National Rivers and Streams Assessment 2023/24
Field Operations Manual
Version 1.0, January 2023
oEPA
Non-Wadeable
United States Environmental Protection Agency
Office of Water
Washington, DC
EPA-841-B-22-007
National Rivers and Streams
Assessment 2023/24
Field Operations
Manual
Non-Wadeable
Version 1.0
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National Rivers and Streams Assessment 2023/24
Version 1.0, January 2023
Field Operations Manual
Non-Wadeable
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 (FOMJ) 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, 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. 2023. National Rivers and Streams Assessment 2023/24: Field Operations Manual - Non-
Wadeable. EPA-841-B-22-007. U.S. Environmental Protection Agency, Office of Water
Washington, DC.
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National Rivers and Streams Assessment 2023/24
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NOTICE II
TABLE OF CONTENTS IV
LIST OF FIGURES VII
LIST OF TABLES VIII
ACRONYMS/ABBREVIATIONS X
DISTRIBUTION LIST XII
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 NON-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 14
2.3.1 General Considerations 14
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 Forms 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 an d Streams 30
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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 37
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 44
9 PERIPHYTON 46
9.1 Summary of Method 46
9.2 Equipment andSupplies 46
9.3 Sampling Procedure 46
9.4 Sample Processing in the Field 48
10 PHYSICAL HABITAT CHARACTERIZATION 49
10.1 Equipment andSupplies 49
10.2 Summary of Methods Approach 49
10.3 Components of the Field Habitat Assessment 50
10.4 Summary of Workflow 51
10.5 Work Flow and Reach Marking 52
10.5.1 Reconnaissance for Physical Habitat Data Collection 53
10.5.2 Thalweg Profile 54
10.6 Channel Margin ("Littoral") and Riparian Measurements 58
10.6.1 Channel Margin Depth and Substrate 58
10.6.2 Large Woody Debris 59
10.6.3 Bank Angle and Channel Cross-Section Morphology 61
10.7 Visual Riparian Estimates 66
10.7.1 Riparian Vegetation Structure 66
10.8 Instream Fish Cover, Algae, and Aquatic Macrophytes 68
10.9 Human Influences 69
10.10 Canopy Cover Measurements 71
10.11 Channel Constraint, Debris Torrents, and Recent Floods 72
10.11.1 Channel Constraint 72
10.11.2 Debris Torrents and Recent Major Floods 74
10.12 Elevation atTransect K 75 ^
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11.1 Summary of Method 76
11.2 Equipment andSupplies 76
11.3 Sampling Procedure 76
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11.4 Sample Processing in the Field 77
12 FISH ASSEMBLAGE 78
12.1 Summary of Method 78
12.2 Equipment andSupplies 78
12.3 Sampling Procedures 79
12.3.1 Irruptive Species 80
12.3.2 Secondary Electrofishing 85
12.3.3 Secondary Seining 87
12.4 Processing Fish 91
12.4.1 Identification and Tallying 91
12.4.2 Unknown Specimens 91
12.4.3 Photovouchering 97
12.4.4 Preparing Preserved Voucher Specimen Samples 97
12.4.5 Preserving Voucher Specimen Samples 98
12.4.6 Processing Unknown/Range Extension (UNK/RNG) Voucher Samples 100
12.4.7 Processing QA Voucher Samples 100
13 FISH TISSUE PLUG SAMPLING METHOD 104
13.1 Method Summary 104
13.2 Equipment andSupplies 104
13.3 Sample Collection Procedures 105
14 FINAL SITE ACTIVITIES 109
14.1 Overview of Final Site Activities 109
14.2 General Site Assessment 110
14.2.1 Elevation at Transect K 110
14.2.2 Watershed Activities and Disturbances Observed 110
14.2.3 Site Characteristics 110
14.2.4 General Assessment 110
14.3 Processing the Fecal Indicator (Enterococc\), Chlorophyll-^, and Periphyton Samples Ill
14.3.1 Equipment and Supplies (Fecal Indicator Filtering) Ill
14.3.2 Procedures for Processing the Fecal Indicator (Enterococci) Sample Ill
14.3.3 Equipment and Supplies (Chlorophyll-a from Water Sample Filtering) 113
14.3.4 Procedures for Processing the Chlorophyll-a Water Sample 113
14.3.5 Equipment and Supplies (Periphyton Sample) 114
14.3.6 Procedures for Processing the Periphyton Samples 115
14.4 Data Forms and Sample Inspection 119
14.5 Launch Site Cleanup 119
15 FIELD QUALITY CONTROL 120
15.1 Revisit Sampling Overview 120
15.2 Revisit Sampling Sites 120
15.3 Field Evaluation and Assistance Visits 121
15.3.1 Specifications for QC Assurance Field Assistance Visits 122
15.4 Reporting 122
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ฃ APPENDIX B SAMPLE FORMS B-l
^ APPENDIX C SHIPPING AND TRACKING GUIDELINES C-l
< APPENDIX D COMMON & SCIENTIFIC NAMES OF FISHES OF THE UNITED STATES D-l
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APPENDIX E EXAMPLE ELECTROFISHING SETTINGS E-l
Figure 1.1 Example Sample Labels for Sample Tracking and Identification 7
Figure 2.1 Field Sampling Scenario for Non-Wadeable Sites 9
Figure 2.2 Overview of Base Site Activities 10
Figure 2.3 Post-sampling Equipment Care 14
Figure 2.4. Example Sample Packing Slips to Accompany Sample Shipments 19
Figure 2.5 Electronic Request Form 20
Figure 3.1 Sampling Reach Features (Non-Wadeable Sites) 28
Figure 8.1 Benthic Macroinvertebrate Collection at Non-Wadeable Sites 41
Figure 8.2 Transect Sample Design for Collecting Benthic Macroinvertebrates (Non-Wadeable Sites) 42
Figure 10.1 Littoral Riparian Plots for Characterizing Riparian Vegetation, human influences, fish cover,
littoral substrate, and littoral depths 54
Figure 10.2 Riparian Zone and Instream Fish Cover Plots for a River Cross-Section Transect 58
Figure 10.3. Large Woody Debris Section of Physical Habitat Form in the NRSA App - Non-Wadeable 60
Figure 10.4 Schematic Showing Bankfull Channel and Incision for Channels 65
Figure 10.5 Determining Bankfull and Incision Heights for 66
Figure 10.6 Proximity Classes for Human Influences in Non-Wadeable Rivers 70
Figure 10.7 Schematic of Modified Convex Spherical Canopy Densiometer 71
Figure 10.8 Types of Multiple Channel Patterns 74
Figure 12.1 Reach Layouts for Fish Sampling at Non-Wadeable Sites 82
Figure 12.2 Unknown/Range Extension Voucher Sample Labels and Tags 99
Figure 12.3 QA Voucher Sample Labels and Tags 101
Figure 14.1 Final Site Activities Summary 109
Figure 15.1 Summary of the Revisit Sampling Design 120
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Table 1.1 Summary Table of Indicators for all NRSA 2023-2024 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 13
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: Site Verification 23
Table 3.2 Procedure: Site Verification 24
Table 3.3 Guidelines to Determine the Influence of Rain Events 26
Table 3.4 Procedure: Laying Out the Sampling Reach at Non-Wadeable Sites 27
Table 3.5 Procedure: Sliding the Sampling Reach 29
Table 3.6 Reach Layout Modifications for Interrupted Streams 30
Table 3.7 Procedure: Modifications for Braided Rivers and Streams 31
Table 4.1 Equipment and Supplies: DO, pH, Temperature, and Conductivity 32
Table 4.2 Procedure: Temperature, pH, Conductivity and Dissolved Oxygen 33
Table 5.1 Equipment and Supplies: Antimicrobial Resistance Samples 34
Table 5.2 Procedure: Antimicrobial Resistance Sample Collection (Non-Wadeable Sites) 34
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 (Non-Wadeable Sites) 36
Table 7.1 Equipment and Supplies: Microcystin Sample 38
Table 7.2 Procedure: Algal Toxin (Microcystin and Cylindrospermopsin) Collection (Non-Wadeable Sites)38
Table 8.1 Equipment and Supplies: Benthic Macroinvertebrate Collection at (Non Wadeable Sites) 40
Table 8.2 Procedure: Benthic Macroinvertebrate Sampling (Non-Wadeable Sites) 43
Table 8.3 Procedure: Compositing Samples for Benthic Macroinvertebrates (Non-Wadeable Sites) 44
Table 9.1 Equipment and Supplies: Periphyton (Non-Wadeable Sites) 46
Table 9.2 Procedure: Collecting Composite Index Samples of Periphyton (Non-Wadeable Sites) 47
Table 10.1 Equipment and Supplies: Physical Habitat 49
Table 10.2 Components of Non-Wadeable River Physical Habitat Protocol 50
Table 10.3 Summary of Workflow Physical Habitat Characterization (Non-Wadeable) 51
Table 10.4 Procedure: Thalweg Profile 55
Table 10.5 Channel Unit Categories Used on Thalweg Form 57
Table 10.6 Procedure: Channel Margin Depth and Substrate 59
Table 10.7 Procedure: Tallying Large Woody Debris 61
Table 10.8 Procedure: Bank Angle and Channel Cross-Section 62
Table 10.9 Procedure: Characterizing Riparian Vegetation Structure 67
Table 10.10 Procedure: Estimating Fish Cover 69
Table 10.11 Procedure: Estimating Human Influence 70
Table 10.12 Procedure: Canopy Cover Measurements 72
Table 10.13 Procedure: Assessing Channel Constraint 73
Table 11.1 Equipment and Supplies: Fecal Indicator Sampling (Non-Wadeable Sites) 76
Table 11.2 Procedure: Fecal Indicator (Enterococci) Sample Collection (Non-Wadeable Sites) 76
Table 12.1 Equipment and Supplies: Fish Sampling (Non-Wadeable Sites) 78
Table 12.2 Summary of Non-wadeable Fishing Protocols 80
Table 12.3 Procedure: Electrofishing (Non-Wadeable Sites) 83
i/i Table 12.4 Procedure: Secondary Electrofishing Methods for Wadeable Areas (Non-Wadeable Rivers)... 86
^ Table 12.5 Procedure: Secondary Seining Methods for Wadeable Areas (Non-Wadeable Rivers) 88
^ Table 12.6 Procedure: Processing Fish (Non-Wadeable Sites) 93
^ Table 12.7 Procedure: Processing Unknown/Range Extension (UNK/RNG) Voucher Samples 102
i_ Table 12.8 Procedure: Processing QA Voucher Samples 103
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Table 13.2 Recommended Target and Alternate Species for Fish Tissue Plug Collection 106
Table 13.3 Procedure: Fish Tissue Plug Samples 107
Table 14.1 Equipment and Supplies: Fecal Indicator (Enterococci) Sample Ill
Table 14.2 Procedure: Processing Fecal Indicator (Enterococci) Sample Ill
Table 14.3 Equipment and Supplies: Chlorophyll-o Processing 113
Table 14.4 Procedure: Chlorophyll-a Sample Processing 114
Table 14.5 Equipment and Supplies: Periphyton Samples 114
Table 14.6 Procedure: ID/Enumeration Samples of Periphyton 115
Table 14.7 Procedure: Preparing Metagenomic Sample of Periphyton 116
Table 14.8 Procedure: Preparing Chlorophyll Samples of Periphyton 117
Table 14.9 Procedure: Preparing Periphyton Biomass Sample 118
Table 15.1 General Information Noted During Field Evaluation 122
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AFDM Ash-Free Dry mass
AMR Antimicrobial resistance
ANC Acid-neutralizing capacity
Ca Calcium
CI Chloride
CN Cellulose nitrate
CPR Cardiopulmonary Resuscitation
CW Channel width(s)
CWA Clean Water Act
DBH Diameter at Breast Height
DELT Deformities, Eroded Fins, Lesions, and Tumors
Dl Deionized water
DO Dissolved Oxygen
DOC Dissolved Organic Carbon
EPA Environmental Protection Agency
FOM Field Operations Manual
GIS Geographic Information System
GPP Generator Powered Pulsator
GPS Global Positioning System
HAB Harmful Algal Bloom
HDPE High-density polyethylene
IBI Index of Biotic Integrity
IM Information Management
K Potassium
LOM Lab Operations Manual
LWD Large Woody Debris
Mg Magnesium
MMI Multimetric Index
MSDS Material Safety Data Sheets
Na Sodium
NAD North American Datum
NARS National Aquatic Resources Survey
NELAP National Environmental Laboratory Accreditation Program
NHD National Hydrology Database
NHs Ammonia
NH4 Ammonium
NIST National Institute of Standards and Technology
NO2 Nitrite
NO3 Nitrate
NRC National Research Council
NRSA National Rivers and Streams Assessment
O/E "Observed" over "Expected"
OSHA Occupational Safety and Health Administration
OW Office of Water
PBS phosphate-buffered saline
PC Personal Computer
PESD-AL Pacific Ecological System Division - Analytical Laboratory
PETG polyethylene terephthalate glycol
PFD Personal Flotation Device
PP Polypropylene
PPT Parts per thousand
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Psig pounds per square inch - gauge
PTD Percent Taxonomic Disagreement
QA Quality Assurance
QAPP Quality Assurance Project Plan
QA/QC Quality Assurance/Quality Control
QCS Quality Control Solution
QRG Quick Reference Guide
RVT Revisit
SEG Site Evaluation Guideline
Si02 Silica
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
USDA United States Department of Agriculture
UNK/RNG Unknown Range Extension
USGS United States Geological Survey
WT Water Tube
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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 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 the project contacts at
participating laboratories, as they are determined.
National Monitoring Coordinators
Richard Mitchell
NRSA Project Leader
mitchell.richard(5)eDa.gov
202-566-0644
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
Bernice L. Smith WRAPD OA
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.michelle(5)eDa.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
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Tom Faber
Region 1
faber. tomPepa. gov
617-918-8672
U.S. EPA - Region 1
11 Technology Drive, North Chelmsford, MA 01863-
2431
Emily Nering
Region 2
nering.emilvPepa.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.mariPeoa.gov
312-353-3017
U.S. EPA-Region V
77 West Jackson Blvd, Chicago, IL 60604-3507
Rob Cook
Region 6
cook, robe rtPeoa.gov
214-665-7141
U.S. EPA-Region VI
1445 Ross Ave -Ste 1200, Dallas, TX 75202-2733
Laura Webb
Region 7
webb.lauraPepa.gov
913-551-7435
U.S. EPA-Region VII
300 Minnesota Ave, Kansas City, KS 66101
Liz Rogers/Shera Reems
Region 8
rogers.lizPepa.gov /reems.sheraPepa.gov
303-312-6974/303-312-6888
U.S. EPA-Region VIII
1595 Wynkoop Street, Denver, CO 80202-1129
Matthew Bolt
Region 9
bo It.matthew Pepa.gov
415-972-3578
U.S. EPA-Region IX
75 Hawthorne Street San Francisco, CA 94105
Lillian Herger
Region 10
herger. lillian (5)eoa.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 Non-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 the 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 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), and
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 the end of September for most ecoregions.
o Sites in the select ecoregions or States can be sampled starting at 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-22-005) 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 conditions, 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 2023-2024 Sites
Indicator
Core or Supplemental
Indicator
Specs/Location in Sampling Reach
In Situ measurements (pH, DO,
temperature, conductivity)
Core Indicator
Measurements taken at Transect A at
mid-channel; readings are taken at 0.5 m
depth, or mid-depth if water depth is less
than 1 meter.
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 Transect A 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 Transect A 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
Core 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 and 0.3 m depth
Antimicrobial resistance (genes
and bacteria)
Supplemental Indicator
Collected at Transect A 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 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 information 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
(Adobeฎ 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.
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
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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).
It is imperative that field and sample information 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 Lead
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 NON-WADEABLE SAMPLING
2.1 Daily Operations
Field methods for the NRSA are designed to be completed in one field day for most sites.
Depending on the time needed for both the sampling and travel for the day, an additional day
may be needed to complete sampling or for pre-departure and post-sampling activities (e.g.,
cleaning equipment, repairing gear, shipping samples, and traveling to the next site). Remote
sites with lengthy or difficult approaches may require more time, and Field Crews will need to
plan accordingly.
A Field Crew for a non-wadeable site will typically consist of four or five people in two boats.
Additional crew members may either help collect samples or may remain on the bank to provide
logistical support. A minimum of two people is always required in a boat together to execute
the sampling activities and to ensure safety.
Typically, at non-wadeable sites, two crew members will work in the "habitat" boat, and two or
three will work in the "fish" boat. One crew member on each boat is primarily responsible for
boat operation and navigation.
A daily field sampling scenario showing how the 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 the Figure 2.1 cannot be changed and is based on the need to
protect some types of samples from potential contamination and to minimize holding times
once samples are collected. For example, at sites where physical habitat assessments are
expected to take longer than fish collections, Group B crew members may collect Group A Tasks
including in situ measurements and water collections. However, these water quality tasks will
need to be completed before fish collections begin.
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Whole Crew
Locate X site
Verify site as target
Set up staging area
Group A Activities:
Group B Activities:
Prepare labels, equipment & supplies
Calibrate multi- parameter probe
Group A or B
Load equipment and supplies onto boat(s)
Locate and travel to Transect A
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)
Conduct physical habitat
characterizations
Collect benthic samples
Conduct fish assessment
V S
Collect periphyton samples
Collect fecal indicator sample at last transect
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
Return to staging area
Clean and organize equipment for loading Inventory supplies for next sampling event
Request additional supplies if needed
Check and prepare all samples for transport or shipment
1
Inspect and clean boats, motors, and trailers to prevent transfer of
nuisance species and contaminants
i
Review App data forms for completeness
I
Submit data via the App / Ship samples / Submit App Tracking
Figure 2.1 Field Sampling Scenario for Non-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
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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 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
one extra site kit in the vehicle in the event replacement items are needed immediately.
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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 4 L 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
u 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
^ the Tracking Form in the App.
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< All samples need to be inspected and appropriately preserved and packaged for transport.
^ Check that all samples are labeled, and all labels are completely filled in. Check that each label is
ง 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
2 are properly sealed. Make sure that all sample containers are properly preserved for storage or
h immediate shipment.
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2.2.2.3 Equipment Cleanup and Check
o All equipment and gear must be cleaned and disinfected between sites to reduce the risk of
ง 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 clean off any plant and animal material. Prior to leaving a site, drain all bilge water
and live wells in the boat. Inspect, clean, and handpick plant and animal remains from vehicle,
boat, motor, and trailer. Before moving to the next site, if a commercial car wash facility is
available, wash 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.
Table 2.2 Post-sampling Equipment Care
Equipment Care after Sampling
1. Clean for biological contaminants.
Prior to departing site, drain all water from live wells and buckets used to hold and process fish, and
drain all bilge water from the boat.
Inspect motor, boat, trailer, sampling gear, waders, boots, etc. for evidence of mud, snails, plant
fragments, algae, animal remains, or debris, and remove using brushes or other tools.
At the base location, inspect and rinse seines, dip nets, kick nets, waders, and boots with water and
dry. Use one of the procedures below to disinfect gear if necessary.
Additional precautions to prevent transfer of Whirling Disease spores, New Zealand mudsnails, and
amphibian chytrid fungus 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.
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Equipment Care after Sampling
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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 409ฎ antibacterial
cleaner for at least 10 minutes, or soak gear in 120ฐF (49ฐC) water for at least 1 minute.
4. Dry gear in direct sunlight (at least 84 ฐF) for at least 4 hours.
Option B:
1. Soak gear in a solution of Sparquatฎ (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 120ฐF (49ฐC) water for at least 1 minute.
3. Dry gear in direct sunlight (at least 84 ฐF) 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 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).
Figure 2.3 Post-sampling Equipment Care
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
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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. If using a backpack shocker as a secondary sampling method, Field Crew
members using electrofishing equipment must be insulated from the water, boat, and
electrodes via rubber boots and linesman gloves. Ensure that the backpack electrofishing
equipment is equipped with a 45ฐ tilt switch that interrupts the current. All personnel should
use chest waders with nonslip soles and linesman gloves. DO NOT wear breathable waders
while electrofishing. If waders become wet inside, stop fishing until they are thoroughly dry or
use a dry pair. Avoid contact with the anode and cathode at all times due to the potential shock
hazard. If you perspire heavily, wear polypropylene or some other wicking and insulating
clothing instead of cotton. If it is necessary for a crew member to reach into the water to pick up
a fish or something that has been dropped, do so only after the electrical current is off and the
anode is removed from the water. Do not resume electrofishing until all individuals are clear of
the electroshock hazard. 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 member has ^
the responsibility to question and modify an operation or decline participation if it is unsafe. ^
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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.
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^ Many hazards lie out of sight in the bottoms of rivers and streams. Broken glass or sharp pieces
1/1 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.
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^ wastes that carry causative agents or pathogens must be immunized against tetanus, hepatitis,
O typhoid fever, and polio. Biological wastes can also be a threat in the form of viruses, bacteria,
q rickettsia, fungi, or parasites,
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t Appropriate safety apparel such as waders, linesman gloves, safety glasses, etc. must be
g available and used when necessary. First aid kits, fire extinguishers, and blankets must be readily
oe available in the field. Cellular or satellite telephones and/or portable radios should be provided
? to Field Crews working in remote areas in case of an emergency. Supplies (e.g., clean water,
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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. ^
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Table 2.4 General Safety Guidelines for Field Operations
ฆ 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.
ฆ 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.
ฆ Use extreme care walking on riprap. Rocks can shift unexpectedly and serious falls are possible.
ฆ 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.
ฆ Electrofishing units may deliver a fatal electrical shock, and should only be used by qualified,
experienced operators.
ฆ 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.
ฆ 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.
ฆ All electrical equipment must bear the approval seal of Underwriters Laboratories and must be
properly grounded to protect against electric shock.
ฆ Use heavy gloves when hands are used to agitate the substrate during collection of benthic
macroinvertebrate samples.
ฆ Use appropriate protective equipment (e.g., gloves, safety glasses) when handling and using
hazardous chemicals.
ฆ 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 venomous snake.
ฆ 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.
ฆ 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.
ฆ 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
10ฐC or 50ฐF) if he or she is exposed to wind or becomes wet.
ฆ 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.
ฆ Handle and dispose of chemical wastes properly. Do not dispose any chemicals in the field.
^ 2.4 Forms (NRSAApp)
^ 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
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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
P specifics of data entry is contained in Section 1.6.
O Electronic Field Forms: This form of data collection can be collected through an Apple
b iOS portable electronic device (tablets). The field forms will be optimized for tablet
q devices. The App will be preloaded on all EPA-owned iPads prior to distribution to Field
ง 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 Forms 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 2.4) 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_ Vlslt#:Q102
Date Collected: f J
Sample ID
Lab Comments (For lab staff only)
CHEM
999000
WCHL
999001
PBIO
999002
PCHL
999003
For Lab Staff only:
Rec'd by:
Date Rec'd:
T2: NRSA23/24 Daily Bacteria Sample Tracking
Site ID: NRS23
Visit #: Q 1 O 2
Date Collected:
/ /
Lab Staff:
Samples In Box? Sample ID Lab Comments (For lab staff only)
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Blank included
in shipment
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T3: NRSA23/24 Frozen Batched Tracking
Site ID: NRS23_ Visit #: O 1 0 2
Date Collected:
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T4: NRSA23/24 Non-Chilled Batched Tracking
Site ID: NRS23_ Visit #: Q 1 O 2
Date Collected:
J. J.
Lab Staff:
Samples In Box? Sample ID Lab Comments ( For lab staff only)
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ENTE 999013
PDNA
999005
MICX
999006
m-o 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 2.4. 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 2.5). 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 before 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 ail 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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SI ale:
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:
Comments:
O Non-chilled batch cooler (includes 1 cooler, T4/T5 shipping label and cooler liner)
How many? Need by: / /
O Replacement Label Packet(s)
How many? Need by: / /
Comments:
O Foil squares (aluminum) - pack of 25
Howmany? Need by: / / Comments:
OIL Benthic/Fish Voucher Bottles (HDPE, white, wide-mouth)
Howmany? Need by: / / Comments:
O Sodium Thiosulfate Tablets - vial of 25
Howmany? Need by: /
O Tape strips - packs of 25
Howmany? Need by:
Comments:
Don't see your item listed above? List items separately below. Refer to equipment lists in FOM for 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 2.5 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: 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
measurements
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).
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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
event, do not sample the site that day. ฃ
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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
wetted width is necessary to characterize the habitat and several biotic assemblages
associated with the sampling reach. Establish the sampling reach about the X-site using the
procedures described in Table 3.4 (non-wadeable sites). It is highly recommended that you lay
out the sampling reach for large, non-wadeable sites before you go in the field using maps,
aerial photos, and/or Geographic Information System (GIS) software. This will save time on the
field day.
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 non-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
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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.
Table 3.4 Procedure: Laying Out the Sampling Reach at Non-Wadeable Sites
Laying out the sampling reach at the base site (recommended at non-wadeable sites)
1. Using GIS, an aerial photo or a 1:100:000 topographic map; locate the X-site using the
coordinates provided for the site.
2. Determine the average wetted width of the channel at the X-site using GIS, available maps,
and/or aerial photographs. To get an average, 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 5 readings together and round to the nearest 1 m.
3. Multiply the average wetted width by 40 to determine the reach length. If the average width is
<4 m, use 150 m as a minimum reach length. If the average width is >100 m, use 4 km as a
maximum reach length. Record both the average channel width and total intended reach length
on the Verification Form in the App
4. From the X-site, measure a distance of one-half the reach length downstream using GIS. Be
careful to measure all of the bends of the river/stream; do not artificially straighten out the line of
measurement. The downstream endpoint is marked as Transect K. Measure one-half the reach
length upstream from the X-site; the upstream end of the reach is marked as Transect A. Record
the upstream and downstream reach lengths on the Verification Form in the App.
5. Measure 1/10 of the reach length downstream from Transect A, and mark this spot as Transect
B. Continue marking the 11 transects A - K in increments of 1/10 of the reach length. Enter the
waypoints for transects into a GPS unit so transects are easy to find on the sampling day.
6. Assign the sampling station at Transect A randomly (e.g., use the seconds display on a digital
watch or other random number selection method to select the initial sampling station: 0 - 4 = Left
Bank, 5 - 9 = Right Bank). From here, three stations will be on the first (randomly selected) side of
the river, then two on the other, then two on the first side, and so on through Transect K (as
shown in Figure 3.1). Note that left and right sides of the stream are determined while you are
facing downstream. It is at these locations that you will collect benthic macroinvertebrate and
periphyton samples.
7. When you are at the site, "ground truth" the wetted width measurements and proceed to Table
3.5 to see if the layout needs to be adjusted.
Laying out the sampling reach in the field
1. Locate the X-site using the coordinates provided for the site.
2. Use a laser range finder to determine the wetted width of the channel at 5 places of "typical"
width within approximately 5 channel widths upstream and downstream from the X-site. Average
the 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 for specific instructions.
Record the upstream and downstream reach lengths on the Verification Form in the App.
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9.
10.
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 upstream and downstream using a GPS
unit, laser rangefinder, or tape measure. Be careful to measure all of the bends of the
river/stream; do not artificially straighten out the line of measurement. Enter the channel to make
measurements only when necessary to avoid disturbing the stream channel prior to sampling
activities. The downstream endpoint is flagged as Transect K. The upstream end of the reach is
flagged as Transect A.
At Transect A, use the seconds display on a digital watch or other random number selection
method to select the initial sampling station for transect samples: 0 - 4 = Left Bank, 5 - 9 = Right
Bank. Mark "L" or "R" on the transect flagging. Note that left and right sides of the stream are
determined while you are facing downstream. It is at these locations that you will collect benthic
macroinvertebrate and periphyton samples.
Measure 1/10 of the reach length downstream from Transect A. Flag this spot as Transect B.
Assign the sampling station systematically after the first random selection as shown in Figure 3.1.
Three stations will be on the first side of the river, then two on the other, then two on the first
side, and so on through Transect K.
Proceed downstream with a GPS unit, laser rangefinder, or tape measure and flag the positions of
nine additional transects (labeled "C" through "K" as you move downstream) at intervals equal to
1/10 of the reach length. Continue to assign the sampling stations systematically.
Total reach length = 40 x mean wetted width (min = 150m; max = 4km)
QC
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Figure 3.1 Sampling Reach Features (Non-Wadeable Sites)
3.2.1 Sliding the Reach
There are some conditions that may require sliding the reach about the X-site (i.e., the X-site is
no longer located at the midpoint of the reach) to avoid features we do not wish to or physically
cannot sample across. Reasons for sliding the reach include:
Lack of landowner permission.
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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.
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 Transect A (even if the
reach has been adjusted by "sliding" it). If Transect A is dry and there is water elsewhere in the
sample reach, collect the sample from a location having water with a surface area >1 m2 and a u
depth >10 cm. ง
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Collect data for the physical habitat indicators along the entire sample reach from interrupted h
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
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percent of length with intermittent pools or no water). Other measurements associated with
characterizing riparian condition, substrate type, etc., are useful to help infer conditions in the
stream when water is flowing.
Table 3.6 Reach Layout Modifications for Interrupted Streams
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3.3.2 Braided Rivers and Streams
Depending upon the geographic area and/or the time of the sampling visit, you may encounter a
stream having "braided" channels, which are characterized by numerous sub-channels that are
generally small and short, often with no obvious dominant channel. If you encounter a braided
stream, establish the sampling reach using the procedures presented in Table 3.7. Figuring the
mean width of extensively braided rivers and streams for purposes of setting up the sampling
reach length is challenging. For braided channels, measure the mean width and bankfull width
as defined in the physical habitat protocols (Section 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 and sketches on the Verification Form in the NRSA App about what you did. Also add
additional details to the optional site map and submit 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
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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 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.
ฆ 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.
ฆ 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.
ฆ 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 and sketches 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 Transect A. Take the
readings at 0.5 m depth. Measure the site depth accurately before taking the measurements. If
the depth at Transect A 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 Multi-parameter water quality meter with pH, DO,
calibrating the water quality meter temperature, and conductivity probes.
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)
prior to launching the boat. 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.
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.
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
pH Meter
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Conductivity Meter
Calibrate the conductivity meter prior to each sampling event. Calibrate the meter in
accordance with the manufacturer's instructions. 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 Transect A, 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 25ฐC.
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 Transect A at the
midpoint of the river. 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 river 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 Transect A. 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
non-wadeable sites.
Table 5.2 Procedure: Antimicrobial Resistance Sample Collection (Non-Wadeable Sites)
Collection of Antimicrobial Resistance Samples
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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 river 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
Transect A at the midpoint of the reach. 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 Transect A.
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
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 describes the sampling procedures for collecting water chemistry samples in non-
wadeable streams and rivers.
Table 6.2 Procedure: Water Chemistry and Chlorophyll-a Sample Collection (Non-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 Transect A in a flowing portion near the middle of the site.
3. Put on nitrile gloves. Make sure not to handle sunscreen or other chemical contaminants until
after the sample is collected 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
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
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.
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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 Transect A as planned, select the
Transect A 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)
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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). ง
HABs can produce toxins known as cyanotoxins, which can be harmful to humans and animals. ง
Microcystin and cylindrospermopsin are two cyanotoxins known to occur in the surface waters u
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.
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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. b
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7.1 Summary of Method ^
The algal toxin (microcystin and cylindrospermopsin) sample is a grab samples taken from
Transect A. 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 h
collected. Store the sample on wet ice in a closed cooler. <
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7.2 Equipment and Supplies
Table 7.1 provides the equipment and supplies needed to collect the algal toxin sample at
Transect A. Record the water sample collection and preservation data on the Sample Collection
Form in the NRSA App.
Table 7.1 Equipment and Supplies: Microcystin Sample
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
measurements
Sample Collection Form in NRSA App
Algal toxin sample label with pre-printed Sample ID
Clear tape strips
Fine tipped indelible markers for labels
7.3 Sampling Procedure
Table 7.2 presents step-by-step procedures for collecting the algal toxin (microcystin and
cylindrospermopsin) sample at non-wadeable sites.
Table 7.2 Procedure: Algal Toxin (Microcystin and Cylindrospermopsin) Collection (Non-Wadeable Sites)
Microcystin Sample Collection
1.
2.
3.
4.
5.
6.
8.
Fill out the pertinent information (Site ID, visit, and date) on the algal toxin label.
Affix the MICX label to the 500 mL PETG clear square Nalgene bottle. Completely cover the label
with clear tape.
Collect the algal toxin sample from Transect A in a flowing portion of the stream near the
middle of the transect.
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.
Rinse the 3 L Nalgene beaker three times with water, and discard the rinse downstream.
Rinse the water sample collection container and lid three times with water, discard the rinse
downstream.
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.
Seal the cap with plastic electrical tape before shipping.
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.
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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 comments 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 3.1). 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 back of Benthic Collection Form in the NRSA App.
Table 8.1 Equipment and Supplies: Benthic Macroinvertebrate Collection at (Non 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
measurements
without preprinted ID Sample ID numbers
Fine-tip indelible markers
Blank labels on waterproof paper for inside
Clear tape strips
of 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 non-wadeable sites. Collect benthic
macroinvertebrate samples at the 11 transects and within the sampling stations for non-
wadeable streams (Figure 8.2). The process for selecting the sample stations is described in the
Initial Site Procedures (Section 3). Collect all benthic samples at non-wadeable sites from the
dominant habitat type within the 10 m x 15 m designated sampling station at each transect.
The procedure for collecting samples at each transect is described in Table 8.2. Take a 1 linear
meter sweep from within the dominant habitat type. Record the benthic macroinvertebrate
collection data on the Benthic Collection Form in the NRSA App. As you proceed from transect
to transect, combine all samples into a bucket.
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Figure 8.1 Benthic Macroinvertebrate Collection at Non-Wadeable Sites ^
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Thoroughly rinse net and proceed to
the next sampling station
Composite the samples from all stations
to create a single sample for the site
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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
Modified D-frame kick net
1 linear meter sweep of dominant habitat
I
COMPOSITE SAMPLES FROM ALL TRANSECTS
Sieve bucket or other bucket(s)
SIEVE SAMPLE
500 [jm 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
concentration of at least 70%
1 L
Figure 8.2 Transect Sample Design for Collecting Benthic Macroinvertebrates (Non-Wadeable Sites)
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Table 8.2 Procedure: Benthic Macroinvertebrate Sampling (Non-Wadeable Sites)
Collecting the Benthic Macroinvertebrate Sample
1. After locating the sampling station at Transect A according to procedures described in the reach
layout section, identify the dominant and secondary habitat type within the plot:
ฆ Coarse (rocky/cobble/gravel/large woody debris) ฆ Organic Fine mud or sand
ฆ Macrophyte beds ฆ Leaf Pack
ฆ OTher (describe in comment bubble)
2. Use the D-frame dip net (equipped with 500 nm mesh) to sweep through 1 linear meter of the
most dominant habitat type within the 10m x 15m sampling station, making sure to disturb the
substrate enough to dislodge organisms.
a) If the dominant habitat is rocky/cobble/large woody debris it may be necessary to exit
the boat and disturb the substrate (e.g., overturn rocks, logs) using your feet while sweeping
the net through the disturbed area.
b) Because a dip-net is being used for sampling, the maximum depth for sampling will be
approximately 0.5 to 1 m; therefore, in cases in which the depth of the river quickly drops off
it may be necessary to sample in the nearest several meters to the shore.
3. After completing the 1 linear meter sweep, remove all organisms and debris from net and place
them in a bucket following sample processing procedures described in the following section.
4. Record the side of the river on which the sampling station is located (left or right, facing
downstream).
5. Record the sampled habitat type (dominant) as well as the secondary habitat type on the Benthic
Collection Form in the App.
6. Record the substrate of the sampled area:
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 bubble.
7. Identify the channel habitat type where the sampling station was located. Indicate the
appropriate channel habitat type for the transect on the Sample Collection Form. The channel
habitat types are:
Pool (P): Still water; low velocity; smooth, glassy surface; usually deep compared to other
parts of the channel
Glide (GL): Water moving slowly, with smooth, unbroken surface; low turbulence
Riffle (Rl): Water moving, with small ripples, waves, and eddies; waves not breaking, and
surface tension is not broken; "babbling" or "gurgling" sound.
Rapid (RA): Water movement is rapid and turbulent; surface with intermittent "white
water" with breaking waves; continuous rushing sound.
Other (OT): Note type of "other" channel habitat in comment bubble on field form.
8. Proceed to the next sampling station and repeat steps 1-7. Between each station remove any
predacious macroinvertebrates and preserve immediately to prevent loss of organisms. Also,
ensure that sample is kept wet between transects. The organisms and detritus collected at each
station on the river should be combined in a single bucket to create a single composite sample for
the river.
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9. If any stations could not be sampled for benthos, leave the habitat, 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.
10. Record the number of transects that were sampled throughout the reach.
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 (Non-Wadeable Sites)
Compositing Benthic Macroinvertebrate Sample
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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
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.
ฆ 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 Collector(s) initials
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Compositing Benthic Macroinvertebrate Sample
Type of sampler and mesh size used Number of stations sampled
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, then drain using the net (or sieve) across the opening to prevent
loss of organisms, and replace with ethanol at the vehicle.
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 electrical
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 near shore shallows at each of the sampling stations located on the
11 cross section transects ("A" through "K") established within the sampling reach. Collect
periphyton samples at the same transect location as the benthic macroinvertebrate samples
(Section 8), directly after the benthic macroinvertebrate samples have been collected. 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
metagenomics 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 (Non-Wadeable Sites)
For collecting samples
Large Funnel (15-20 cm diameter)
12 cm2 area delimiter (3.8 cm diameter pipe, 3 cm tall)
Stiff-bristle toothbrush with handle bent at 90ฐ angle
1 L wash bottle for Dl water
500 mL graduated plastic bottle for the composite sample with
marked volume gradations
60 mL plastic syringe with tip removed, and length of tubing (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 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 (left or right). Collect the substrate selected for sampling from a depth no deeper
than 0.5 m. If you cannot collect a sample because the entire location is too deep, skip the
transect. 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
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produce one composite sample for each site. Record the number of transects sampled and the
total volume of the composite sample on the Sample Collection Form in the NRSA App.
Table 9.2 Procedure: Collecting Composite Index Samples of Periphyton (Non-Wadeable Sites)
Periphyton Composite Sample
1) Starting with Transect "A", collect a single sample from the assigned sampling station using the
procedure below.
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.):
i) 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.
ii) 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 20mL in the tube equals the target volume of 45 mL.
iii) 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.
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.)
c) If no coarse sediments (cobbles or larger) are present:
i) Use the area delimiter to confine a 12 cm2 area of soft sediments in a shallow area of the
sampling station.
ii) 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 25mL line of the syringe.
iii) 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.
iv) Put the bottle in a cooler on ice while you travel between transects and collect the o
subsequent samples. (The sample needs to be kept cool and dark because a chlorophyll ^
sample will be filtered from the composite.) o.
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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
1.
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.
2.
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.
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 bubbles.
3.
Clean up
4.
After preparing the four types of laboratory samples (see Section 14.3), 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 samples: an
ID/enumeration sample (to determine taxonomic composition and relative abundances), a
metagenomic sample, 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 river 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
river. Use this checklist to ensure that equipment and supplies are organized and available at
the river site in order to conduct the activities efficiently.
Table 10.1 Equipment and Supplies: Physical Habitat
For making Convex spherical canopy densiometer (Lemmon Model B), modified with taped "V"
measurements gpg
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
SONAR unit
Laser rangefinder (400 ft. distance range) and clear waterproof bag
Clinometer
Binoculars (optional)
Surveyor's telescoping leveling rod
Sounding rod
Field Operations Manual
For recording
data
In the NRSA App:
Physical Habitat Form
Channel Constraint Form
Torrent Evidence Form
Site Assessment Form
10.2 Summary of Methods Approach
Physical habitat in rivers includes all those physical attributes that influence or provide
sustenance to organisms within the stream. The physical habitat of a river varies naturally, thus
expectations differ even in the absence of anthropogenic disturbance. River sample reach
lengths are defined as 40 times the mean wetted width near the X-site, with a minimum of 150
m and maximum of 4 km, as described in Section 3. Measurement points are systematically
placed to statistically represent the entire reach. River depth and wetted width are measured at
very tightly spaced intervals, whereas channel cross-section profiles, substrate, bank
characteristics and riparian vegetation structure are measured at larger intervals. Woody debris
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is tallied in 11 sampling locations placed evenly throughout the sampling reach and the presence
of snags is recorded along the thalweg between transects. 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.
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10.3 Components of the Field Habitat Assessment
Field data collection for the physical habitat assessment is accomplished in a single float down
each sampling reach. The physical habitat methods are made up of the following components:
Thalweg profile, Littoral/Riparian Cross-Sections, and assessments of the entire reach. Table
10.2 describes the components of physical habitat in non-wadeable systems and gives an
overview of how the data is collected. Physical habitat measurements are recorded using the
NARS App. Similarly, specific forms in the App are used for assessing the degree of channel
constraint and recording evidence of debris torrents or recent major flooding.
Table 10.2 Components of Non-Wadeable River Physical Habitat Protocol
1. Thalweg Profile:
At 10 equally spaced intervals between each of 11 transects (100 along entire reach):
Classify habitat type, record presence of backwater and off-channel habitats.
Determine dominant substrate visually or using sounding rod.
Record the presence of mid-channel snags
Measure thalweg (maximum) depth using Sonar or sounding rod
2. Littoral/Riparian Cross-Sections: at 11 transects at equal intervals along reach length:
Measure/estimate from one chosen bank on 11 transects:
Wetted width and Mid-channel bar width (laser range finder).
Bankfull width (laser) and height (pole and clinometer used as level).
Incision height (pole and clinometer used as level).
Bank angle (estimate)
Riparian canopy cover (densiometer) in four directions from chosen bank.
Shoreline Substrate in the first 1 m above waterline (dominant and subdominant size
class).
In 20 m long Littoral Plot extending streamward 10 m from chosen bank:1
Littoral depth at 5 locations systematically-spaced within plot (Sonar or sounding rod).
Dominant and Subdominant substrate size class at 5 systematically-spaced locations (visual
or sounding rod).
Tally large woody debris in littoral plot and in bankfull channel by size and length class.
Areal cover class of fish concealment and other features, including:
filamentous algae overhanging vegetation aquatic macrophytes
undercut banks large woody debris boulders and rock ledges
brush/small woody debris live trees or roots artificial structures
In 20 m long Riparian Plot extending 10 m landward starting at bankfull margin-both sides of
river:
Estimate areal cover class and type (e.g., woody) of riparian vegetation in Canopy,
Understory, and Ground Cover layers
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Observe and record human activities and disturbances and their proximity to the channel.
3. For the whole sampling reach, after completing thalweg and littoral/riparian measurements:
Classify channel type and degree of constraint, identify features causing constraint,
estimate the percentage of constrained channel margin for the whole reach, and estimate
the bankfull and valley widths.
*Note: Boundaries for visual observations are estimated by eye.
10.4 Summary of Workflow
Table 10.3 lists the activities performed at and between each transect for the physical habitat
characterization. The activities are performed along the chosen riverbank and mid-channel
(thalweg profile).
Table 10.3 Summary of Workflow Physical Habitat Characterization (Non-Wadeable)
A. At the chosen bank on first transect (farthest upstream):
1. Access the Physical Habitat Form in the NRSA App and select the button that indicates the transect
at which you are located. Select the bubble to indicate the bank (left or right) that was assigned
during reach layout.
2. Use a GPS to determine the coordinates where the transect intersects the chosen bank and record
them in the Bank Latitude and Longitude fields on the form.
3. Record the dominant and secondary substrate of the shoreline 1 meter above the water's edge,
based on visual observations.
4. Move boat in a "loop" within the 10 x 20 m littoral plot, measuring and recording 5 littoral depths
and probing substrate. Use the 'sonar' or 'pole' columns based on the method used.
5. Estimate dominant and secondary littoral substrate, based on probing the 5 locations wherever
possible. If probing is not possible due to depth, flow, or other conditions, mark the 'judgement'
bubble and use whatever evidence is available to assign values to the substrate.
6. Tally Large Woody Debris (LWD) within or partially within the 10 x 20-meter littoral plot that is all
or partially in the wetted channel.
7. Tally LWD within or partially within the 10 x 20-meter littoral plot that is entirely out of water but
at least partially within the bankfull channel.
Measure or estimate the wetted width and (if present) bar width, estimate bankfull height, width
and channel incision. (Note that width and incision estimates incorporate both left and right
banks.).
Choose bank angle class.
Make visual riparian vegetation cover estimates for the 10 x 20-meter riparian plot on both sides
of the channel. (Riparian plot starts where perennial vegetation begins or at bankfull channel
margin, whichever is closest to the wetted river margin. The plot continues 10 m back from the
bankfull line).
Make visual human disturbance tally on both sides of the river. Use the same plot dimensions as
for riparian vegetation - except that if a disturbance item is observed in the river or within the
bankfull channel, the proximity code is "B", use the close rating; "C" if disturbance is present
within the riparian plot. If the item is only observed beyond (outside) the riparian plot, the
proximity code is "P". If the disturbance is not present in or adjacent to the plot, mark as "0", not
present.
12. Estimate areal cover of fish concealment features in the 10 x 20-meter littoral plot.
13. Estimate and record distance to riparian vegetation on the chosen bank.
14. Indicate the level of channel constraint present in the area of the transect and indicate whether
you could readily see over the bank.
8.
9.
10.
11.
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17.
18.
15. Collect densiometer measurements at bank (facing upstream, downstream, left, right).
16. Proceed to a midstream point on the transect and record the GPS coordinates in the Midstream
Latitude and Longitude fields on the form.
Record the intended transect spacing in the field provided. This should equal l/10th the total reach
distance.
If the next transect has not already been marked during reach layout, get out far enough from the
bank so you can see downstream. Then use the laser rangefinder to sight and record the distance
to the intended position of the next downstream transect.
B. Thalweg Profile:
1. As soon as you get out from the bank after doing transect activities, take the first of 10 thalweg
depth measurements and substrate/snag probes using sonar and pole -- also classify habitat type
and record presence of side-channels and backwaters.
2. Estimate thalweg measurement distance increments using the GPS course-tracking and trip-
meter functions. Alternatively, estimate these distances by keeping track of boat lengths or
channel-width distances traversed; each thalweg measurement is l/10th the distance between
transects, which can help you keep track of your downstream progress).
3. After completing the thalweg measurements, record the actual transect spacing in the field
provided. In fast flowing water or rivers with obstructions or hazards, it may be necessary to place
one or more transects based on safe access. In these instances, the intended and actual transect
spacing values will be different. If the next transect is placed as intended, the two values will be
equal.
C. Repeat the Whole Process (for the remaining 10 transects and subreaches).
D. Channel Constraint Assessment
After completing the Thalweg Profile and Littoral-Riparian measurements and observations at all
11 Transects, complete the classification and estimation of channel constraint type, frequency of
contact with constraining features, and the width ratio of bankfull channel divided by valley
width. You may wish to refer to the individual transect assessments of incision and constraint.
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10.5 Work Flow and Reach Marking
In a single midstream float down the 40 channel-width reach, the 2-person habitat crew
accomplishes a reconnaissance, a sonar/pole depth profile, and a pole-drag to note the
presence of snags and characterize mid-channel substrate. The float is interrupted by stops at
11 transect locations for littoral/riparian observations. Crews determine (and mark - optional,
but recommended) the intended position of each successive downstream transect using a global
positioning system (GPS) and/or a laser range finder. The crew then floats downstream along
the thalweg to the new transect location, making thalweg profile measurements and
observations at 10 evenly-spaced increments along the way. When they reach the new
downstream transect location, they stop to perform cross-section, littoral, and riparian
measurements, recording the actual GPS latitude/longitude of the transect position (bank and
midstream). They will also collect biological samples at each transect.
GPS coordinates are determined for the actual locations of each transect stop. If the GPS unit
also has course tracking, trip-meter (accumulated distance and bearing), and waypoint
setting/navigation features, we recommend using it to locate thalweg measurement points
(use course tracking and trip meter). Equipping the boat with a bow or stern anchor to stop at
transect locations can greatly ease the shore marking operation and shoreline measurement
activities, though such equipment can be dangerous to use in some rivers.
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10.5.1 Reconnaissance for Physical Habitat Data Collection
The habitat crew will also record reconnaissance and safety notes at this time. They will inform
the second boat of the route, craft, and safety precautions needed during its subsequent
electrofishing activities. They also assist the electrofishing boat crew over log jams or other
obstacles and help to conduct shuttles (this can take considerable time where put-ins and take-
outs are distant). As the crew floats downstream, they may choose and communicate to the
electrofishing crew the most practical path to be used when fishing with a less maneuverable
boat, taking into consideration multiple channels, blind channels, backwaters, alcoves,
impassible riffles, rapids, jams, and hazards such as dams, bridges and power lines.
Measurements are made at two scales along the length of the reach; the results are later
aggregated for the entire reach. 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). The
thalweg profile measurements must be spaced evenly over the entire reach.
The sampling reach length is 40 times the wetted width at the X-site, with a 150 m minimum
and 4-kilometer maximum reach. Section 3.1.1 describes the procedures for locating the X-site,
or the midpoint of the sample reach. Section 3.2 describes the protocol for delineating a sample
reach length that is 40 times its width. Those sections also describe the protocol for measuring
out (with a laser range finder or GIS software) and locating the 11 littoral/riparian stations
where many habitat measurements will be made.
The thalweg profile measurements are spaced as evenly as practicable over the entire sample
reach length. They must also be sufficiently close together to not "miss" deep areas and habitat
units that are in a size range of about 1/3 to 1/2 of the average channel width. To set the
interval between thalweg profile measurements:
Divide the reach length by 100 to set the thalweg increment distance. You will be
making 100 evenly-spaced thalweg profile measurements, 10 between each detailed
channel cross-section where littoral/riparian observations are made (the 11 established
transect locations).
If the thalweg is too deep or it is not physically possible to measure the thalweg,
estimate the depth to the best of your ability and flag it on the field form.
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littoral substrate, and littoral depths
10.5.2 Thalweg Profile
"Thalweg" refers to the flow path of the deepest water in a river channel. The thalweg profile is
a longitudinal survey of maximum depth and several other selected characteristics at 100 near-
equally spaced points along the centerline of the river between the two ends of the river reach
(Figure 10.1). For practical reasons, Field Crews will approximate a thalweg profile by sounding
along the river course that they judge is deepest, but also safely navigable. Locations for
observations and measurements along the path of this profile are determined using the GPS
course-tracking and trip-meter features (recommended), or by visually estimating distances
based upon the river width. Data from the thalweg profile allows calculation of indices of
residual pool volume, river size, channel complexity, and the relative proportions of habitat
types such as riffles and pools. The procedure for obtaining thalweg profile measurements is
presented Table 10.4. Record data on the lower portion of the Physical Habitat Form in the
NRSA App.
10.5.2.1 Thalweg Depth Profile
A thalweg depth profile of the entire 40 channel-width reach is approximated by a sonar or
sounding rod while floating downstream along the deepest part of the channel (or closest
navigable path). In the absence of a recording fathometer (sonar depth sounder with strip-chart
output or electronic data recorder), the crew records depths at frequent, relatively evenly-
spaced downstream intervals while observing a sonar display and holding a surveyor's rod off
the side of the boat. The sonar screen is mounted so that the crew member can read depths on
the sonar and the rod at the same time. The sonar sensor may need to be mounted at the
opposite end of the boat to avoid mistaking the rod's echo for the bottom, though using a
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narrow beam (16 degree) sonar transducer minimizes this problem. It is easy to hold the
sounding rod vertically if you are going at the same speed as the water. If the thalweg is too
deep or the water is too rough to safely measure depth, estimate the depth to the best of your
ability and record descriptive notes in the adjacent comment bubble.
10.5.2.2 Snags and Substrate Characteristics
The procedure for identifying snags by either dragging the thalweg pole to detect underwater
snags or observing snags throughout the wetted channel and substrate characteristics are
presented in Table 10.4. While floating downstream, one crew member holds a calibrated PVC
sounding rod or surveying rod down vertically from the gunwale of the boat, dragging it lightly
on the bottom to simultaneously "feel" the substrate, detect snags, and measure depth with the
aid of sonar. The crew member shall record the dominant substrate type sensed by dragging the
rod along the bottom (bedrock/hardpan, boulder, cobble, gravel, sand, silt & finer, or other) on
the Thalweg Profile section of the Physical Habitat Form in the App. Substrate characteristics are
recorded at every thalweg depth measurement (e.g., 10 determinations between each
transect). In shallow, fast-water situations, where pole-dragging might be hazardous, crews will
estimate bottom conditions the best they can visually and by using paddles and oars. If
unavoidable, suspend measurements until out of Whitewater situations, but make notes and
appropriately flag observations concerning your best judgments of depth and substrate. In
addition to noting snags during the thalweg measurements, crew member will note the
presence of snags throughout the wetted channel.
10.5.2.3 Channel Habitat Classification
Classify and record channel habitat types shown in Table 10.5 and check for the presence of off-
channel and backwater habitat at a spatial resolution of about 0.4 channel-width increments.
Designate side channels, backwaters and other off-channel areas independent of the main-
channel habitat type. Main channel habitat units are at least half as long as the channel is wide,
(e.g., if there is a small, deep, pool-like area at the thalweg within a large riffle area, don't record
it as a pool unless it occupies an area about half as wide or long as the channel is wide). For dry
and intermittent rivers, record the channel habitat type as dry channel (DR) in places where no
water is present in the channel.
Table 10.4 Procedure: Thalweg Profile
Taking the Thalweg Profile
1. Record GPS coordinates (Lat/Long) midstream and at shoreline location on the Physical Habitat Form
at the transect located at the upstream end of the thalweg subreach about to be assessed.
2. Determine the interval between transects based on the mean wetted width used to determine the
reach length. Transects are at 4 channel-width spacing; thalweg depth, snags, off-channel habitats
and other downstream longitudinal profile observations are recorded at intervals of 0.4 channel-
widths.
3. Record the calculated interval in the intended transect spacing field.
4. Begin at the upstream transect (station "0" of "10"). Determine the locations at which to take
measurements using the course-tracking and trip-meter functions of the GPS. Alternatively, estimate
your position.
Thalweg Depth Profile
a) While floating downstream along the thalweg, record depths at frequent, evenly-spaced
intervals while observing a sonar display and holding a surveyor's rod off the side of the boat.
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b) A depth recording every 0.4 channel-width distance is required, yielding 10 measurements
between channel/riparian cross-section transects.
c) If the depth is >0.5 meters, or contains a lot of air bubbles, the sonar fathometer may not give
reliable depth estimates. In this case, record depths using a calibrated sounding rod. In shallow,
fast-water situations depths may have to be visually estimated to the nearest 0.5 m.
d) Measure depths to nearest 0.1 m and record in either the "SONAR" or "POLE" column based on
the method used. You may use different methods in the same thalweg profile, but each station
should only have a depth recorded in one column.
Snags and Substrate Characteristics
a) From the gunwale of the boat, hold a surveying rod or calibrated PVC sounding rod down vertically
into the water. (CAUTION: Hold the rod over the side or upstream end of the raft; otherwise it
could be jerked out of your hands if it catches on an obstruction in fast water.)
b) Lightly drag the rod on the river bottom to "feel" the substrate and detect snags.
c) Record the presence of any snags, whether they are hit by the rod or seen visually in the
thalweg or anywhere within the channel. Also record the dominant substrate type sensed by
dragging the rod along the bottom.
d) Choose the appropriate "SUBSTRATE" type and record the presence/absence of "SNAGS" using
the provided checkboxes (checked boxes indicate presence, unfilled boxes indicate absence). If
snags are encountered between stations, mark snags as present on the line for the next station
assessed.
e) If it is too deep to safely measure the substrate type, estimate the type based on knowledge
and surrounding measurements and provide a comment in the adjacent comment bubble.
Channel Habitat Classification
a) Classify and record the channel habitat type at increments of every 0.4 channel width.
b) Check for off-channel and backwater habitat at increments of every 0.4 channel width.
c) If channel is split by a bar or island, navigate and survey the channel with the most flow.
d) When a side channel is encountered, select the checkbox in the "OFF-CHANNEL" column
beginning with the point of divergence from the main channel, continuing downriver until the
side channel converges with the main channel.
5. Proceed downriver to the next station and repeat the above procedures.
6. Repeat the above procedures until you reach next transect. Record the distance between the
transects you just traversed in the actual transect spacing field. Ideally this distance is equal to the
intended transect spacing, but various conditions such as fast-moving water or obstructions may
necessitate a move of the transect, in which case the two values will be different.
Use the bubbles at the top of the Physical Habitat Form to select the next transect. Record
coordinates for transect mid-stream- and transect shoreline locations corresponding to the
downstream end of the thalweg sub-reach you just traversed.
NOTE: if you have taken measurements at all 10 primary thalweg stations and there is still a significant
amount of river before the next transect, you should evenly space lor 2 additional thalweg profile stations
in the remaining area to maintain the stations as evenly spaced as possible and not miss thalweg data in the
downstream end of the sub-reach. Record these additional stations in the data rows labeled 10 and/or 11 on
the form.
After completing activities at the shoreline, repeat the above procedures for each of the reach segments,
until you reach the downriver end of the reach (Transect "K").
7.
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Table 10.5 Channel Unit Categories Used on Thalweg Form
Class (Code)a
Description
Pool (PO)
Still water, low velocity, smooth, surface, deep
compared to other parts of channel
Glide (GL)
Water moving slowlv. with a smooth, unbroken
surface. Low turbulence.
Riffle (Rl)
Water moving, with small ripples, waves and eddies-
waves not breaking, surface tension not broken.
Sound: "babbling", "gurgling".
Rapid (RA)
Water movement rapid and turbulent, surface with
intermittent Whitewater with breaking waves. Sound:
continuous rushing, but not as loud as cascade.
Cascade (CA)
Water movement rapid & very turbulent over steep
channel bottom. Most of the water surface is broken
in short, irregular plunges, mostlv Whitewater. Sound:
roaring.
Falls (FA)
Free falling water over vertical or near vertical drop
into plunge, water turbulent and white over high falls.
Sound: splash to roar. (Don't navigate raft over a
waterfall!).
Dry channel (DR)
No water in the channel.
Off-channel
Side-channels, sloughs, backwaters, and alcoves
separated from the main channel.
0 In order for a channel habitat unit to be distinguished, it must be at least as wide or long as the channel is
wide.
Mid-channel bars, islands, and side channels within a thalweg profile require some guidance.
Mid-channel bars are defined as channel features below the bankfull flow level that are dry
during baseflow conditions (Section 10.6.3 defines bankfull channel). Islands are channel
features that are dry even when the river is at bankfull flow. If a mid-channel feature is as high
as the surrounding flood plain, it is considered an island. Both mid-channel bars and islands
cause the river to split into side channels. If a bar or island is encountered along the thalweg
profile, navigate and survey the channel that carries the most flow. Note that side channels are
present but do not sample them. q
When side channels are present, select the checkbox in the Thalweg Profile section of the <
Physical Habitat Form in the "Off-Channel" column. These notations will begin at the point of ^
divergence from the main channel, continuing downstream to the point of convergence with the [3
main channel. In the case of a slough or alcove, the "off-channel" notation should continue from o;
the point of divergence downstream to where the off-channel feature is no longer evident. 1
When major side channels occur, indicate in the comment bubble(s) that the feature is a side 1-
channel. h
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10.6 Channel Margin ("Littoral") and Riparian Measurements
This section covers channel margin depth and substrate, large woody debris, bank angle,
channel cross-section morphology, canopy cover, riparian vegetation structure, fish cover, and
human influences. Record measurements on the Physical Habitat Form in the NRSA App.
10.6.1 Channel Margin Depth and Substrate
Channel margin depths are measured along the designated shoreline at each transect within a
10m x 20m littoral plot that is centered on the transect (Figure 10.2). Dominant and sub-
dominant bottom substrates are determined and recorded at 5 systematically-spaced locations
that are located by eye within the 10m x 20m plot. The procedure for obtaining channel margin
depth and substrate measurements is described in more detail in Table 10.6. Record these
measurements in the Littoral and Shoreline Substrate Information section of the Physical
Habitat Form in the NRSA App. Identify the dominant and subdominant substrate present along
a shoreline swath 20 meters long and 1 meter back from the waterline. The substrate size class
choices are as shown in Table 10.6.
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UPSTREAM END
<^FLOW |
LEFT
Om BANK
bankfull line
Riparian
Plot
unvegetated bank
flow flow line
littoral plot
with depth
measures
| FLOW
Thalweg Profile
FLOW
unvegetated bank
Riparian
Plot
RIGHT
BANK
Thalweg
Profile
Increments
DOWNSTREAM END
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Table 10.6 Procedure: Channel Margin Depth and Substrate
1.
2.
3.
Access the Physical Habitat Form in the NRSA App. Be sure the correct letter designating the transect
location is selected at the top of the form. Indicate the assessed bank (left or right) as designated
during reach layout activities. Also ensure that GPS coordinates have been recorded for each
transect at the bank and midstream locations.
Determine and record the depth and the dominant and secondary substrate size class at 5
systematically-spaced locations estimated by eye within this 10m x 20m plot as well as on the
shoreline lm back from the waterline. If the substrate particle is "artificial" (e.g., concrete,
asphalt), choose the appropriate size class, record the observation and note that it is artificial in
the comment bubble.
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
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)
On the Physical Habitat Form, indicate the appropriate shore and bottom substrate type by tapping
the blue data field and selecting the correct size class. Record the depth measurements in either the
"SONAR" or "POLE" column based on the method used.
4. Repeat Steps 1 through 3 at each new cross-section transect.
10.6.2 Large Woody Debris z
o
Large Woody Debris (LWD) is defined as woody material with a small end diameter of >30 cm (1 p
ft) and a length of >5 m (15 ft). This includes any portion of woody material that meets those m
minimum size categories (e.g., include any LWD piece with at least 5 meters of its length having 2j
a diameter of 30 cm or greater). These size criteria are larger than those used in wadeable ^
streams because of the lesser role that small wood plays in controlling velocity and morphology ^
of larger rivers. The procedure for tallying LWD is presented in Table 10.7. For each tally (Wood jj
All/Part in Wetted Channel and Dry but All/Part in Bankfull Channel), the field form (Figure 10.3)
provides 12 entry boxes for tallying debris pieces visually estimated within three length and four ^
diameter class combinations. Tally each LWD piece in only one box. Tally all LWD that is wholly ^
or partially within each 10x20 littoral plot. Do not tally woody debris in the area between ^
channel cross-sections, but the presence and location of large debris dams and accumulations ^
should be mapped (sketched) and noted in the thalweg profile comments. ^
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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 10.3) refer to the large end diameter. Sometimes LWD is not cylindrical, so it has no
clear "diameter". In these cases, visually estimate what the diameter would be for a piece of
wood with circular cross-section that would have the same volume. When evaluating length,
include only the part of the LWD piece that has a diameter >0.3 m (1 ft). Count each of the LWD
pieces as one tally entry and include the whole piece when assessing dimensions, even if part of
it is outside of the bankfull channel. If you encounter massive, complex debris jams, estimate
their length, width, and height. Estimate the diameter and length of large "key" pieces and
judge the average diameter and length of the other pieces making up the jam. Record this
information in the comments section of the form.
LARGE WOODY DEBRIS (10 x 20m Plot) - tally each piece
Populate blank boxes with zeros
DIAMETER
LARGE END
0.3 - 0.6m
0.6 - 0.8m
0.8 - 1.0m
> 1.0m
WOOD ALLVPART IN WETTED CHANNEL
5-15m 15-30m >30m
DRY BUT ALL7PART IN BANKFULL CHANNEL
5-15m 15-30m >30m
Figure 10.3. Large Woody Debris Section of Physical Habitat Form in the NRSA App - Non-Wadeable
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Table 10.7 Procedure: Tallying Large Woody Debris
Note: Tally pieces of large woody debris (L WD) within the 11 transect littoral plots of the river reach at
the same time the shoreline measurements are being determined. Include all pieces whose large end is
located within the littoral plot in the tally. Tally wood that is at least partially within the wetted channel
separately from that that is not presently wetted, but still within or directly above (bridging) the bankfull
channel
1. LWD is tallied in 11 "plots" systematically spaced over the entire length of the sampling reach. These
plots are each 20 m long in the upstream-downstream direction (10 m up, 10 m down from the
transect). They are positioned along the chosen bank and extend from the shore 10 m towards mid-
channel and then all the way to the bankfull margin.
2. Tally all LWD pieces within the plot that are at least partially within the presently wetted (baseflow)
channel. First, determine if a piece is large enough to be classified as LWD (any portion with a
diameter of at least 30 cm [1 ft.] and a length of at least 5 m [15 ft.]).
3. For each piece of LWD, determine its diameter class based on the diameter of the large end (0.3 m
to < 0.6 m, 0.6 m to <0.8 m, 0.8 m to <1.0 m, or >1.0 m), and the length class of the LWD pieces
based on the part of its length that has diameter >30 cm. Length classes are 5 m to <15 m, 15 m to
<30 m, or >30 m.
If the piece is not cylindrical, visually estimate what the diameter would be for a piece of wood
with circular cross-section that would have the same volume.
When estimating length, include only the part of the LWD piece that has a diameter >0.3 m (1 ft).
4. Tally each piece of LWD in the appropriate diameter and length class tally box in the "WOOD
All/Part in WETTED Channel" columns in the Large Woody Debris section of the Physical Habitat
Form in the App.
5. Tally all shoreline LWD pieces along the littoral plot that are at least partially within or above
(bridging) the bankfull channel, but not in the wetted channel. For each piece, determine the
diameter class based on the diameter of the large end (0.3 m to < 0.6 m, 0.6 m to <0.8 m, 0.8 m to
<1.0 m, or >1.0 m), and the length class based on the length of the piece that has diameter >30 cm.
Length classes are 5m to <15 m, 15 m to <30 m, or >30 m.
6. Tally each piece of LWD in the appropriate diameter and length class tally box in the "DRY BUT
ALL/PART IN 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. When using paper forms, place a
tally mark for each piece in the appropriate diameter x length class tally box in the
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.
8. Repeat Steps 1 through 6 for the next river transect.
10.6.3 Bank Angle and Channel Cross-Section Morphology
Bank angles of undercut, vertical, steep, and gradual are visually estimated as defined on the
Physical Habitat Form. Observations are made from the wetted channel margin up 5 m (a
canoe's length) into the bankfull channel margin on the previously chosen side of the stream.
You will measure or estimate the wetted width, mid-channel bar width, bankfull height and
width, the amount of incision, and the degree of channel constraint. These are assessed for the
whole channel (left and right banks) at each of the 11 cross-section transects. The procedures
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for obtaining bank angle and measurements of channel cross-section morphology are presented
in Table 10.8.
Wetted width is the width of the channel containing free-standing water; if >15 m, it can be
measured with a laser rangefinder. Mid-channel bar width, the width of exposed mid-channel
gravel or sand bars, is included within the wetted width, but is also recorded separately. In
channel cross-section measurements, the wetted and bankfull channel boundaries include mid-
channel bars. Therefore, the wetted width is measured as the distance between wetted left and
right banks. If islands are present, treat them like bars, but flag these measurements by
indicating in the adjacent comment bubble that the "bar" is an island. If you are unable to see
across the full width of the river when an island separates a side channel from the main channel,
record the width of the main channel, flag the observation, and note in the comment bubble
that the width pertains only to the main channel.
Table 10.8 Procedure: Bank Angle and Channel Cross-Section
Bank Angle and Cross Section Methods
Record the wetted width of the river and the width of exposed mid-channel bars (if present) in
the Bank Characteristics section of the Physical Habitat Form in the App. Also determine the
bankfull channel width.
Visually estimate the bank angle (undercut, vertical, steep, gradual), as defined on the form.
Bank angle observations refer to the area from the wetted channel margin up 5 m (canoe's
length) into the bankfull channel margin on the previously chosen side of the river. Mark the
angle in the "Bank Measurements" section of the Physical Habitat Form.
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 bankfull height 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.
Hold the surveyor's rod vertically, with its base planted at the water's edge. Examine both banks,
then determine the channel incision as the height up from the water surface to elevation of the
first terrace of the valley floodplain (Note this is at or above the bankfull channel height).
Whenever possible, use the clinometer as a level (positioned so it reads 0% slope) to measure
this height by transferring (backsighting) it onto the surveyor's rod. Record this value in the
Incised Height field of the "Bank Measurements" section of the form.
Repeat Steps 1 through 4 at each cross-section transect. Record data for each transect on a
separate field data form.
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Bankfull flows are large enough to erode the stream bottom and banks, but frequent enough
(every one to two years) to not allow substantial growth of upland terrestrial vegetation.
Consequently, in many regions, it is these flows that have determined the width and depth of
the channel. Estimates of the bankfull dimensions of stream channels are extremely important
in the NRSA. They are used to calculate shear stress and bed stability (see Kaufmann et al.,
1999). Unfortunately, we have to depend upon evidence visible during the low-flow sampling
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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 deposi-
tional surface that frequently is flooded and experiences sediment deposition under
the current climate and hydrological regime.
Then look specifically for:
An obvious break in the slope of the banks.
A change from water-loving and scour-tolerant vegetation to more
drought-tolerant vegetation.
A change from well-sorted stream sediments to unsorted soil materials.
In the absence of clear bankfull indications, consider the previous season's flooding as the best
evidence available (note: you could be wrong if very large floods or prolonged droughts have
occurred in recent years.). Look for:
Drift debris ("sticky wickets" left by the previous seasons flooding).
The level where deciduous leaf-fall is absent on the ground (carried away by
previous winter flooding).
Unvegetated sand, gravel or mud deposits from previous year's flooding.
In years that have experienced large floods, drift material and other recent high flow markers
may be much higher than other bankfull indicators. In such cases, base your determination on
less transient indicators such as channel form, perennial vegetation, and depositional features.
In these cases, flag your data entry and also record the height of drift material in the comments
section of the field data form.
We use the vertical distance (height) from the observed water surface up to the level of the first
major valley depositional surface (Figure 10.4) 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
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floodplain, making "Bankfull Height" smaller than "Incision" (Figure 10.5). Bankfull height is
never greater than incision height. Look for evidence of recent flows (within about 1 year) to
distinguish bankfull and incision heights, though recent flooding of extraordinary magnitude
may be misleading. In cases where the channel is cutting a valley side slope 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 determine incision height and bankfull
height. Remember that incision height is measured as vertical distance to the first terrace
above bankfull; if terrace heights differ on left and right banks, choose the lower of the two
terraces. Even when quite constrained by their valley side slopes, large rivers often have flood
terraces above bankfull height. In some cases, though, your sample reach may be in a steep "V"
shaped valley or gorge formed over eons, and the slopes of the channel banks simply extend
uphill indefinitely, not reaching a terrace before reaching the top of a ridge. In such cases,
record incision height values equal to bankfull values and make appropriate comments that no
terrace is evident. Similarly, when the river is extremely incised below an ancient terrace or
plateau,(e.g., the Colorado River in the Grand Canyon), you may crudely estimate the terrace
height if it is the first one above bankfull level. If you cannot estimate the terrace height, make
appropriate comments describing the situation.
Finally, assess and record the local degree of river channel constraint (i.e., at the transect) by
following the guidelines on the Channel Constraint Section of the form regarding the
relationships among channel incision, valley side slope, and width of the valley floodplain. Mark
whether you could or could not readily see over the bank and record the distance from shore to
riparian vegetation in the field provided. You will also do an overall assessment of channel
constraint for the whole river reach; see Section 10.11 for a discussion of constraint concepts
and assessment procedures.
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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
Former second
terrace becomes
Former active floodplain
Former first third terrace
no longer connected terrace becomes
becomes new first terrace seconcl terrace
above bankfull level
(Record this height)
Recent incision-
bankfull level below
first terrace of valley
bottom
Valley Fill
Figure 10.4 Schematic Showing Bankfull Channel and Incision for Channels
(A) not recently incised, and (B) recently incised into valley bottom. Note level of bankfull stage relative to
elevation of first terrace on valley bottom (stick figure included for scale)
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A) Deeply Incised Channel
Incision Height (Always
equal to or greater than
bankfull height)
From Figure 7-9 (B)
Dowtcutting over
Second Terrace
V
First Terrace
Height
(When
channel form
is not a good
indicator, use
evidence of
recent
flooding)
Former second
terrace becomes
Former first ^1,rci twracc
Bankfull Height -
(when channel form is
not a good indicator,
use evidence of recent
flooding, lack of
permanent flood-
intolerant vegetation)
Incision Heปght=
Bankful Height
"Hood-
to leiant
jfegetation
No incision:
No evidence of
downcutting,
vertical bank
angle, etc.)
B) Small stream constrained in V-shaped valley
Flood-
ntoterant
vegetation
Figure 10.5 Determining Bankfull and Incision Heights for
(A) Deeply Incised Channels, and (B) Streams in Deep V Shaped Valleys (Stick figure included for scale)
10.7 Visual Riparian Estimates
10.7.1 Riparian Vegetation Structure
Riparian vegetation observations apply to the riparian area upstream 10 m and downstream 10
m from each of the 11 transects. They include the visible area from the river bankfull margin
back a distance of 10 m (30 ft) shoreward from both the left and right banks, creating a 10 m x
20 m riparian plot on each side of the river (Figure 10.2). The riparian plot dimensions are
estimated, not measured. Table 10.9 presents the procedure for characterizing riparian
vegetation structure and composition.
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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, broadteaf 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
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 data 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) rankings. 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 10.9 Procedure: Characterizing Riparian Vegetation Structure
Riparian Vegetative Cover
4.
5.
Anchor or tie up at the river margin at a cross-section transect; then make the following observa-
tions to characterize riparian vegetation structure.
Estimate the distance from the shore to the edge of the riparian vegetation plot (i.e., the distance
from the current water level to the bankfull margin); record it in the "Channel Constraint" section
of the Physical Habitat Form in the App.
Facing the left bank (left as you face downstream), estimate a distance of 10 m back into the
riparian vegetation, beginning at the bankfull channel margin. Estimate the cover and structure
of riparian vegetation within an estimated 10 m x 20 m plot centered on the transect and starting
where perennial vegetation begins or at the bankfull river margin (whichever is closest to the
river shoreline). On steeply-sloping channel margins, estimate the riparian plot dimensions as if
they were projected down from an aerial view.
Within this 10 m x 20 m area, conceptually divide the riparian vegetation into 3 layers: a CANOPY
(>5 m high), an UNDERSTORY (0.5 to 5 m high), and a GROUND COVER layer (<0.5 m high).
Within this 10 m x 20 m area, determine the dominant woody vegetation type for the CANOPY
LAYER (vegetation > 5 m high) as either Deciduous, Coniferous, broadleaf Evergreen, Mixed, or
None. Consider the layer "Mixed" if more than 10% of the areal coverage is made up of the
alternate vegetation type. If the 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.
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
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of shadow that would be cast by a particular layer alone if the sun were directly overhead.
Record the appropriate cover class on the data form ("0" = absent, zero cover; "1" = sparse,
<10%; "2" = moderate, 10-40%; "3" = heavy, 40-75%; or "4" = very heavy, >75%).
7. Look at the UNDERSTORY layer (vegetation between 0.5 and 5 m high). Determine the dominant
woody vegetation type for the understory layer as described in Step 5 for the canopy layer. If the
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".
8. 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 6 for
the canopy layer.
9. Look at the GROUND COVER layer (vegetation < 0.5 m high). Determine the areal cover class for
woody shrubs and seedlings (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 6 for the
canopy layer. Include all non-vegetative materials such as buildings, pavement, standing water,
etc. In the "barren, bare dirt, or duff category.
10. Repeat Steps 1-9 for the plot on the opposite side of the river and repeat the entire process for
all remaining transects.
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10.8 Instream Fish Cover, Algae, and Aquatic Macrophytes
Over a defined length and distance from shore at the sampling locations (Figure 10.2), 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 fish cover is outlined in Table 10.10.
Record data in the Fish Cover section of the Physical Habitat Form in the NRSA App. Crews will
estimate the areal cover of all of the fish cover and other listed features that are in the water
and on the banks within the 10 m x 20 m plot only on the side of the river previously chosen for
assessment during reach layout (Figure 10.1). The areal cover classes offish concealment and
other features are the same as those described for riparian vegetation (Section 10.7.1).
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 river which could provide
cover for fish or macroinvertebrates. If the river channel contains live wetland grasses, include
these as macrophytes. Woody debris are the larger pieces of wood that can provide cover and
influence stream morphology (i.e., those pieces that would be included in the large woody
debris tally [Section 10.6.2]). Brush/woody debris refers to the smaller wood pieces that
primarily affect cover but not morphology. Live Trees or Roots are living trees that are within the
< channel - estimate the areal cover provided by the parts of these trees or roots that are
< inundated. Overhanging vegetation includes tree branches, brush, twigs, or other small debris
u 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
E would be inundated during bankfull flows. Boulders are typically basketball to car-sized
in 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.
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Table 10.10 Procedure: Estimating Fish Cover
Fish Cover
1. Stop at the designated shoreline at a cross-section transect and estimate a 10 m distance
upstream and downstream (20 m total length), and a 10 m distance out from the banks to define
a 20 m x 10 m littoral plot.
2. Examine the water and the banks within the 20 m x 10 m littoral plot for the following features
and types offish cover '.filamentous algae, macrophytes, large woody debris, in-channel live trees
or roots, brush and small woody debris, overhanging vegetation, undercut banks, boulders, and
artificial structures.
3. For each cover type, estimate its areal cover by eye and/or by sounding with a pole. Record the
appropriate cover class in the "Fish Cover" 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.
10.9 Human Influences
For the left and right banks at each of the 11 detailed Channel/Riparian Cross-Sections, evaluate
the presence/absence and the proximity of 11 categories of human influences outlined in Table
10.11. Record data in the Human Influence section of the Physical Habitat Form in the App.
Relate your observations and proximity evaluations to the river and riparian area within 10 m
upstream and 10 m downstream from the transect (Figure 10.6). Four proximity classes are
used: In the river or on the bank within 10 m upstream or downstream of the cross-section
transect (B), contained within the 10 m x 20 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 an
another transect, or its 10x20 meter riparian plot.
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Cwithin riparian plot
B on bank or in stream
Poutside plot
(but do not sight through next
/ transect or plot)
Figure 10.6 Proximity Classes for Human Influences in Non-Wadeable Rivers
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Table 10.11 Procedure: Estimating Human Influence
Human Influence
1. Stop at the designated shoreline at a cross-section transect, look toward the left bank (left when
facing downstream), and estimate a 10 m distance upstream and downstream (20 m total length).
Also, estimate a distance of 10 m back into the riparian zone to define a riparian plot area.
2. Examine the channel, bank and riparian plot area adjacent to the defined river segment for the
following human influences: (1) walls, dikes, revetments, riprap, & dams; (2) buildings; (3) cleared
lot, pavement (e.g., paved, graveled, dirt parking lot, foundation); (4) roads or railroads, (5) inlet or
outlet pipes; (6) landfills or trash (e.g., cans, bottles, trash heaps); (7) parks or maintained lawns; (8)
row crops; (9) pastures, rangeland, or hay fields; (10) logging; and (11) mining (include gravel
mining). 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.
3. For each type of influence, determine if it is present and what its proximity is to the river and
riparian plot area. Consider human disturbance items as present if you can see them from the cross-
section transect. Do not include them if you have to site through another transect or its 10 m x 20 m
riparian plot.
4. For each type of influence, record the proximity class in the Human Influence part of the Visual
Riparian Estimates section of the Physical Habitat Form in the App. If a disturbance is within more
than one proximity class, record the one that is closest to the river. Proximity classes (listed in order
of priority) are:
B (Bank) Present within the defined 10 m stream segment and located in the stream
or on the stream bank. The bank is defined as the area between the water's
edge and the bankfull margin.
C (Contained) Present within the 10 x 20 m riparian plot area, but away from the bank.
P (Present) Present, but outside the riparian plot area.
0 (Absent) Not present within or adjacent to the 20 m stream segment or the riparian
plot area at the transect
5. Repeat Steps 1 through 4 for the opposite bank.
6. Repeat Steps 1 through 5 for each cross-section transect.
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10.10 Canopy Cover Measurements
Canopy cover over the river is determined on the near bank 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).
Measure vegetative cover over the reach at the chosen bank at each of the 11 transects (A-K).
Four measurements are obtained at each cross-section transect (upriver, downriver, left, and
right).
The procedure for obtaining canopy cover data is presented in Table 10.12. 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 four measurement points, record the number of
intersection points (maximum=17) that have vegetation or other overhead objects covering
them in the Canopy Density @ Bank section of the Physical Habitat Form in the NRSA App.
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."
TAPE
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Table 10.12 Procedure: Canopy Cover Measurements
Canopy Cover Methods
1. Take densiometer readings at a cross-section transect while anchored or tied up at the river
margin.
2. Hold the densiometer 0.3 m (1 ft) above the surface of the river. Holding the densiometer level
using the bubble level, move it in front of you so your face is just below the apex of the taped "V".
3. At the channel margin measurement locations, count the number of grid intersection points within
the "V" that are covered by either a tree, a leaf, a high branch, or the bank itself.
4. Take 1 reading each facing upstream (UP), downstream (DOWN), left bank (LEFT), and right bank
(RIGHT). Right and left banks are defined with reference to an observer facing downstream.
5. Record the UP, DOWN, LEFT, and RIGHT values (0 to 17) in the "CANOPY COVER @ BANK" section
of the form.
6. Repeat Steps 1 through 5 at each cross-section transect.
10.11 Channel Constraint, Debris Torrents and Recent Floods
10.11.1 Channel Constraint
After completing the thalweg profile and riparian/channel cross-section measurements and
observations, envision the stream at bankfull flow and evaluate the degree, extent and type of
channel constraint, using the procedures presented in Table 10.13. 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 10.8):
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.
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Table 10.13 Procedure: Assessing Channel Constraint
Channel Constraint
NOTE: These activities are conducted after completing the thalweg profile and littoral-riparian
measurements and observations, and represent an evaluation of the entire stream reach.
Channel Constraint: Determine the degree, extent, and type of channel constraint based on envisioning the
stream at bankfull flow.
1. Classify the stream reach channel pattern as predominantly a single channel, an
anastomosing channel, or a braided channel.
Single channels may have occasional in-channel bars or islands with side channels, but
feature a predominant single channel, or a dominant main channel with a subordinate
side channel.
Anastomosing channels have relatively long major and minor channels branching and
rejoining in a complex network separated by vegetated islands, with no obvious
dominant channel.
Braided channels also have multiple branching and rejoining channels, separated by
unvegetated bars. Subchannels are generally small, short, and numerous, often with no
obvious dominant channel.
2. After classifying the channel pattern, determine whether the channel is constrained within
a narrow valley, constrained by local features within a broad valley, unconstrained and
free to move about within a broad floodplain, or free to move about, but within a
relatively narrow valley floor.
3. Then examine the channel to ascertain the bank and valley features that constrain the
stream. Entry choices for the type of constraining features are bedrock, hillslopes,
terraces/alluvial fans, and human land use (e.g., a road, a dike, landfill, rip-rap, etc.).
4. Based on your determinations from Steps 1 through 3, select and record one of the
constraint classes shown on the Channel Constraint Form.
5. Estimate the percent of the channel margin in contact with constraining features (for
unconstrained channels, this is 0%). Record this value on the Channel Constraint Form.
6. Finally, estimate the "typical" bankfull channel width, and visually estimate the average
width of the valley floor. Record these values on the Channel Constraint Form.
NOTE: To aid in this estimate, you may wish to refer to the individual transect assessments of incision and
constraint that were recorded on the 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 field form that indicates this fact.
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A) Anastomosing channel pattern
222 Vegetated islands above bankfull flow. Multiple
channels remain during major flood events.
B) Braided channel pattern
FLOW
channel pattern disappears during major flood events.
DVP
Figure 10.8 Types of Multiple Channel Patterns
10.11.2 Debris Torrents and Recent Major Floods
Debris torrents, or lahars, differ from conventional floods in that they are flood waves of higher
magnitude and shorter duration, and their flow consists of a dense mixture of water and debris.
Their high flows of dense material exert tremendous scouring forces on streambeds. For
example, in the Pacific Northwest, flood waves from debris torrents can exceed 5 meters deep
in small streams normally 3 m wide and 15 cm deep. These torrents move boulders in excess of
1 m diameter and logs >1 m diameter and >10 m long. In temperate regions, debris torrents
occur primarily in steep drainages and are relatively infrequent, occurring typically less than
once in several centuries.
Because they may alter habitat and biota substantially, infrequent major floods and torrents can
confuse the interpretation of measurements of stream biota and habitat in regional surveys and
monitoring programs. Therefore, it is important to determine if a debris torrent or major flood
has occurred within the recent past. After completing the thalweg profile and channel/riparian
measurements and observations, examine the stream channel along the entire sample reach,
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including its substrate, banks, and riparian corridor, checking the presence of features described
on the Torrent Evidence 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.
10.12 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 (Non-Wadeable Sites)
For collecting samples
nitrile gloves
pre-sterilized, 250 ml sample bottle
sodium thiosulfate tablet
Wet ice
cooler
For recording measurements
Sample Collection Form 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 (Non-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.
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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.
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. 2009, USEPA 2013a) and
possibly Observed/Expected (O/E) taxa richness. In addition, the fish assemblage data provides a
starting point for developing potential indicators of ecosystem services related to fish.
In non-wadeable rivers, collect fish using boat (or raft) electrofishing over a defined fish
sampling reach (which will be between 20 and 40 times the mean channel width [CW] recorded
on the Verification Form). The fish sampling reach is less than or equal to the sampling reach
established for the site (See Section 3.2). Use secondary fish collection methods in habitat that
cannot be adequately sampled by boat (or raft). Secondary methods may include backpack or
some type of towed (e.g., a towed barge, small watercraft, or float tube) electrofishing unit,
using your boat (or raft) as a towed barge, or seining as a last option (only if conductivity is too
high or too low for electrofishing). Conduct sampling in a downstream direction, allocating effort
(button time) within subreaches (areas between the cross-section transects). If you have not
collected 500 individuals at the end of the minimum fish sampling reach (20 CW; 5 subreaches),
sample additional subreaches in their entirety until you obtain at least 500 individuals or sample
the entire sampling reach (40 CW; 10 subreaches). Record information related to sampling
effort on the front of the Fish Gear Form in the NRSA App. Record species identification and
enumeration data on the Fish Collection Form in the NRSA App.
While conducting fish assemblage sampling, crews will also aim to collect two individuals from
which a fish tissue plug will be collected (see Section 13). The fish plug sample will consist of two
fish tissue plugs for mercury analysis and will be collected from two fish of the same species at
all sites where suitable fish species and lengths are available. These fish are often collected
during the fish assemblage sample collection effort, but additional effort can be used to collect
specimens for tissue samples if desired. If crews do not sample the reach for fish assemblage
data (due to permit restrictions, site conditions, etc.), they are still encouraged to conduct
sampling for fish tissue plug specimens using any effective gear, including hook and line. If you
do not conduct fish assemblage sampling, do not record any gear or sampling details on the Fish
Gear Form, other than to indicate the reason the site was not fished in the "Fish Sampling - IF
NOT ROUTINE OR SUSPENDED" section. In this case you will also leave the Fish Collection Form
blank except to enter the pertinent details about the fish plug specimen(s).
12.2 Equipment and Supplies
Table 12.1 shows the checklist of equipment and supplies required to complete the non-
wadeable fish sampling. This checklist is similar to the one presented in Appendix A, which is
used at the base location to ensure that all of the required equipment is brought to the site.
Table 12.1 Equipment and Supplies: Fish Sampling (Non-Wadeable Sites)
For collecting
Boat, motor, and trailer (and necessary
GPS with transect waypoints
fish
safety equipment)
preloaded
Gasoline and oil (if using a 2 cycle motor or
generator)
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For recording
measurements
Boat electrofishing equipment
Pulsator Control Box
Foot Pedal
Anode Droppers
Generator
Linesman's Gloves
Hearing Protection
Towed electrofishing equipment (e.g.,
towed barge, or mounted on a small
watercraft or float tube), if needed for
subreaches too shallow to sample from a
boat or raft used as a towed unit.
Probes with extensions.
Appropriate switching box
Pulsator control box (5.0 GPP or less)
Backpack electrofisher (as used for
wadeable streams)
Dip nets (non-conductive handles) mesh
Scientific collection permit(s)
10 ft x 6 ft minnow or bag seine with % in.
mesh (additional 4 ft depth seine may also
be used)
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 NRSAApp
Fish Collection Form in NRSA App
Several Leak proof HDPE jars for fish
voucher specimens (various sizes
from 250 mL- 4 L)
1 scalpel for slitting open large fish
before preservation
1 container of 10% buffered formalin
1 aquarium net for dipping small fish
from live well
2 measuring boards (3 cm size
classes) (optional; needed only if
quantitative length data are needed)
1 set Fish ID keys
Field Operations Manual
Digital camera with extra memory
card & battery
20 ft x 6 ft minnow or bag seine with
V* inch mesh (additional 4 ft depth
seine may also be used)
Polarized sunglasses and hats
Buckets
Soft (#2) lead pencils for writing on
voucher inner labels
Fine-tip indelible markers
Clear tape strips
12.3 Sampling Procedures
Table 12.2 describes the procedure for collecting fish in non-wadeable streams. The sampling
crew should consist of one boat operator (who also controls the electrofishing unit) and one dip
netter (equipped with a 1/4" mesh dip net) situated at the bow. Begin sampling at the upstream
end of the sampling reach defined for the site (See Section 3.2) and proceed downstream. 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.
The reach length sampled for fish varies based on the mean width of the river used to establish
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).
In small rivers (mean channel width [CW] recorded on the Verification Form is < 12 meters),
conduct sampling in the same fashion as for small wadeable streams, sampling the entire g
sampling reach (150 m minimum or 40 CW; 10 subreaches) and moving the boat (or raft) within <
each subreach to sample both shorelines as well as the mid-channel. ^
LU
In large rivers (mean channel width [CW] recorded on the Verification Form > 13 meters) the $
minimum length for fish sampling is 20 CW (5 subreaches). If a minimum of 500 fish are not ^
collected after sampling the minimum fishing reach, sample additional subreaches in their
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entirety until 500 fish are collected or all 10 subreaches have been sampled. For rivers where
the CW is > 13 m wide, restrict sampling to shoreline habitats within each subreach. Start on the
same bank as the habitat crew, and move to the opposite bank after every two subreaches.
Within each subreach, sample for ~700 seconds of "button time." Prior to sampling each
subreach, determine the most appropriate gear for the subreach (e.g., boat or raft vs. barge or
backpack electrofishing units, or seines if the conductivity is too high or too low). When
electrofishing, proceed downstream at a pace equal to or slightly greater than the prevailing
current to maximize capture efficiency. Maneuver the electrofishing unit in and around complex
habitat when necessary; however, use discretion in sampling these habitats to maintain equal
effort among subreaches.
Table 12.2 summarizes the fishing protocols for each of two sizes of non-wadeable rivers and
Table 12.3 provides detailed steps for fishing non-wadeable rivers.
Whenever possible, process fish at the end of each subreach to minimize mortality and stress to
fish. You can use multiple lines per species on the Fish Collection Form if necessary (e.g., you
collect a large number of individuals and need additional space for tallying).
If seining is used, record all fish collected with seining protocols on separate lines of the Fish
Collection Form.
At the end of the minimum fish sampling reach, determine if you have collected at least 500
individuals. If so, stop sampling. If not, sample additional subreaches (one at a time and in their
entirety) until at least 500 individuals are captured. 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). Stop sampling when you reach Transect K (the end of the
sampling reach), regardless of the number of individuals collected.
It is important that you record the total reach length that was sampled for fish, as this is used
along with the number offish 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.
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 Non-wadeable Fishing Protocols
Small Non-Wadeable Rivers (mean channel width [CW1 from Verification Form < 12 meters)
Fish Sampling reach will be between 150 and 480 meters
Subreaches will be between 15 and 48 meters each
S Sample all 10 subreaches in their entirety from bank to bank starting at Transect A
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If islands are encountered, crews should still fish bank to bank, including both sides of
the island.
S Total button time will range from 500-700 seconds per subreach
You do not have to expend equal button time among the 10 subreachesyou can
devote more button time to subreaches with more complex habitat.
S No minimum fish number
Large Non-Wadeable Rivers (CW >13 meters)
Fish Sampling reach will be between 260 and 4000 meters
Subreaches will be 1/10 of the sampling reach
Minimum fishing length = 5 subreaches
S Fish each subreach along bank in pairs of subreaches starting at the random PHab bank at
Transect A
If islands are encountered, crews will only fish the edge of the island that occurs along
the main channel.
S Button time is roughly 700 seconds per subreach
Depending upon the habitat complexity, you can vary the distance actively fished to
allocate the available button time throughout the subreach.
S Minimum fish number is 500 unless all 10 subreaches have been fished.
> After fishing 5 subreaches, if 500 fish have not been collected, add subreaches one at
a time (but fish them in their entirety) until 500 fish are collected or all 10 subreaches
(40 CW) have been fished.
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Small Nonwadeable River: Mean Channel Width < 12 rn
40x Channel Width
Large Nonwadeable River: Mean Channel Width > 13 m
< 20 x Channel Width ป
CW (5 subreaches), continue fishing next
subreach (alternating banks) until either 500
individuals are collected, or Transect K is
reached (10 subreaches sampled; 40 CW)
Figure 12.1 Reach Layouts for Fish Sampling at Non-Wadeable Sites
Dark shaded areas indicate the minimum length of the fish sampling reach. Light shaded areas are
sampled as needed to meet the required 500 individuals.
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Table 12.3 Procedure: Electrofishing (Non-Wadeable Sites)
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 boat or raft electrofishing (e.g., no access for boat or raft, safety
concerns, ambient conductivity is too high or too low to use a boat electrofishing unit),
determine if you can sample with the use of secondary methods (backpack or barge shocker,
or by seining).
i) If yes, follow the procedures presented in Section 12.3.2 and/or Section 12.3.3.
ii) If not, select Not Fished - Site Conditions Prohibit Sampling. Provide a comment describing
why site conditions prohibit sampling.
c) If you determine that > 50% of the required fish sampling reach is physically inaccessible or
otherwise unsampleable by any means, 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) In a non-wadeable river, you should always attempt to sample; never 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 either Large Nonwadeable or Small Nonwadeable in the Fish
Sampling Protocol section based on the mean wetted width reported on the Verification Form.
a) For safety, everyone must wear PFDs, foot protection, and insulated linesman's gloves.
b) To aid vision while netting fish, wear polarized sunglasses and a hat or visor.
4) Determine the minimum length for the fish sampling reach based on the width used to define the
sampling reach for the river (rounded to the nearest meter and recorded on the Verification Form).
a) If the mean channel width (CW) from the Verification Form is <12 m, the fish sampling reach
length is 150 m or 40 CW (10 subreaches).
b) If the CW is >13 m, the minimum fish sampling reach length is 20 channel widths (5
subreaches).
5) 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 ฐC).
6) Mark either Boat or Raft as the Primary Electrofishing Gear. Boats use outboard motors and can
travel upstream if needed, while rafts are oar-powered and generally cannot move upstream.
a) Only one netter should be used. Record this along with the number of anodes, and their
diameter (mark the units as either inches or centimeters) on the Fish Gear Form. Diameter
refers to the diameter of a hoop or sphere or the distance between the outer edge of the
shape formed by an array of droppers (which is usually round).
b) Set the boat electrofishing unit to pulsed DC and mark it in the Wave Form section of the Fish
Gear Form.
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c) Test the other settings outside of the sampling area. Start the electrofisher, set the timer, and
depress the switch to begin fishing. See Tables in Appendix E for example starting settings. If
fishing success is poor, increase the pulse width first and then the voltage. Increase the pulse
rate last to minimize mortality or injury to large fish. If mortalities occur, first decrease pulse
rate, then the voltage, then the pulse width.
i) 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.
ii) If the electrofishing system only lets you adjust the percent of power, record the value on
the Fish Gear Form.
iii) If your conductivity meter cannot measure ambient conductivity, you can "uncorrect"
specific conductance at 25 ฐC to ambient conductivity using the following equation:
(1) Ambient conductivity=Specific conductance x (l+([water temp-25 ฐC] x 0.02)).
d) Record the final settings that will be used for sampling on the Fish Gear Form.
7) Begin sampling at the upstream end of the reach (Transect A). Start on the bank (right or left as you
face downstream) selected at random to begin the physical habitat sampling.
a) For small rivers (< 12 m wide), move the boat (or raft) within each subreach to sample both
shorelines as well as the mid-channel areas (similar to what is done for a small wadeable
stream using a backpack electrofishing unit).
b) For rivers > 13m wide, sample along the shoreline only. Move to the opposite bank after every
two subreaches. Within each subreach, proceed downstream in close proximity to the bank
and at a pace equal to or slightly greater than the prevailing current to maximize capture
efficiency. You can "nose in" your boat or raft into shoreline habitat to effectively sample but
do not remain in that habitat for too long.
8) Generally, effort (i.e., button time) should be ~500 -700 seconds per subreach.
a) At sites with maximum reach length (4000 m) it is likely that the entire length within each
subreach (400 m) will not be fished. Depending upon the habitat complexity, you can vary the
distance actively fished to allocate the available button time throughout the subreach. You do
not have to expend equal time among the subreaches.
b) Avoid the temptation to focus sampling in the richest habitat types.
9) Use a 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.
a) Irruptive species: If you encounter a large school of a single species (e.g., shad, certain shiners,
etc.), quickly move through it to ensure you can sample the entire subreach within the allotted
button time.
10) Sampling with Secondary Gear: If shallow habitat exists within a subreach that is inaccessible to
your boat or raft, use secondary collection methods to sample the habitat thoroughly. If you use a
secondary electrofishing gear to sample some portion of a subreach, follow the directions provided
in Table 12.4. As a last option, if the conductivity is too high or too low to use a backpack or barge
electrofishing units, seine those areas you cannot access with a boat, following the directions
presented in Table 12.5.
11) Process fish at the completion of each transect or as often as necessary to reduce mortality.
Release fish in a location that eliminates the likelihood of recapture.
12) Repeat Steps 7 through 11 until you have sampled the minimum reach length determined in Step
4.
13) If you have sampled the required minimum fishing reach length (20 CW; 5 subreaches) determine
the total number of individuals collected.
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a) If the total is < 500, sample one or more additional subreaches until at least 500 individuals
have been collected and processed, or you sample all 10 subreaches (40 CW). Go to Step 14.
b) If you collect > 500 individuals, determine if a single irruptive species comprises > 50% of the
total number of individuals.
i) If irruptive species make up > 50% of the sample, sample one or more additional
subreaches to bring the proportion of the irruptive species below 50%. Go to Step 14.
ii) If not, go to Step 14.
c) If you have sampled all 10 subreaches (i.e., you have reached Transect K; 40 CW, go to Step 14.
14) After sampling all the required subreaches, record the final length of the fish sampling reach (as
tracked by GPS or measured by range finder) sampled with the primary gear type in the Primary
Electrofishing Gear section of the Fish Gear Form.
a) If you suspend fish sampling before completing the minimum distance (20 CW; 5 subreaches),
record the length that was sampled in the Primary Electrofishing Gear section, and mark the
reason for the suspension in the Fish sampling - Not Routine or Suspended section of the Fish
Gear Form.
15) In the Primary Electrofishing Gear section of the Fish Gear Form, record the total button time
expended for boat electrofishing, the total time spent sampling, and the total fish sampling reach
sampled by the boat electrofishing unit.
a) In the Secondary Electrofishing Gear section of the Fish Gear Form, record the total button
time expended for secondary electrofishing, the total time spent sampling, and length of the
total fish sampling reach sampled by the secondary electrofishing method (if applicable).
b) If seining was used as an additional collection method, record the total number of hauls, the
average haul length, the total time spent seining, and the total fish sampling reach sampled for
each type of seine in the Primary and Secondary Seine Gear sections of the Fish Gear Form.
16) 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.
17) Note the general response behavior offish 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).
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).
18) 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.
12.3.2 Secondary Electrofishing
If shallow habitat exists within a subreach that is inaccessible to your boat or raft, it will be
necessary to use secondary collection methods to sample the habitat thoroughly. Table 12.4
presents the procedure for collecting fish from large wadeable areas of subreaches through the
use of secondary electrofishing techniques. The intent of the secondary electrofishing methods
is to provide comparable data to boat electrofishing. Do not use your boat as a wadeable fj
fishing unit (by attaching an anode pole to the box) if your electrofishing unit is larger than ^
5000 V (e.g., Generator Powered Pulsator [GPP] 5.0). Record information about the gear used ^
in the Secondary Electrofishing Gear section of the Fish Gear Form in the NRSA App. At the end <
of sampling, record the button time of the secondary gear, the total amount of time spent using <ฑi
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the secondary method, and the length of the total fish sampling reach that was sampled with
the secondary method. The total button time for boat and wadeable electrofishing combined
should be ~700 seconds per subreach. 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.
Table 12.4 Procedure: Secondary Electrofishing Methods for Wadeable Areas (Non-Wadeable Rivers)
1) Use secondary electrofishing as a collection method whenever an area within a subreach is
inaccessible by boat (e.g., a large riffle area). The area must be safe to wade. Do not use your boat
as a wadeable fishing unit (by attaching an anode pole to the box) if your electrofishing unit is
larger than 5000 V (e.g., GPP 5.0).
2) Mark either Backpack or Bank or Towed Unit in the Secondary Electrofishing Gear section of the Fish
Gear Form.
a) Only one netter and one anode should be used. Record this along with the anode diameter
(mark the units as either inches or centimeters) on the Fish Gear Form.
3) Operation of Backpack Electrofisher
a) For backpack electrofishing there may be from 2 to 3 people involved (depending upon the crew
size).
b) Set unit to pulsed DC and mark it in the Wave Form section of the Fish Gear Form.
c) Select the initial voltage setting based on the ambient conductivity of the river,
i) See Tables in Appendix E.
d) 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.
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.
4) Operation of Towed Electrofisher
a) When using a towed electrofishing unit, the minimum crew size for electrofishing is three.
i) The operator must remain actively at the control box and navigate the towed electrofishing
unit.
ii) The probe operator will use one probe.
b) Set unit to pulsed DC and mark it in the Wave Form section of the Fish Gear Form.
c) Select the initial voltage setting based on the ambient conductivity of the river (i.e., not
corrected to 25 ฐC).
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.
d) Select the initial pulse rate and width.
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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.
e) If the electrofishing system only lets you adjust the percent of power, record the value on the
Fish Gear Form.
f) 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.
g) Once you have determined the appropriate settings, record them on the Fish Gear Form. Start
cleared clocks and resume fishing.
h) Note: some electrofishers do not meter all the requested settings; provide what you can.
i) If button time is not metered, estimate it with a stop watch and provide a comment in the NRSA
App.
5) Once the settings on the electrofisher are adjusted properly to sample effectively and minimize
injury and mortality, begin sampling the wadeable area at the downstream end and work upstream.
a) Depress the switch and slowly sweep the electrode from side to side sampling the wadeable
area.
b) In slack water areas (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.
c) In fast, shallow water, sweep the anode and fish downstream into a net.
d) 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.
e) Keep the cathode near the anode if fish catch is low.
6) 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.
7) The total button time for boat and wadeable electrofishing combined should be -700 seconds per
subreach.
8) After you finish sampling the wadeable area, return to Step 11 of Table 12.2.
a) You need to track the button time, total fishing time, and the length of stream sampled by
wading in each subreach. At the end of all sampling, record the total button time and total
fishing time expended for wadeable electrofishing, and the length of the fish sampling reach
that was sampled by wading in the Secondary Electrofishing Gear section of the Fish Gear
Form.
12.3.3 Secondary Seining
In waters where high conductivity would require the use of an electrofishing unit larger than
5000 V (e.g., a 7.5 or 9.0 GPP system), use seining as a secondary method in large wadeable ^
areas that are inaccessible to a boat. Use seining only as the last option for collecting fish at a ^
non-wadeable site. If conditions are such that seining is the only method used, provide a ^
justification in the comment bubble provided in the Sampling Protocol section at the top of the $
Fish Gear Form. Table 12.5 presents the procedure for seining large wadeable areas of
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subreaches. The intent of the supplemental seining is to provide comparable data to boat
electrofishing. Do not use seining to sample small "microhabitats" within a subreach.
Record all fish collected with seining protocols on separate lines on the data form from those
lines used for fish collected by electrofishing.
Although wadeable electrofishing techniques typically work best in an upstream direction,
seining may work best moving downstream. Depending upon habitat types and complexity, use
2 to 3 crew members. Two crew members move the seine; a third person creates and maintains
a bag in the seine in areas with higher velocities, or agitates rocks in riffles or snags. To avoid
mortality, process fish after each seine haul. Record all fish collected by seining 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: Secondary Seining Methods for Wadeable Areas (Non-Wadeable Rivers)
1) Use secondary seining as a last option only (e.g., when electrofishing is ineffective due to high
conductivity or extremely high turbidity). Area must be safely wadeable.
a) If site conditions are such that only seining is used, 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 required sampling reach.
i) At some sites, seining may be more effective while working downstream (from Transect A)
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 PFDs 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 ฐC.
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:
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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.
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.
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iii) Because of decreased complexity and shallower depths, seine hauls in glides or runs are
typically longer and use wider nets. You can use a 1.25 m deep seine in shallow glides.
iv) Pull the bag completely to shore at a predesignated point.
i) 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 near shore 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.
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, and 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.
b) 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) After you finish seining the wadeable area of the subreach, return to Step 11 of Table 12.3.
a) At the end of all sampling, use the seining information form to 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 Net
section of the Fish Gear Form.
e) Edge Habitats
f) Snag Habitats
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12.4 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.
12.4.1 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
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 12.4.6).
12.4.2 Unknown Specimens
If you cannot positively identify individuals to species in the field, record taxonomic information
of the collection form using scientific names rather than common names. If you can identify a
specimen only to family, record the scientific rather than the common family name (e.g.,
UNKNOWN PERCID A, not UNKNOWN PERCH A) on the Fish Collection Form 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
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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 12.4.6) or take good digital photographs (Section 12.4.3) for laboratory
identification. If you are only able to identify an unknown to Family, retain as many of the
individuals as possible for later identification. Use the UNK/RNG Voucher label(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 (Non-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, 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 river 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 flag and comment for the line
if you believe the population is stunted.
7) Examine each individual for external anomalies. Readily identify external anomalies including missing
organs (eye, fin), skeletal deformities, shortened operculum, eroded fins, irregular fin rays or scales,
tumors, lesions, ulcerous sores, blisters, cysts, blackening, white spots, bleeding or reddening,
excessive mucus, and fungus. After you process all of the individuals of a species, record the total
number 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 Tog# 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 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 formalinthe 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., NRS23_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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12.4.3 Photovouchering
Use digital imagery for fish species that cannot be preserved as voucher specimens (e.g., rare,
threatened, and endangered species; very large bodied). Ideally, take photos of all species
collected a site (that are not preserved) to provide a minimal level of documentation of
occurrence. Take photographs of entire specimens and additional specific morphological
features that are appropriate and necessary for an independent taxonomist to accurately
identify the specimen. Additional detail for these guidelines is provided in Stauffer et al. (2001),
which is provided to all Field Crews 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.
12.4.4 Preparing Preserved Voucher Specimen Samples
There are two different types of samples for preserved voucher specimens. The UNK/RNG
voucher samples are used to identify specimens that cannot be confidently identified in the
field, and to provide physical specimens of suspected range extensions. After submitting the
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 12.4.7) 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.
12.4.5 Preserving Voucher Specimen Samples
Preserve UNK/RNG and QA voucher specimens in the field with a 10% buffered formalin
solution. The volume of formalin must be equal to or greater than the total volume of
specimens. Use additional jars if necessary to ensure proper preservation. For individuals having
a total length larger than 200 mm (~8 in.), make a slit along the right side of the fish in the lower
abdominal cavity to allow penetration of the preservative solution. Follow all the precautions for
handling formalin outlined in the 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
UNK/RNG VOUCHER
Site ID: NRS23
/
_/202
/
_/202
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK RNG OUCHER
BAG
TAG: 41
iซG;
TAG: 43
TซG. h
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
TAG: 37
TAG: 38
TAG: 39
TAG: 40
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
TAG: 33
TAG: 34
TAG: 35
TAG: 36
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
TAG: 29
TAG: 30
Tflซ' 51
TAG: 32
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK.RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
TAG: 25
TAG: 26
TAG: 27
TAG: 28
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
TAG: 21
TAG: 22
TAG: 23
TAG: 24
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
TAG: 17
TAG: 18
TAG: 19
TAG: 20
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
IAซ: 13
TAG: 14
lAli: IB
1AU: IS
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
TAG: 09
TAG: 10
TAG: 11
TAG: 12
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
TAG: 05
TAG: 06
TAG: 07
TAG: 08
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/RNG VOUCHER
BAG
UNK/FNG OUCHER
BAG
TAC. 0.
TAG: 02
TAG: 03
TAC. 04
Figure 12.2 Unknown/Range Extension Voucher Sample Labels and Tags ^
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12.4.6 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 App to
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.
12.4.7 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 VC UCHER SAG
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
LU
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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 METHOD
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
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
Fine-tipped indelible markers for filling out sample labels
Clear tape strips for covering labels
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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
Blue catfish
Ictalurus furcatus
Flathead catfish
Pylodictis olivaris
Percidae
Sauger
Sander canadensis
Walleye
Sander vitreus
Yellow perch
Perca flavescens
Moronidae
White bass
Morone chrysops
Esocidae
Northern pike
Esox lucius
Chain pickerel
Esox niger
Muskellunge
Esox masquinongy
Salmonidae
Brown trout
Salmo trutta
Cutthroat trout
Oncorhynchus clarkii
Rainbow trout
Oncorhynchus mykiss
Brook trout
Salvelinus fontinalis
Largescale sucker
Catostomus macrocheilus
Quillback
Carpiodes cyprinus
CatostomidaeA
River carpsucker
Carpiodes carpio
River redhorse
Moxostoma carinatum
Smallmouth buffalo
Ictiobus bubalus
White sucker
Catostomus commersonii
Centrarchidae
Bluegill
Lepomis macrochirus
Rock bass
Ambloplites rupestris
Redbreast sunfish
Lepomis auritus
Redearsunfish
Lepomis microlophus
m
Northern pikeminnow
Ptychocheilus oregonensis
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Table 13.3 Procedure: Fish Tissue Plug Samples
Fish Tissue Plug Methods
1.
2.
5.
6.
7.
8.
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.
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.
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.
Remove one fish selected for plug sampling from the clean holding container(s) (e.g., livewell) using
clean nitrile gloves.
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).
Weigh the fish in grams using the fish weigh scale.
Note any anomalies (e.g., lesions, cuts, sores, tumors, fin erosion) observed on the fish.
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
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.
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Fish Tissue Plug Methods
15. Place the sample immediately on dry ice for shipment.
16. Dispose of gloves, scalpel and biopsy punch.
17. Keep the samples frozen on dry ice or in a freezer at <-20ฐC 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-o, 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 Figure 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 "I" 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
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: Processing Fecal Indicator (Enterococci) Sample
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.
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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).
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(s) 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 procedurefecal 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 river sample.
1. Set up sample filtration apparatus using same procedure as used for the river sample. Chill
Filter Extraction tubes with beads on dry ice.
2. Aseptically transfer 1 polycarbonate filter from filter box to base of opened Petri dish. Close
filter box and set aside (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).
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5. Measure 10 mL of the chilled phosphate buffered saline (PBS) in the sterile graduated PP tube
and pour into filter funnel.
6. Replace cover on filter funnel and pump to generate a vacuum (do not generate more than 7
inches of Hg of 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 j-j
<
The procedures for processing chlorophyll-a water samples are presented in Table 14.4. llj
Whenever possible, sample processing should be done in subdued light, out of direct sunlight.
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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
(do not cover the volume markings on the tube). Ensure that all written information is
complete and legible. Cover with a strip of clear tape.
11. Wrap the tube in aluminum foil and place in 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
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60 mL syringe with tip removed
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 (ฑ1 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 (ฑ1 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 ฑ1 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-a 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 (ฑ1 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 (ฑ1 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 ฑ1 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.
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
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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 ofPeriphyton 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
Load the boat on the trailer and inspect the boat, motor, and trailer for evidence of weeds and
other macrophytes. Clean the boat, motor, and trailer as completely as possible before leaving
the launch site. Inspect all nets for pieces of macrophyte or other organisms and remove as
much as possible before packing the nets for transport. Pack all equipment and supplies in the
vehicle and trailer for transport. Keep equipment and supplies organized so they can be
inventoried using the equipment and supply checklists presented in Appendix A. Lastly, be sure
to clean up all waste material at the launch site and dispose of or transport it out of the site if a
trash can is not available. ^
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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 revisit
sampling design is provided in Figure 15.1.
Revisit Sites (4 per state)
Visit 1
Space revisit events a minimum of
2 weeks and a maximum of 1
month apart
1
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 Revisit Sampling Design
15.2 Revisit Sampling Sites
A total of 192 (approximately 10%) of the target sites visited will be revisited during the same
field season year by the same Field Crew that initially sampled the site. Revisit samples and
O measurements are taken from the same reach as the first visit. Each state has four revisit sites:
cc
^ two likely wadeable and two likely non-wadeable sites. For each state these sites are:
O
<-> Wadeable Revisit sites:
The two likely wadeable revisit sites are resample sites from the NRSA 2018/19; one
g from the Large Stream (LS) panel and one from the Small Stream (SS) panel. The revisit
q sites are labeled as NRS23_18RVT2LS and NRS23_18RVT2SS, respectively, and are
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located in the Large and Small Streams tab 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
river sample is filtered and the filter blank will be sent to the lab along with the river samples.
Detailed description of the filter blanks 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.
If the Evaluator's findings indicate that the Field Crew is not performing the
Corrective procedures correctly, safely, or thoroughly, the Evaluator must continue working
^ctjon with this Field Crew until certain of the Crew's ability to conduct the sampling
Procedures 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.
ง 15.4 Reporting
O 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
Zi and problems (i.e., weaknesses [might affect data quality] or deficiencies [would adversely
3 affect data quality]). The Evaluator will ensure that the Crew understands the findings and will
Q be able to perform the procedures properly in the future. The Evaluator will record responses or
m concerns, if any, on the Field Evaluation and Assistance Check List. After the Evaluator
U_
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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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Stauffer, Dr. Jay R., J. Karish and T.D. Stecko. 2001. Guidelines for Using Digital Photos as Fish
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