the number of individuals collected (Figure 12.1). For small
wadeable streams (mean CW from the Verification Form <12 meters), follow the protocol
presented in Section 12.3.2. Sample the entire reach (150 m to 40 CW, 10 subreaches) and
move the electrofishing unit (towed barge or backpack) within each subreach to sample both
shorelines as well as the mid-channel.
For large wadeable streams (mean CW from the Verification Form > 13 m), follow the protocols
described in Section 12.3.3. For large wadeable streams the minimum length for fish sampling is
20 CW (5 of the 10 subreaches). If a minimum of 500 fish are not collected after sampling this
minimum fishing reach, sample additional subreaches in their entirety until at least 500 fish are
collected, or all 10 subreaches have been sampled. Stop sampling when you reach Transect K
(the end of the sampling reach), regardless of the number of individuals collected.
Table 12.2 summarizes the fishing protocols for small and large wadeable streams. More
detailed steps for fishing small and large wadeable streams are found in the following sections.
If conditions prohibit any type of electrofishing, collect fish by seining as described in Table 12.5.
The objective of seining is to collect species and relative abundance data that is comparable to
what would have been obtained by electrofishing at the site. If seining is used, record all fish
collected with seining protocols on separate lines of the Fish Collection Form.
It is important that you record the total reach length that was sampled for fish, as this is used
along with the number of fish collected to determine sampling sufficiency. Data from streams
that were not sufficiently sampled for fish cannot be used to assess stream condition based on
the fish assemblage.
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12.3.1 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
abundance of fish 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
than or equal to 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 12.2 Summary of Wadeable Fishing Protocols
Small Wadeable (mean channel width [CW1 from Verification Form < 12 meters)
• Fish sampling reach length will be between 150 m (CW < 4 m) and 40 CW.
• Subreaches will be between 15 meters and 4 channel widths long
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
Large Wadeable (mean channel width [CW1 from Verification Form > 13 meters)
• Initial minimum fish sampling reach will be 20 CW (5 subreaches).
• Subreaches will be 1/10 of the sampling reach in length
S Fish each subreach in a swath 8 meters from bank in pairs of subreaches starting at a
random 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. Put another way, sample
for 700 seconds and if you have not reached the next transect, stop fishing and move to
that transect.
S Minimum fish number is 500 or until 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 Wadeable Stream: Mean Channel Width < 12 m
Large Wadeable Stream: Mean Channel Width > 13 m
< 20 x Channel Width >
20 CW (5 subreaches), continue fishing next subreach
(alternating bank after every two subreaches) until
either 500 individuals are collected, or Transect K is
reached (10 subreaches [40 CW] have been sampled)
Figure 12.1 Reach Layouts for Fish Sampling at 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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12.3.2 Small Wadeable Streams
Table 12.3 describes the procedure for collecting fish in small wadeable streams. The sampling
crew should consist of one electrofisher operator, one dip-netter (1/4" mesh dip net), and an
optional bucket carrier (who may also have a net to aid in transferring fish to the livewell). An
anode with a net cannot substitute for a netter. For safety, all crew members are required to
wear non-breathable waders and insulated gloves. To aid vision, wear polarized sunglasses and
a hat or visor. See Appendix E for example starting settings for electrofishing using backpack,
towed barge, and boat (Temple 2018). These are only suggestions; the final determination of
settings is decided by the lead fish taxonomist.
Begin sampling at the downstream end of the sampling reach defined for the site (Figure 12.1)
and proceed upstream. Sample the entire reach, which will be between 150 m and 40 channel
widths (10 subreaches). Total button time will vary between 500 and 700 seconds per subreach.
Conduct sampling by subreach (area between transects), but you do not have to allocate effort
equally among all 10 subreaches. Whenever possible, process fish at the end of each subreach
to minimize mortality and stress to fish.
Table 12.3 Procedure: Electrofishing (Small Wadeable Streams)
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Electrofishing Procedures in Small Wadeable Streams
1) Verify or enter the fish sampling date in the header section of the Fish Gear Form in the NRSA App. The
date in this field will be automatically entered based on the date recorded on the Verification Form. If
you have not filled out the Verification Form or if the date of the fish collection is different, you can
edit the date here. Note however that changing the value on this form will not change the value on
the Verification Form.
2) Decide if you will be able to sample the site for fish.
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, select Not Fished - No Permit.
b) If site conditions prevent barge or backpack electrofishing (e.g., no access, safety concerns,
ambient conductivity is too high or too low to use a barge or backpack electrofishing unit),
determine if you can sample by seining.
i) If yes, follow the procedures presented in Table 12.5.
ii) If not, select Not Fished - Site Conditions Prohibit Sampling. Provide a comment describing
why site conditions prohibit sampling.
c) If you can determine that > 50% of the required fish sampling reach (75 m or 20 CW; 5 subreaches)
cannot be sampled, select Not Fished/Fishing Suspended - Can't sample >50% of required reach.
d) If you cannot sample because of equipment problems, select Not Fished - Equipment Failure.
e) At a very small and very shallow stream, if you cannot attempt to sample, but are very confident
that no fish are present (i.e., you do not observe any at any point along the sampling reach), then
select Not Fished - No Fish Observed.
f) If you cannot sample for any other reason, select Not Fished - Other and provide a comment
indicating why sampling cannot occur.
3) If you can begin to sample, select Small Wadeable in the Fish Sampling Protocol section.
a) Proceed to the downstream end of the sampling reach (Transect A).
b) For safety, everyone must wear personal floatation devices, non-breathable waders, foot
protection, and insulated linesman's gloves.
c) To aid vision while netting fish, wear polarized sunglasses and a hat or visor.
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4) Select 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 (note whether the conductivity value is corrected to 25 °C).
5) Mark either Backpack, Bank or Towed Barge in the Primary Electrofishing Gear section of the Fish Gear
Form. Mark Towed Barge for any electrofishing unit that is towed (e.g., canoe, kayak or float tube).
a) Do not use any secondary electrofishing gear in a small wadeable stream.
6) Operation of Bank or Towed 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 stream.
i) See Tables in Appendix E.
ii) If the electrofishing system only lets you select High and Low voltage (rather than a specific
voltage), record the setting used on the Fish Gear Form.
iii) If your conductivity meter cannot measure ambient conductivity, you can "uncorrect" specific
conductance at 25 °C to ambient conductivity using the following equation:
(1) Ambient conductivity=Specific conductance x (l+([water temp-25 °C] x 0.02)).
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 pulse
width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-120 Hz.
d) If the electrofishing system only lets you adjust the percent of power, record the value on the Fish
Gear Form.
e) 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.
f) Once you have determined the appropriate settings, record them on the Fish Gear Form. Start
cleared clocks and resume fishing.
g) Note: Some electrofishers do not meter all the requested settings; provide what you can.
h) If button time is not metered, estimate it with a stop watch and provide a comment in the NRSA
App.
7) 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 stream.
i) See Tables in Appendix E.
ii) If your conductivity meter cannot measure ambient conductivity, you can "uncorrect" specific
conductance at 25 °C to ambient conductivity using the following equation:
(1) Ambient conductivity=Specific conductance x (l+([water temp-25 °C] x 0.02)).
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 pulse
width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-120 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.
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e) Once you have determined the appropriate settings, record them on the Fish Gear Form. Start
cleared clocks and resume fishing.
f) Note: some electrofishers do not meter all the requested settings; provide what you can.
g) If button time is not metered, estimate it with a stop watch and provide a comment in the NRSA
App.
b)
c)
8) Once the settings on the electrofisher are adjusted properly to sample effectively and minimize injury
and mortality, begin sampling at the downstream end of the reach (Transect A) and fish in an
upstream direction.
a) The minimum reach length is 150 m. The maximum reach length for this protocol is 40 CW.
i) Search for fish even if the stream is extremely small, and it appears that sampling may
produce no specimens.
ii) Button time should range from 500 to 700 sec per subreach. Actual time will depend upon
the water conditions, the diversity and complexity of the available habitat, and on the
number offish present in the reach.
(1) If the electrofisher is highly effective and the fish are staying stunned longer and the
water is clear and flowing slowly then button time will be much lower than it will be in a
system where it is more turbid, flowing faster, and the fish are not being stunned as well.
(2) 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 and less time to subreaches
with simple habitat.
Depress the switch and slowly sweep the electrode from side to side.
Sample all habitat types (deep, shallow, fast, slow, complex, and simple). Avoid the temptation to
focus sampling only in the richest habitat types.
i) For available cut-bank and snag habitats, 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.
ii) In fast, shallow water, sweep the anode and fish downstream into a net.
iii) In stretches with deep pools, fish the margins of the pool as much as possible, being
extremely careful not to step or slide into deep water.
d) Keep the cathode near the anode if fish catch is low.
9) The netter, holds the net 1 to 2 ft from the anode, follows the operator, nets stunned individuals, and
places them in a bucket.
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) Continue upstream until you reach the next transect (end of subreach).
a) Process fish and/or change water after each subreach to reduce mortality and stress.
i) Although not required, you may note amphibians and reptiles captured on the Fish Collection
Form.
b) Release fish in a location that eliminates the likelihood of recapture.
11) Repeat Steps 8-10 until all 10 subreaches are sampled (i.e., you reach transect K).
12) After sampling all 10 subreaches, record the final length of the fish sampling reach in the Primary
Electrofishing Gear section of the Fish Gear Form.
a) If you suspend sampling before completing all 10 subreaches, record the actual length that was
sampled, and mark the reason for the suspension in the Fish sampling - Not Routine or Suspended
section of the Fish Gear Form.
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b) If you did not collect any fish, mark Fished - None Collected in the Fish sampling - Not Routine or
Suspended section of the Fish Gear Form.
13) In the Primary Electrofishing Gear section of the Fish Gear Form, record the total button time
expended for electrofishing, the total time spent sampling, and the length of the total fish sampling
reach (recorded in Step 12) sampled by electrofishing.
14) Indicate whether conditions allowed for sufficient sampling on the Fish Gear Form (Yes, No, Not Sure).
If you marked No or Not Sure, provide an explanatory comment.
15) Note the general response of fish to your final electrofishing settings as either:
a) Immobilized (no swimming motions due to electrical field). Includes narcosis (slack muscles) and
tetany (rigid muscles).
b) Inhibited Swimming (unbalanced swimming induced by the electrical field). Includes taxis
(movement, usually towards the anode), pseudo-taxis (movement, but fish are unconscious and
belly-up), and oscillotaxis (movement without orientation).
c) Escape (upright avoidance swimming).
16) Record the total length of the stream that was sampled for fish on the Fish Gear Form.
12.3.3 Large Wadeable Streams
Table 12.4 describes the procedure for collecting fish in large wadeable streams. The
electrofishing crew should consist of one electrofishing operator, and one dip netter and an
optional bucket carrier (who may also have a net to aid in transferring fish to the live well). An
anode with a net cannot substitute for a netter. For safety, all crew members are required to
wear non-breathable waders and insulated gloves. Polarized sunglasses and caps to aid vision
are also required. See Appendix E for example starting settings for electrofishing using
backpack, towed barge, and boat (Temple 2018). These are only suggestions; the final
determination of settings is decided by the lead fish taxonomist.
For large wadeable streams with a mean channel width (from the Verification Form) >13 m, the
minimum fish sampling reach is 20 channel widths (5 subreaches). As shown in Figure 12.1,
begin sampling at Transect A on a randomly determined bank and fish a section of the subreach
that extends approximately 8 m from the bank in an upstream direction. Within each subreach,
fish the near bank habitat as well as midstream habitat within the 8 meter sampling area for a
button time of ~700 seconds. When 700 seconds are reached, stop electrofishing unless you are
"pushing" a large school offish, in which case continue fishing until you capture them at a break.
To reduce stress and mortality, net immobilized fish immediately and deposit into a bucket or
live-well for processing. Whenever possible, process fish at the end of each subreach to
minimize mortality and stress to fish. At the end of the minimum fish sampling reach (20 CW or
5 subreaches), 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. 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%. Stop sampling when you reach Transect
K (the end of the entire 40 CW sampling 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).
Table 12.4 Procedure: Electrofishing (Large Wadeable Sites)
Electrofishing Procedures in Large Wadeable Streams
1) Verify or enter the fish sampling date in the header section of the Fish Gear Form in the NRSA App.
The date in this field will be automatically entered based on the date recorded on the Verification
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Form. If you have not filled out the Verification Form or if the date of the fish collection is different,
you can edit the date here. Note however that changing the value on this form will not change the
value on the Verification Form.
2) Decide if you will be able to sample the site for fish.
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, select Not Fished - No Permit.
b) If site conditions prevent towed or backpack electrofishing (e.g., no access, safety concerns,
conductivity is too high or too low to use a barge or backpack electrofishing unit), determine if
you can sample by seining.
i) If yes, follow the procedures presented in Table 12.5.
c) If not, select Not Fished - Site Conditions Prohibit Sampling. Provide a comment indicating why site
conditions prohibit sampling.
d) If you can determine that > 50% of the required fish sampling reach cannot be sampled (20 CW; 5
subreaches), select Not Fished/Fishing Suspended - Can't sample >=50% of required reach.
e) If you cannot sample because of equipment problems, select Not Fished - Equipment Failure.
e) If you cannot sample for any other reason, select Not Fished - Other and provide a comment
about why sampling cannot be completed.
3) If you can begin to sample, select Large Wadeable in the Fish Sampling Protocol section.
a) Proceed to the downstream end of the reach (Transect A).
b) For safety, everyone must wear personal floatation devices, non-breathable waders, foot
protection, and insulated linesman's gloves.
c) To aid vision while netting fish, wear polarized sunglasses and a hat or visor.
4) Select 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.
5) Select either Backpack or Bank or Towed Unit in the Primary Electrofishing Gear section of the Fish
Gear Form.
a) Do not use any secondary electrofishing gear in a large wadeable stream.
6) Operation of Bank or Towed 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 ambient conductivity of the river (i.e., not corrected
to 25 °C).
i) See Tables in Appendix E.
ii) If the electrofishing system only lets you select High and Low voltage (rather than a specific
voltage), record the setting used on the Fish Gear Form.
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 pulse
width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-120 Hz.
d) If the electrofishing system only lets you adjust the percent of power, record the value on the Fish
Gear Form.
e) 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
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Electrofishing Procedures in Large Wadeable Streams
minimize mortality or injury to large fish. If mortalities occur, first decrease pulse rate, then
voltage, then pulse width.
f)
Once you have determined the appropriate settings, record them on the Fish Gear Form. Start
cleared clocks and resume fishing.
g)
Note: some electrofishers do not meter all the requested settings; provide what you can.
h)
If button time is not metered, estimate it with a stop watch and provide a comment in the NRSA
App.
7)
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 ambient conductivity of the river.
i) See Tables in Appendix E.
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 pulse
width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-120 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.
f)
Note: some electrofishers do not meter all the requested settings; provide what you can.
g)
If button time is not metered, estimate it with a stop watch and provide a comment in the NRSA
App.
8)
Once the settings on the electrofisher are adjusted properly to sample effectively and minimize injury
and mortality, begin sampling at the downstream end of the reach (Transect A). Randomly choose a
bank on which to start and fish in an upstream direction within 8 m of the shoreline.
For the large stream protocol, the minimum initial fish sampling reach length is 5 subreaches
9)
When using a towed electrofishing unit, the minimum crew size for electrofishing is three.
a)
The operator must remain actively at the control box and navigate the towed electrofishing unit.
b)
The probe operator will use one probe.
10) When using a backpack electrofishing unit, the minimum crew size is two (one operator with the
probe and one netter). An anode outfitted with a net cannot substitute for a netter.
11) Depress the switch and slowly sweep the electrode from side to side.
a)
Search for fish even if it appears that sampling may produce no specimens.
b)
Sample all habitat types (deep, shallow, fast, slow, complex, and simple). Avoid the temptation to
focus sampling only in the richest habitat types.
i) In slack water area (e.g., available cut bank and snag habitats), 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.
ii) In fast, shallow water, sweep the anode and fish downstream into a net.
iii) In stretches with deep pools, fish the margins of the pool as much as possible, being
extremely careful not to step or slide into deep water.
c)
Keep the cathode near the anode if fish catch is low.
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12) The netter, holds the net 1 to 2 ft from the anode, follows the operator, nets stunned individuals, and
places them in a bucket.
a) Use a 6 mm (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.
b) 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.
13) Continue upstream until you reach the next transect (end of subreach).
a) The total button time within each subreach should be -700 sec. depending upon the habitat
complexity, you can vary the distance actively fished to allocate the available button time
throughout the subreach.
b) Process fish after each subreach to reduce mortality and stress. Release fish in a location that
eliminates the likelihood of recapture.
i) Although not required, you may note amphibians and reptiles captured on the Fish Collection
Form.
14) Move to the opposite bank when necessary (see Figure 12.1). Repeat Steps 11-14 until you have
sampled the required length of stream.
a) If you have to suspend sampling before completing the required fish sampling reach, record the
actual length that was sampled, and mark the reason for the suspension in the Fish sampling -
Not Routine or Suspended section of the Fish Gear Form.
15) If the minimum reach length has been met (20 X mean stream width) determine the total number of
individuals collected.
a) If the total is < 500, sample one or more additional subreaches in their entirety until at least 500
individuals have been collected and processed, or you sample all 10 subreaches. Go to Step 16.
b) If you collect > 500 individuals, determine if a single irruptive species comprises > 50% of the
total number of individuals.
i) If an irruptive species makes up > 50% of the sample, sample one or more additional
subreaches to bring the proportion of the irruptive species below 50%. Go to Step 16.
ii) If not, go to Step 16.
c) If you have sampled all 10 subreaches (i.e., you have reached Transect K), go to Step 16.
16) After sampling, record the final length of the fish sampling reach in the Primary Electrofishing Gear
section of the Fish Gear Form.
a) If you did not collect any fish, select Fished - None Collected in the Fish sampling - Not Routine or
Suspended section of the Fish Gear Form.
17) In the Primary Electrofishing Gear section of the Fish Gear Form, record the total button time
expended for electrofishing (should be -700 sec per subreach sampled), the total time spent sampling,
and the length of the total fish sampling reach (recorded in Step 15 or 16) sampled by electrofishing.
18) Indicate whether conditions allowed for a sufficient sampling effort on the Fish Gear Form (Yes, No,
Not Sure). If you marked either No or Not Sure, provide an explanatory comment.
19) Note the general response behavior of fish to your final electrofishing settings on the Fish Gear Form
as either:
a) Immobilized (no swimming motions due to electrical field). Includes narcosis (slack muscles) and
tetany (rigid muscles).
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b) Inhibited Swimming (unbalanced swimming induced by the electrical field). Includes taxis
(movement, usually towards the anode), pseudo-taxis (movement, but fish are unconscious and
belly-up), and oscillotaxis (movement without orientation).
c) Escape (upright avoidance swimming).
20) Record the total length of the stream that was sampled for fish on the Fish Gear Form. This total
length should coincide with the end of a subreach unless fishing was suspended prematurely.
12.4 Seining
In small or large wadeable streams where conditions prohibit electrofishing, use seining only as
the last option for collecting fish. Seining is not to be used in concert with electrofishing. If
conditions are such that seining is the only method used, provide a justification in the comment
bubble provided in the Sampling Protocol section at the top of the Fish Gear Form. Table 12.5
presents the procedure for seining wadeable streams. The intent of the seining effort is to
provide comparable data to electrofishing.
Although wadeable electrofishing techniques typically work best in an upstream direction,
seining may work best moving downstream. Allocate seine hauls so that the snag, edge, and
mid-channel habitats are fished thoroughly. In general, edge and snag habitats will be sampled
using narrower seines over shorter distances, while mid-channel habitats will be sampled using
longer seines over longer distances. Generalized habitat seining procedures are presented in
Table 12.5. Depending upon habitat types and complexity, use 2 to 3 crew members. Two crew
members move the seine; an optional third person creates and maintains a bag in the seine in
area with higher velocities, or agitates rocks in riffles or snags.
To avoid mortality, process fish after each seine haul. Record all fish collected with seining
protocols on separate lines on the Fish Collection Form from those lines used for fish collected
by electrofishing.
If you seine, record information for the seine hauls on the Seine Gear sections of the Fish Gear
Form to track effort.
Table 12.5 Procedure: Seining (Wadeable Sites)
Procedures for Seining at All Wadeable Sites
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1) Use seining as a last option only (e.g., when electrofishing is ineffective due to high conductivity or
extremely high turbidity). Do not use seining as a supplementary method to electrofishing. Fish
sampling reach must be safely wadeable.
a) If site conditions are such that only seining is used, provide an explanation in the comment bubble
provided in the Sampling Protocol Comments section at the top of the Fish Gear Form.
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) Verify or enter the fish sampling date in the header section of the Fish Gear Form in the NRSA App.
The date in this field will be automatically entered based on the date recorded on the Verification
Form. If you have not filled out the Verification Form or if the date of the fish collection is different,
you can edit the date here. Note however that changing the value on this form will not change the
value on the Verification Form.
3) Select the pertinent protocol and size class in the Fish Sampling Protocol section,
a) Proceed to the downstream end of the reach (Transect A).
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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 12.1
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) Select 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.
Note whether the conductivity value is corrected to 25 °C.
5) Select 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 crew members (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, select the type and
record the dimensions in the Secondary Seine Net section of the Fish Gear Form.
6) Determine the length of the fish sampling reach and the number of subreaches that should be
sampled (refer to Table 12.2).
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.
8) 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 offish (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.
b) Riffle Habitats
i) Use two crew members, 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.
ii) Starting no more than 3 m upstream, a third crew member kicks the substrate and overturns
rocks, proceeding quickly downstream toward the net.
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iii) When the area is thoroughly kicked, quickly raise and bag the net.
c) Pool, Backwater, and Bar Habitats (Slack water)
i) Use two crew members, each tending a seine pole. Pull the seine across the pool using
shallow riffles or banks as barriers. A third crew member 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.
iv) Pull the bag completely to shore at a predesignated point.
d) Glide or Run Habitats (noticeable current)
i) Use two crew members, each tending a seine pole. Pull the seine diagonally across the glide
towards the bank. If necessary, a third crew member 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.
e) Edge Habitats
i) Edge habitats may be shallow too deep with complex to uniform habitat, and may include
undercut banks.
ii) Use two crew members, each tending a seine pole. Seine along the nearshore area.
iii) The near bank crew member moves along the shore while jabbing along any undercut or
small structure. The other crew member 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.
iv) 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.
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 crew members, 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, then quickly lift the seine to the water surface. You can use a third crew member 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 crew
member moves along the snag, while jabbing along its length. The other crew member 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.
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a) Record identifications, tallies, and voucher information on the Fish Collection Form. Record all fish
collected with seining protocols on separate lines on the field Fish Collection Form from those
lines used for fish collected by electrofishing.
10) At the end of all sampling, determine the total number of hauls, the average haul length, the total
time spent seining, and the total fish sampling reach length sampled for each type of seine. Record the
totals in the Primary and Secondary Seine Wet sections of the Fish Gear Form.
11) Indicate whether conditions allowed for sufficient sampling on the Fish Gear Form (Yes, No, Not Sure).
If you marked No or Not Sure, provide an explanatory comment.
12) Record the total length of the river that was sampled for fish on the Fish Gear Form. This total length
should coincide with the end of a subreach unless fishing was suspended prematurely.
12.5 Processing Fish
Process the fish at the end of each subreach or pairs of subreaches, as described in Table 12.6.
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.
If you use a seine to collect fish, please record all fish collected with seining protocols on
separate lines on the field Fish Collection Form from those lines used for fish collected by
electrofishing.
12.5.1 Identification and Tallying
Record species identifications, tallies, and other information for individuals collected on the Fish
Collection Form in the NRSA App. While recording data on the Fish Collection Form in the App,
select the species from the drop-down list and use the tally boxes to enter the number of each
fish species in each size category. As you collect additional fish, edit the tallies in the existing
lines of data or add new lines of data for new species. If more than 14 lines of fish data are
needed, tap the Add New Fish Collection Line button to add a new data line to the form.
When entering data in the tally boxes, numbers can be typed directly into tally boxes, or the "+"
and buttons can be used to incrementally change the number. The same species can be
entered on multiple rows if necessary (e.g., you did not realize the species had already been
collected or an unknown fish is later determined to be a species already collected). As fish are
tallied, a running total of fish collected is automatically calculated at the top of the form.
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 (15 cm [6 inch] 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, Page et al. 2013). Appendix D provides a
list of species names to be used, based on the current cumulative taxa list developed for NRSA.
If you believe a specimen is nonindigenous to the site, mark it as Introduced on the Fish
Collection Form. If you suspect it represents a potential range extension for the species, prepare
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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 12.5.6).
12.5.2 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 by typing directly
into the Common Name field. 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 Section 12.5.6) or take good digital photographs (Section 12.5.3) 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(s) to label your
jar(s) of unknown to track from which sites the unknowns originated.
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Table 12.6 Procedure: Processing Fish (Wadeable Sites)
Fish Processing
1) Ensure that the date offish collection is accurately recorded on the top of the Fish Gear Form. This
date will be attributed to all fishing activities including the collection of vouchers and fish plug tissue
samples. If fishing occurs over multiple days, use the first day of fishing as the date entered on the
Fish Gear Form and include a comment in the Sampling Protocol section of the Fish Gear Form that
provides all dates offish collection.
2) Process individuals collected as often as necessary to avoid mortality. You may record a single species
on multiple lines of the collection form (e.g., use separate lines for individuals collected in multiple
subreaches or collected with a secondary gear type) or simply amend the tallies of fish on data rows
previously used. Fish do not need to be separated by subreach.
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 stream immediately after processing. Release fish in an area
where recapture is unlikely.
3) Only identify and process individuals > 25 mm (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.
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 previous NRSA surveys.
(1) Record the complete common name. Avoid using shortened names (e.g., stoneroller,
carp, bass, etc.). Select the common name from the provided drop-down list to ensure
correct spelling, etc. The search bar in the drop-down selection window can be used to
search for any part of a fish name to speed data entry. Once you see the common name
you are searching for, tap on it to enter it into the data form.
(2) If you use a non-standard name, you must type the name directly into the Common
Name field and include a comment which provides the taxonomic reference for the
name.
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) by typing in the Common Name field.
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 field 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 by selecting it from the drop-down list. For other hybrids
record the common name of both species (e.g., Green sunfish x Bluegill, Cutthroat trout x
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Rainbow trout) which may also be included in the drop-down list. 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
field and see Step 9.
5) If you know the species is not native to this location, mark as Introduced? If you cannot evaluate the
native/introduced status of fish collected, select the checkbox at the top of the collection form. This
will confirm that blank values for the Introduced? field are missing, rather than being presumed as
native.
6) Visually estimate the total length of each individual (a measuring board is not necessary but can be
used to calibrate one's eye to the size groupings or can be used to verify a fish length). Keep a
running tally in the appropriate Tally and Counts section (< 6 in., 6-12 in., 12-18 in., or > 18 in.) of the
Fish Collection Form. When entering data in the tally boxes, numbers can be typed directly into tally
boxes, or the "+" and buttons can be used to incrementally change the number.
a) If all individuals of a species appear to be the same size, provide a 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 of individuals observed with one or more anomaly in the Anom Count field of the Fish
Collection Form.
a) NOTE: Do not include injuries from collecting, handling, or processing fish, or from parasites in the
external anomaly tally.
8) If an individual has died due to electrofishing or handling, include it in the running tally for the species
as normal. After you process all of the individuals of a species, record the total mortality for the
species in the Mortality/Count field of the Fish Collection Form.
9) If you are retaining individuals of the species as part of the preserved Unknown/Range Extension
Unk/Rng) voucher sample:
a) Select the Voucher Unk/Rng checkbox on the form.
b) Assign the species the next available voucher specimen tag, and record the number in the
Voucher Tag # field.
i) If you take one or more photographs of the species instead of or in addition to preserving
specimens, select the Voucher Photo box. Include the specimen tag in all photos of the
species. 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 field of the Fish 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). Line numbers
and tag numbers do not need to be the same because the same species may occur on
multiple lines, but any given species should only be assigned a single tag number.
10) If you are retaining specimens as part of a preserved QA voucher sample for the site:
a) Select the Voucher QA checkbox on the form.
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i) NOTE: This should be marked at least once for all species collected at the site (including
unknowns) to create a complete voucher collection for the site.
b) Use the sheet of labels and tags for the QA voucher sample (the primary jar label has a
preprinted sample ID number and is found on the same label sheet as other sample labels
provided in the site kit and extra jar labels are provided on a separate sheet). Assign the species
the next available voucher specimen tag number (start with Tag 01 and assign tag number
sequentially). Record the specimen tag number in the Voucher Tag # field of the Fish Collection
Form.
c) If you take one or more photographs of the species instead of or in addition to preserving
specimens, select the Voucher Photo box. Include the specimen tag in all photos of the species.
d) Record the number of individuals retained for the preserved voucher sample in the Vouchers
Retained field of the Fish 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 (i.e., one tag per line on the collection form). Line numbers and tag numbers do
not need to be the same because the same species may occur on multiple lines, but any
given species should only be assigned a single tag number.
11) Repeat Steps 2 through 10 for each subreach sampled. Add to the tally counts by editing the numbers
in the tally field or by using the "+" and buttons. The same species can be entered on multiple
rows if necessary (e.g., you did not realize the species had already been collected or an unknown fish
is later determined to be a species already collected). As fish are tallied, a running total offish
collected is automatically calculated at the top of the form.
12) Record the fish collected with seining methods on a separate lines from those lines used for fish
collected by electrofishing.
13) At the end of sampling, follow the appropriate procedure to prepare the preserved voucher samples
(UNK/RNG and/or QA Vouchers) and/or select specimens for tissue (fish plug) 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. Note that sample IDs are not assigned to UNK/RNG Voucher samples.
15) Complete an inner label by filling in the site ID, date and visit number with a pencil. For QA Vouchers,
also write the sample ID in the space provided.
16) If you did not prepare a QA voucher sample, mark No Voucher Preserved on the bottom of the Fish
Collection Form (this is akin to the 'no sample collected' checkbox associated with other sample
types).
a) Otherwise complete a sample jar label for the QA Voucher sample. Attach it to the sample jar
and cover it with clear tape. Extra jar labels are provided that will be used to label additional jars
belonging to the same voucher sample by writing the Voucher Sample ID on the label and
indicating the number of the jar on the label (e.g., Jar 1 of 2).
b) Record the total number of jars in the "# of Jars" field. Also select the "Preserved" checkbox to
confirm that the voucher sample has been properly preserved.
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17) Record the file names of any photovouchers taken on the Fish Collection Form in the NRSA App by
completing the Following steps.
a) Enter the tag number assigned to the species in the Voucher Tag # field. Use 2-digit format (e.g.,
01, 02, etc.).
b) Select the Voucher Photo checkbox. Doing so will open a Photo File Name field that is
automatically populated with the base portion of the photo naming convention (Site ID + Visit
number + tag number).
c) Each photo taken of the same individual or species will need to be assigned a letter (starting with
"a"). Indicate the beginning and the end of the sequence in the Sequence field. For example, a
series of four photos of an individual would be assigned a sequence of "a-d".
d) Prior to uploading the images to the NARS SharePoint site, name each image file as: Site ID + Visit
number + tag number + sequence (e.g., NRS238_WY-10001_Vl_tag01a).
18) If you did not collect any fish from the entire fish sampling reach, select Fished - None Collected in
the Fish Sampling - Not Routine or Suspended section of the Fish Gear Form.
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12.5.3 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 in electronic format.
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, if the specimen is part of
either the unknown/range extension or QA voucher collection, 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, ocular/oral
orientation, fin rays, gill arches, mouth structures, etc.). It may also be necessary to photograph
males, females, or juveniles to depict key identifying features.
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 personal computer 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., NRS23_WY-10001_Vl_tag01a.jpg). Record the file names on the Fish Collection
Form. You should review your photos to confirm that they provide sufficient details to allow
someone else to confidently confirm your identification using only your image files.
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 post
all images to the NRSA SharePoint site.
12.5.4 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
initial fish collection data via the NRSA App (which should be done immediately after leaving the
site), the data can easily be amended by editing the Fish Collection Form and resubmitting that
form to update the database.
In addition to a UNK/RNG voucher sample (if needed), you will prepare an additional QA voucher
sample (Section 12.5.7) at some sites. A QA voucher sample will be performed at a pre-
designated set of sites and includes preserved specimens (and/or photographs) of all species
collected at a site (including the unknowns). Use the voucher specimen tags and sample labels
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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.
12.5.5 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 SDS. 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
inner and outer jar labels (Figure 12.2 for UNK/RNG samples, and Figure 12.3 for QA voucher
samples). Attach the completed label to the jar and cover with clear shipping tape. Extra jar
labels are provided for each type of voucher collection. If you have multiple jars of either type of
sample, use the extra jar labels provided to prepare a label for each additional jar. For the QA
voucher sample, write the unique sample ID number on the extra jar label (this is found on the
pre-printed QA voucher label). On each jar label, use the spaces provided to record "Jar N of X",
where "N" is the individual jar number, and "X" is the total number of jars for the sample.
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UNK/RNG VOUCHER
Site ID: NRS23
/ / 202
UNK/RNG VOUCHER
Site ID: NRS23
/ / 202
UNK/RNG VOUCHER
BAG
TAG: 41
UNK/RNG VOUCHER
BAG
TAG: 42
UNK/RNG VOUCHER
BAG
TAG: 43
UNK/RNG VOUCHER
BAG
TAG: 44
UNK/RNG VOUCHER
BAG
TAG: 37
UNK/RNG VOUCHER
BAG
TAG: 38
UNK/RNG VOUCHER
BAG
TAG: 39
UNK/RNG VOUCHER
BAG
TAG: 40
UNK/RNG VOUCHER
BAG
TAG: 33
UNK/RNG VOUCHER
BAG
TAG: 34
UNK/RNG VOUCHER
BAG
TAG: 35
UNK/RNG VOUCHER
BAG
TAG: 36
UNK/RNG VOUCHER
BAG
TAG: 29
UNK/RNG VOUCHER
BAG
TAG: 30
UNK/RNG VOUCHER
BAG
TAG: 31
UNK/RNG VOUCHER
BAG
TAG: 32
UNK/RNG VOUCHER
BAG
TAG: 25
UNK/RNG VOUCHER
BAG
TAG: 26
UNK/RNG VOUCHER
BAG
TAG: 27
UNK/RNG VOUCHER
BAG
TAG: 28
UNK/RNG VOUCHER
BAG
TAG: 21
UNK/RNG VOUCHER
BAG
TAG: 22
UNK/RNG VOUCHER
BAG
TAG: 23
UNK/RNG VOUCHER
BAG
TAG: 24
UNK/RNG VOUCHER
BAG
TAG: 17
UNK/RNG VOUCHER
BAG
TAG: 18
UNK/RNG VOUCHER
BAG
TAG: 19
UNK/RNG VOUCHER
BAG
TAG: 20
UNK/RNG VOUCHER
BAG
TAG: 13
UNK/RNG VOUCHER
BAG
TAG: 14
UNK/RNG VOUCHER
BAG
TAG: 15
UNK/RNG VOUCHER
BAG
TAG: 16
UNK/RNG VOUCHER
BAG
TAG: 09
UNK/RNG VOUCHER
BAG
TAG: 10
UNK/RNG VOUCHER
BAG
TAG: 11
UNK/RNG VOUCHER
BAG
TAG: 12
UNK/RNG VOUCHER
BAG
TAG: 05
UNK/RNG VOUCHER
BAG
TAG: 06
UNK/RNG VOUCHER
BAG
TAG: 07
UNK/RNG VOUCHER
BAG
TAG: 08
UNK/RNG VOUCHER
BAG
TAG: 01
UNK/RNG VOUCHER
BAG
TAG: 02
UNK/RNG VOUCHER
BAG
TAG: 03
UNK/RNG VOUCHER
BAG
TAG: 04
Figure 12.2 Unknown/Range Extension Voucher Sample Labels and Tags
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12.5.6 Processing Unknown/Range Extension (UNK/RNG) Voucher Samples
Table 12.7 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 2023/24 laboratory operations manual. Identify
unknown specimens using whatever resources are necessary (magnification, literature,
reference collections/specimens, including dissected anatomical features or in-house
colleagues).
Following positive laboratory identification, update the Fish Collection Form in the NRSA Appto
reconcile the unknown records to reflect revisions to the actual species identifications, counts,
and any other information recorded on the Fish Collection Form. Resubmit the form to update
the database with the revised data.
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 Common Name field, and enter
any changes to the original counts in the appropriate Tally/Counts fields. If you determine that a
single unknown record is actually >1 species, update the original UNKNOWN record with the
information for the most abundant species. Record the information for additional species from
this original unknown in a new data line but retain the voucher specimen tag number of the
original unknown record. For example, if a sample of 20 specimens of UNKNOWN COTTUS A is
later identified as 15 individuals of one species and 5 individuals of another, record the common
name for the first (most abundant) species on the same line as the original unknown record, and
assign 75% of the original total count to it. Record the common name of the second species on
the first available blank line, and assign 25% of the original total count to this second species.
If you use a non-standard name (i.e., one that is not listed in Appendix D), use the adjacent
comment bubble to provide the taxonomic reference for the name. Submit the revised form via
the NRSA App as soon as possible after completing the laboratory identifications. Retain the
preserved UNK/RNG 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(a)glec.com. 715-829-3737). There are provisions under which fish can be identified by a
contracted lab and the results returned to you.
12.5.7 Processing QA Voucher Samples
Prepare the QA voucher sample as outlined in Table 12.8. Prepare the QA voucher sample
separately from the UNK/RNG voucher sample. Processing involves ensuring that the sample
jar(s) and photovoucher files include representative 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 number assigned (Figure 12.3).
Record information about the preserved QA voucher sample on the bottom of the Fish
Collection Form in the App.
Retain all of your QA voucher samples (including digital image files) until given direction by EPA
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
"dangerous goods," and packing and documentation requirements will differ depending on
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whether the samples contain formalin or ethanol, the size of individual bottles, and on the
particular shipping service used.
QA VOUCHER (VERT)
Site ID: NRS23
/ / 202
999012
QA VOUCHER (VERT)
Site ID: NRS23
/ / 202
999012
QA VOUCHER-BAG
TAG: 41
QA VOUCHER - BAG
TAG: 42
QA VOUCHER-BAG
TAG: 43
QA VOUCHER - BAG
TAG: 44
QA VOUCHER - BAG
TAG: 37
QA VOUCHER - BAG
TAG: 38
QA VOUCHER - BAG
TAG: 39
QA VOUCHER - BAG
TAG: 40
QA VOUCHER-BAG
TAG: 33
QA VOUCHER-BAG
TAG: 34
QA VOUCHER-BAG
TAG: 35
QA VOUCHER - BAG
TAG: 36
QA VOUCHER-BAG
TAG: 29
QA VOUCHER - BAG
TAG: 30
QA VOUCHER - BAG
TAG: 31
QA VOUCHER - BAG
TAG: 32
QA VOUCHER-BAG
TAG: 25
QA VOUCHER-BAG
TAG: 26
QA VOUCHER-BAG
TAG: 27
QA VOUCHER - BAG
TAG: 28
QA VOUCHER - BAG
TAG: 21
QA VOUCHER - BAG
TAG: 22
QA VOUCHER - BAG
TAG: 23
QA VOUCHER - BAG
TAG: 24
QA VOUCHER-BAG
TAG: 17
QA VOUCHER-BAG
TAG: 18
QA VOUCHER-BAG
TAG: 19
QA VOUCHER - BAG
TAG: 20
QA VOUCHER - BAG
TAG: 13
QA VOUCHER - BAG
TAG: 14
QA VOUCHER - BAG
TAG: 15
QA VOUCHER - BAG
TAG: 16
QA VOUCHER-BAG
TAG: 09
QA VOUCHER-BAG
TAG: 10
QA VOUCHER - BAG
TAG: 11
QA VOUCHER - BAG
TAG: 12
QA VOUCHER - BAG
TAG: 05
QA VOUCHER - BAG
TAG: 06
QA VOUCHER - BAG
TAG: 07
QA VOUCHER - BAG
TAG: 08
QA VOUCHER-BAG
TAG: 01
QA VOUCHER-BAG
TAG: 02
QA VOUCHER - BAG
TAG: 03
QA VOUCHER - BAG
TAG: 04
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Figure 12.3 QA Voucher Sample Labels and Tags
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Table 12.7 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) 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 voucher specimen tag number you recorded on the Fish Collection Form.
5) Make any corrections to the original collection data by revising names and/or counts in the Fish
Collection Form in the App.
a) Use common names from the standard list (Appendix D) as revised names.
b) If you must use a non-standard name, provide the taxonomic reference in the adjacent comment
bubble.
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 (the most abundant) on the line of the original
unknown.
b) Record the revised name and count for the second species as a new line of data on the Fish
Collection Form.
i) Record the specimen tag number from the original unknown record in the new line of data.
7) After reconciling all of the unknowns, and correcting any other information on the Fish Collection
Form, submit the revised form via the NRSA App to update the database with the new data. Retain the
preserved UNK/RNG voucher samples. Contact your regional EPA Regional Coordinator if you cannot
store the samples at your facility.
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Table 12.8 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 number 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 Contract 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
Tracking Form matches the preprinted label attached to each sample jar, and that the number of jars
recorded on the Fish Collection Form is correct. Also ensure that each extra jar is labelled with the
same Sample ID as the primary jar.
4) Complete an inner label by filling in the site ID, date, visit number and sample ID with a pencil. Place a
completed inner label inside each jar of specimens.
5) Retain the QA voucher samples in appropriate storage space for formalin until you receive information
regarding where to send them from the NRSA staff at EPA Office of Water or EPA Regional
Coordinator.
6) 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 Regional Coordinator if your
laboratory does not have the capability of handling waste formalin.
7) 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.
8) Decant the tap water. Use 45%-50% isopropyl alcohol or 70% ethanol as a final preservative for
specimens.
9) When ready to ship all of the QA voucher samples, complete a sample Tracking Form as described in
Appendix C.
10) Package the preserved samples properly for either formalin or ethanol and prepare all required
documentation and safety measures for the shipment.
11) Post all photovoucher files for each QA voucher sample to SharePoint. Use the file names that are
recorded on the Fish Collection Form.
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13 FISH TISSUE PLUG SAMPLING METHODS
13.1 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
for a single contaminant of concern (mercury), which uses a tissue plug instead of a skin-on
fillet. A plug sample consisting of 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 all sites where
suitable fish species and lengths are available. Specimens must be at least 190 mm to qualify for
tissue plug sampling. DO NOT collect tissue plugs from any fish less than 190 mm in total length.
These fish are often collected during the fish assemblage sample collection effort (Section 12)
but additional effort can be used to collect specimens for tissue samples if desired. 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. If only one qualifying specimen is collected, collect both tissue plugs from the same
individual.
13.2 Equipment and Supplies
Table 13.1 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 Collection Form in the NRSA App.
Table 13.1 Equipment and Supplies: Fish Tissue Plug Sample
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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
Sterile disposable scalpel
Sterile forceps
Laboratory pipette bulb.
Antibiotic salve.
Fish collection gear (electrofisher, nets, livewell,
etc.)
Dip net
Field Operations Manual
For recording measurements
Fish tissue plug sample labels
Fish Collection Form in NRSA App
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Fine-tipped indelible markers for filling out
sample labels
Clear tape strips for covering labels
13.3 Sample Collection Procedures
Collection of individual fish specimens for the fish tissue plug indicator occurs in the sample
reach during the fish assemblage sampling effort, using the same gear used to collect the fish
assemblage samples. Fish tissue plug samples should be taken from the species listed in the
target list found in Table 13.2. 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 plug sample. Recommended and
alternate target species are given in Table 13.2. The procedures for collecting and processing
fish plug samples are presented in Table 13.3.
If the fish assemblage sample collection effort does not yield specimens that are appropriate for
the collection of the tissue plugs, the crew should expend a reasonable amount of additional
effort to collect specimens for the tissue plug samples. This additional effort will not be included
as part of the fish assemblage effort and specimens collected during this additional effort should
not be recorded as part of the assemblage collection. Crews may target areas of habitat that
were not sampled as part of the fish assemblage effort or concentrate additional effort on
habitat likely to hold target fish.
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Table 13.2 Recommended Target and Alternate Species for Fish Tissue Plug Collection
FAMILY NAME
COMMON NAME
SCIENTIFIC NAME
MINIMUM LENGTH
GUIDELINE
Centrarchidae
Spotted bass
Micropterus punctulatus
The minimum length
Largemouth bass
Micropterus salmoides
guideline is >190 mm
Smallmouth bass
Micropterus dolomieu
for ALL species.
Black crappie
Pomoxis nigromaculatus
White crappie
Pomoxis annularis
Ictaluridae
Channel catfish
Ictalurus punctatus
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Blue catfish
Ictalurus furcatus
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Table 13.3 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 plug sample has been collected.
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 (minimum size for all species is 190 mm). Select fish based on the following criteria:
• is on the target list
• both the same species
• both satisfy legal requirements of harvestable size for the sampled river, or at least be of
consumable size if no legal harvest requirements are in effect
• 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 selected for plug sampling 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 the collection method, species, and specimen length and weight in the Fish Tissue Plug
section of the Fish Collection Form in the NRSA App. If the specimen is one of the fish species listed
in the assemblage portion of the form, it will appear in the drop-down list in the fish tissue plug
common name field. If the species was not collected as part of the assemblage effort, type the
common name directly into the box, taking care to spell it correctly.
9. Prepare a Sample Label for the sample with the Site ID, date and visit number. Affix the 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 offish 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.
If only one qualifying specimen is collected, collect both tissue plugs from the same individual (one plug
from each side of the fish). Leave the second set of length and weight fields blank and include a comment
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Fish Tissue Plug Methods
in the adjacent comment bubble to verify that both plugs were collected from the single fish listed and
the blank fields were left blank on purpose.
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 of collection.
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14 FINAL SITE ACTIVITIES
14.1 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, submit any and all data collected and ship or store the samples. Making an initial
data submission as soon as possible is important so that the data is stored in the EPA database,
preventing data loss due to a lost or broken iPad. Data can easily be updated in the NRSA App
and resubmitted at any time. Activities described in this section are summarized in Table 14.1.
Figure 14.1 Final Site Activities Summary
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14.2 General Site Assessment
Complete the Site Assessment Form in the NRSA App at the end of the sampling day, 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.
14.2.1 Elevation at Transect K
Ensure that the elevation at Transect K has been taken with your GPS and is recorded on the
Assessment Form. To record this information, record the elevation holding the GPS at
approximately 3 feet above the surface of the water.
14.2.2 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. 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 and be sure to verify that blank field indicate
absence by filling in the bubble at the top of the section. The distinction between low,
moderate, and heavy will be subjective. For example, if there are two to three houses on a site,
select "L" for low next to "Residences." If the site is lined 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.
To confirm that lines left blank were done so to indicate absence of the activity or disturbance,
select the "ALL activities were evaluated" bubble at the top of this section of the Site
Assessment Form in the App.
14.2.3 Site Characteristics
Record observations regarding the general characteristics of the site on the Site Assessment
Form. 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.
14.2.4 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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14.3 Processing the Fecal Indicator [Enterococci), Chlorophyll-a,
and Periphyton Samples
14.3.1 Equipment and Supplies (Fecal Indicator Filtering)
Table 14.1 provides the equipment and supplies needed for Field Crews to collect the fecal
indicator sample.
Table 14.1 Equipment and Supplies: Fecal Indicator (Enterococci) Sample Processing
For processing samples
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)
Millipore 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
Bubble bag
Zip-top bag
Dry ice
Cooler
Field Operations Manual
For recording measurements
Sample Collection Form in NRSA App
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
14.3.2 Procedures for Processing the Fecal Indicator (Enterococci) Sample
The fecal indicator sample must be filtered before the chlorophyll-a and periphyton samples,
since the filtering apparatus needs to be sterile for this sample. The procedures for processing
the fecal indicator sample are presented in Table 14.2. The sample must be filtered and frozen
within six hours of collection.
Table 14.2 Procedure: Fecal Indicator (Enterococci) Sample Processing
Filtering for the fecal indicator (Enterococci) 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, two bottles of chilled phosphate buffered saline
(PBS), Millipore 47 mm polycarbonate sterile filter box, and two filter forceps.
3. Chill Filter Extraction tubes with beads on dry ice.
4. Aseptically transfer two polycarbonate filters from filter box to base of opened Petri dish.
Close filter box and set aside (this step is to prevent wind from disturbing the remaining
filters in the box but is optional in calm/controlled environments).
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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.
6. Load filtration funnel with sterile polycarbonate filter on support pad (shiny side up).
7. Gently shake sample bottle 25 times to mix well.
8. Measure 25 mL of the mixed water sample in the sterile graduated sterile 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 vacuum [3.44 psig]). 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 mLand
continue filtration. If it was very difficult to filter the first 25 mL, proceed to step 11. If the
filter clogs before completely filtering the first or second 25 mL volume, discard the filter
and repeat the filtration using a lesser volume.
11. Pour approx. 10 mL of the chilled phosphate buffered saline (PBS) into the graduated PP
tube used for the sample. Cap the tube and shake 5 times. Remove the cap and pour
rinsate into filter funnel to rinse filter.
12. Filter the rinsate and repeat with another 10 mL of phosphate buffered saline (PBS).
13. Remove filter funnel from base without disturbing filter. Using sterile disposable forceps
remove the filter (touching only the filter edges) and fold it in half, in quarters, in eighths,
and then in sixteenths (filter will be folded four 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.
15. Record the volume of sample filtered through the filter on the small yellow label and
apply the label to the extraction tube (DO NOT cover with clear tape).
16. Record the volume of sample filtered through the filter on the outer bag label and apply
the label to the bubble bag (DO NOT cover with clear tape).
17. Insert tube into bubble bag and zip-top bag on dry ice for preservation during transport
and shipping.
18. Record the volume of water sample filtered through each filter on the Sample Collection
Form in the App. Record the filtration start time and finish time for the sample as well as
the time the filters were frozen.
19. Repeat steps 6 to 15 for the remaining 50 mL sub-sample volume to be filtered. Make
every effort to filter the same volume of sample through each of the two filters.
Processing Procedure—fecal indicator (Enterococci) filter blank
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Enterococci filter blanks will be prepared at all revisit sites during the first visit. Prepare the filter
blanks before filtering the stream 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 (this step is to prevent wind from disturbing the remaining filters in the
box but is optional in calm/controlled environments).
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 mL of the chilled phosphate buffered saline (PBS) in the sterile graduated PP tube
and pour into filter funnel.
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6. Replace cover on filter funnel and pump to generate a vacuum (do not generate more than 7
inches of Hg of vacuum [3.44 psig]). 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.
9. Record the volume of PBS filtered through the filter on the small yellow label and apply the
label to the extraction tube (DO NOT cover with clear tape). Note that there is a specific label
for the blank sample. At sites where a blank is not collected, this label will be discarded.
10. Insert tube into bubble bag and zip-top bag on dry ice for preservation during transport and
shipping.
11. Package and submit this sample to the lab with the standard samples.
12. Indicate that you have collected a filter blank by selecting the "Blank Collected" checkbox on
the Sample Collection Form.
14.3.3 Equipment and Supplies (Chlorophyll-a from Water Sample Filtering)
Table 14.3 provides the equipment and supplies needed to process the chlorophyll-a water
sample.
Table 14.3 Equipment and Supplies: Chlorophyll-a Processing
For filtering chlorophyll-a sample
Whatman GF/F 0.7 nm 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
Zip-top bag
Plastic electrical tape
For recording measurements
Sample Collection Form in NRSA App
Sample labels
Fine-tipped indelible markers
Clear tape strips
14.3.4 Procedures for Processing the Chlorophyll-a Water Sample w
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The procedures for processing chlorophyll-a water samples are presented in Table 14.4. >
Whenever possible, sample processing should be done in subdued light, out of direct sunlight. tj
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Table 14.4 Procedure: Chlorophyll-a Sample Processing
Filtering for the chlorophyll a water sample
1. Put on nitrile gloves.
2. Use clean forceps to place a Whatman GF/F 0.7 nm 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 gently shake the bottle to
homogenize the sample. While filtering sample, keep the bottle in the cooler on ice.
4. Measure 250 mL of 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 [3.44 psig]). 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.
5. Observe the filter for visible color. If there is readily 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 in the App.
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 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.
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 the provided leak-proof zip-top plastic bag
labelled with the completed chlorophyll outer bag label. Cover the outer label with clear tape.
Place this bag immediately on dry ice to freeze.
14.3.5 Equipment and Supplies (Periphyton Sample)
Table 14.5 lists the equipment and supplies needed to process the periphyton sample.
Table 14.5 Equipment and Supplies: Periphyton Samples
For preparing
Whatman 47 mm 0.7 micron GF/F glass fiber filter
periphyton samples
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
125 mL sterile PETG bottle
60 mL syringe with tip removed
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Aluminum foil squares
Forceps
Nitrile gloves
deionized water in wash bottle
plastic electrical tape
dry ice
wet ice
coolers
formalin
For data recording
Sample Collection Form
Sample labels
Fine-tipped indelible markers
Clear tape strips
For cleaning
10% Bleach solution
equipment
14.3.6 Procedures for Processing the Periphyton Samples
Four different types of laboratory samples are prepared from the composite periphyton sample:
an ID/enumeration sample (to determine taxonomic composition and relative abundances),
periphyton metagenomics sample, 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.
14.3.6.1 ID/Enumeration Sample
Prepare the ID/Enumeration sample as a 50 mL aliquot from the composite periphyton sample,
following the procedure presented in Table 14.6. Preserve each sample with 2 mL of formalin.
Record the sample ID number from the container label and the total volume of the periphyton
composite sample in the appropriate fields on the Sample Collection Form in the NRSA App.
Store the preserved samples upright in a container containing absorbent material.
Table 14.6 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 site
ID, date, and visit number as well as the total volume of the composite sample on the label. Attach
completed label to a 50 mL centrifuge tube. Cover the label completely with a clear tape strip.
2. Record the volume of the subsample (typically 50 mL) and the total volume of the composite
sample on the Sample Collection Form.
3. Thoroughly mix the bottle containing the composite sample.
4. Immediately after mixing, pour 50 mL of sample into pre-labeled 50 mL centrifuge tube.
5. 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.
6. Shake gently to distribute preservative.
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14.3.6.2 Periphyton Metagenomic Sample
Prepare the periphyton metagenomic sample as a 100 mL aliquot from the composite index
sample, following the procedure presented in Table 14.7.
Table 14.7 Procedure: Preparing Metagenomic Sample of Periphyton
Periphyton metagenomic Sample Processing Procedure
1. Prepare a sample label (with pre-printed sample ID number sample type "PDNA"). Record the
site ID, date, and visit number on the label. Attach completed label to the sterile 125 mL PETG
bottle. Cover the label completely with a clear tape strip.
2. Record the total volume of the composite sample on the Sample Collection Form.
3. Put on nitrile gloves.
4. Remove the cap from the bottle.
5. Do not rinse the bottle and avoid touching the inside of the bottle or the inside of the cap.
6. Thoroughly mix the bottle containing the composite sample and immediately pour 100 mL of the
mixed sample into the labeled 125 mL PETG bottle. Use the graduations on the bottle to gauge
the volume of sample poured.
7. Carefully replace the cap on the sample bottle. Seal the cap with plastic electrical tape.
8. Immediately after sample is collected, place in a cooler with ice to minimize exposure to light
and begin chilling the sample. The sample should be frozen as soon as is practicable and should
remain frozen until and during shipping.
14.3.6.3 Periphyton Chlorophyll a Sample
Prepare the periphyton chlorophyll-o sample by filtering a 25 mL aliquot of the composite
sample through a 47 mm 0.7 micron GF/F glass fiber filter. The procedure for preparing the
periphyton chlorophyll-o sample is presented in Table 14.8. Chlorophyll-o 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 (3.44 psig) 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.
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Table 14.8 Procedure: Preparing Chlorophyll Samples of Periphyton
Periphyton Chlorophyll a Sample Processing Procedure
3.
4.
5.
6.
7.
10.
11.
12.
13.
14.
Rinse the sides of the filter funnel and the filter with a small volume of deionized water to
prevent contamination from the previously filtered samples.
Using clean forceps, place a Whatman GF/F 0.7 nm 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
the filter properly. Attach the filter funnel to the filter holder and filter chamber, and then attach
the vacuum pump to the filter flask.
Rinse a 25 mL or 50 mL graduated cylinder three times with small volumes of deionized water
and discard.
Mix the composite sample bottle thoroughly.
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.
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
(3.44 psig) 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.
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.
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.
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.
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. Seal the cap with plastic electrical tape.
Prepare a sample label (with pre-printed sample ID number sample type "PCHL") including the
site ID, date, visit number, and volume filtered, and attach it to the centrifuge tube. Cover the
label completely with a strip of clear tape.
Place the centrifuge tube into the provided leak-proof zip-top plastic bag with the water column
chlorophyll sample.
Record the total volume of the composite 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.
Place the centrifuge tube containing the filter on dry ice.
14.3.6.4 Periphyton Biomass Sample
Prepare the periphyton biomass 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 the
biomass sample is presented in Table 14.9. Using the same filtration chamber that was used for
Enterococci and chlorophyll-o samples, rinse it with deionized water prior to filtering the
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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 (3.44 psig) 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 14.9 Procedure: Preparing Periphyton Biomass Sample
Periphyton AFDM Sample Processing Procedures
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1. 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.
2. 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.
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 (3.44
psig) 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. 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.
9. 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.
10. 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. Seal the cap with plastic electrical tape.
11. Prepare a sample label (with pre-printed sample ID number sample type "PBIO"), including the site
ID, date, visit number, and volume filtered, and attach it to the centrifuge tube. Cover the label
completely with a strip of clear tape. Place the centrifuge tube into the provided leak-proof zip-top
plastic bag with the water column and periphyton chlorophyll samples.
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12. Record the total volume of the composite 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.
13. Place the centrifuge tube containing the filter on dry ice.
14.3.6.5 Cleaning of Periphyton Equipment
Once all four laboratory samples have been prepared, discard any remaining sample and
thoroughly clean all periphyton sampling equipment (including brush, delimiter, composite
bottle, funnel, and syringe) with a 10% bleach solution to disinfect the equipment and limit the
possible spread of periphyton DNA to future samples. After cleaning, thoroughly rinse all the
equipment with tap or Dl water. Store the equipment in a clean plastic bag.
14.4 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.
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 each form, verify that all information has been
recorded accurately and that any data needing additional explanation has a comment
associated with it.
In each data form you will find a number of data validation routines that will help find missing or
possibly incorrect data. These routines can be accessed at any time by tapping the data review
button at the top of each form.
Ensure that all samples are labeled, all labels are completely filled in, and each label is covered
with clear plastic tape (with the exception of Enterococci labels). 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. Ensure that the water chemistry (CHEM) sample ID is entered into the top of the
Tracking Form to allow the App to populate the other IDs for samples that were collected.
14.5 Launch Site Cleanup
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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National Rivers and Streams Assessment 2023/24
Version 1.0, January 2023
Field Operations Manual
Wadeable
15 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
NRSA.
15.1 Revisit Sampling Overview
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 repeat
sampling design is provided in Figure 15.1.
Revisit Sites (4 per state)
r
I
Visit 1
Space revisit events a minimum of
2 weeks and a maximum of 1
month apart
:—i
Visit 2
Collect all samples:
In situ measurements
Antimicrobial resistance (AMR)
Water chemistry
Chlorophyll-a
Periphyton
Benthos
Enterococci
Fish assemblage
Fish plugs
Physical habitat
AMR Field
Blank
(perform before
collecting AMR
sample)
Enterococci
filter blank
(before filtering
other samples)
Collect all samples:
In situ measurements
Antimicrobial resistance (AMR)
Water chemistry
Chlorophyll-a
Periphyton
Benthos
Enterococci
Fish assemblage
Fish plugs
Physical habitat
Figure 15.1 Summary of the Repeat Sampling Design
15.2 Revisit Sampling Sites
A total of 192 (approximately 10%) of the target sites visited will be revisited during the same
sample year 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 likely wadeable and two likely non-wadeable sites. For each state these sites are:
Wadeable Revisit sites:
• The two likely wadeable revisit sites are resample sites from the NRSA 2018/19; one
from the Large Stream (LS) panel and one from the Small Stream (SS) panel. The revisit
sites are labeled as NRS23_18RVT2LS and NRS23_18RVT2SS, respectively, and are
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located in the Large and Small Streams tabs within the state specific site evaluation
spreadsheet.
Non-Wadeable Revisit sites:
• The two likely non-wadeable revisit sites are resample sites from the NRSA 2018/19
from the River (RV) panel. The revisit sites are labeled as NRS23_18RVT2RV_FT and are
located in the Rivers tab within the state specific site evaluation spreadsheet.
If a site selected for revisit sampling is dropped, then the next available base site in the same
state and panel should be revisited and the next available oversample site in the state and panel
is used to replace the re-designated base site. If there are no base sites remaining in the panel,
replace the revisit site with the first available oversample site in the panel and consider it the
new 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 (Figure 15.1). Label
the data and samples Visit 2 to indicate that they are from the second sampling event at a revisit
site.
At each revisit site, a field blank will be collected for the antimicrobial resistance sample during
the first sampling visit (Visit 1). The crews will pour a provided bottle of ultra-pure water into a
500mL PETG pre-sterilized bottle. Detailed description of the AMR field blank is found in Section
5.4.
Additionally 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 it with the "filter blank" label and use the provided checkbox on the Sample
Collection Form to indicate the blank was collected. The filter blank should be run before the
stream sample is filtered and the filter blank will be sent to the lab along with the stream
samples. Detailed description of the filter blank is found in Table 14.2.
15.3 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 QAPP for the NRSA. The following sections will
focus only on the field evaluation and assistance visits.
These visits provide a QA/QC check for the uniform evaluation of the data collection methods,
and an opportunity to conduct procedural reviews as required 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).
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15.3.1 Specifications for QC Assurance Field Assistance Visits
Field evaluation and assistance personnel are trained in the specific data collection methods
detailed in this FOM. 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 15.1 summarizes the plan, the checklist, and corrective action procedures.
It is anticipated that evaluation and assistance visits will be conducted with each Field Crew
early in the sampling and data collection process, and that corrective actions will be conducted
in real time. If the Field Crew misses or incorrectly performs a procedure, the Evaluator will note
this on the checklist and immediately point this out so the mistake can be corrected on the spot.
The role of the Evaluator is to provide additional training and guidance so that the procedures
are being performed consistent with the FOM, all data are recorded correctly, and paperwork is
properly completed at the site.
Table 15.1 General Information Noted During Field Evaluation
Field
Evaluation
Plan
• EPA 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.
o 15.4 Reporting
I—
g 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 Field Crew understands the findings and
CI will be able to perform the procedures properly in the future. The Evaluator will record
9 responses or concerns, if any, on the Field Evaluation and Assistance Check List. After the
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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:
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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