National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
United States Environmental Protection Agency
Office of Water
Washington, DC
EPA-841-B-17-003a
National Rivers and Streams
Assessment 2018/19
Field Operations
Manual
Wadeable
Version 1.1
June 2018
-------
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
i
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
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 2018/19: Quality Assurance Project Plan EPA-841-B-17-
001
National Rivers and Streams Assessment 2018/19: Site Evaluation Guidelines EPA-841-B-17-003
National Rivers and Streams Assessment 2018/19: Field Operations Manual EPA-841-B-17-003a
National Rivers and Streams Assessment 2018/19: Laboratory Methods Manual EPA-841-B-17-
004
This document (Field Operations Manual (FOM)) contains a brief introduction and procedures to
follow at the base location and on-site, including methods for sampling water chemistry (grabs
and in situ measurements), periphyton, benthic macroinvertebrates, algal toxins, fish
assemblage, fish tissue plugs, whole fish tissue, Enterococci, and physical habitat. These
methods are based on the guidelines developed and followed in the National Rivers and Streams
Assessment 2008-2009 (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
2018/19. All Project Cooperators must follow these guidelines. Mention of trade names or
commercial products in this document does not constitute endorsement or recommendation for
use. Details on specific methods for site evaluation and sample processing can be found in the
appropriate companion document.
The suggested citation for this document is:
USEPA. 2017. National Rivers and Streams Assessment 2018/19: Field Operations Manual -
Wadeable. EPA-841-B-17-003a. U.S. Environmental Protection Agency, Office of Water
Washington, DC.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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 2
1.5 Supplemental Material to the Field Operations Manual 3
1.6 Recording Data and Other Information 4
2 INTRODUCTION TO WADEABLE SAMPLING 8
2.1 Daily Operations 8
2.2 Base Site Activities 10
2.2.1 Pre-departure Activities 10
2.2.2 Post Sampling Activities 12
2.3 Safety and Health 14
2.3.1 General Considerations 14
2.3.2 Safety Equipment 16
2.3.3 Safety Guidelines for Field Operations 16
2.4 Forms (Paper or Electronic) 18
2.4.1 Field Forms 18
2.4.2 Tracking Forms 18
2.4.3 Equipment and Supplies 19
3 INITIAL SITE PROCEDURES 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 26
3.1.4 Sampling During or After Rain Events 26
3.1.5 Site Photographs 26
3.2 Laying out the sampling reach 27
3.2.1 Sliding the Reach 30
3.3 Modifying Sample Protocols for High or Low Flows 31
3.3.1 Streams with Interrupted Flow 31
3.3.2 Braided Rivers and Streams 32
i/i
Z 4 WATER CHEMISTRY / CHLOROPHYLL-A SAMPLE COLLECTION AND PRESERVATION 34
LU '
I—
2 4.1 In Situ Measurements of Dissolved Oxygen, pH, Temperature, and Conductivity 34
u 4.1.1 Summary of Method 34
q 4.1.2 Equipment and Supplies 34
y 4.1.3 Sampling Procedure 35
^ 4.2 Water Chemistry Samples 37
i—
iv
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
4.2.1 Summary of Method 37
4.2.2 Equipment and Supplies 37
4.2.3 Water Chemistry and Chlorophyll-a Sampling Procedure 37
5 ALGAL TOXINS (MICROCYSTINS AND CYLINDROSPERMOPSIN) 40
5.1 Summary of Method 40
5.2 Equipment and Supplies 40
5.3 Sampling Procedure 41
6 BENTHIC MACROINVERTEBRATES 42
6.1 Summary of Method 42
6.2 Equipment and Supplies 42
6.3 Sampling Procedure 44
6.4 Sample Processing in Field 48
7 PERIPHYTON 50
7.1 Summary of Method 50
7.2 Equipment and Supplies 50
7.3 Sampling Procedure 50
7.4 Sample Processing in the Field 52
8 PHYSICAL HABITAT CHARACTERIZATION 53
8.1 Equipment and Supplies 53
8.2 Summary of Methods Approach 54
8.3 Components of the Habitat Characterization 54
8.4 Work Flow for the Physical Habitat Components 55
8.4.1 Thalweg Profile and Large Woody Debris Tally 55
8.4.2 Channel/Riparian Cross-Sections 55
8.4.3 Channel Constraint and Torrent Evidence 56
8.4.4 Stream Discharge 56
8.5 Habitat Sampling Locations within the Reach 56
8.5.1 Thalweg Profile and Large Woody Debris Measurements 59
8.6 Channel and Riparian Measurements at Cross-Section Transects 66
8.6.1 Slope and Bearing 66
8.7 Substrate Size and Channel Dimensions 73
8.7.1 Bank Characteristics 76
8.8 Canopy Cover Measurements 83
8.9 Visual Riparian Estimates 84
8.9.1 Riparian Vegetation Structure 84
8.10 Instream Fish Cover, Algae, and Aquatic Macrophytes 86
8.11 Human Influence 87
8.12 Cross-section Transects on Side Channels 89
8.13 Channel Constraint, Debris Torrents, Recent Floods, and Discharge 92
8.13.1 Channel Constraint 92
8.13.2 Debris Torrents and Recent Major Floods 95
8.14 Stream Discharge 98 £
8.14.1 Velocity-Area Procedure 98
8.14.2 Timed Filling Procedure 101 ^
8.14.3 Neutrally Buoyant Object Procedure 102 O
8.15 Elevation atTransect K 103 u.
O
9 FECAL INDICATOR (ENTEROCOCCI) 104 ^
CO
9.1 Summary of Method 104 k
V
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
9.2 Equipment and Supplies 104
9.3 Sampling Procedure 104
9.4 Sample Processing in the Field 105
10 FISH ASSEMBLAGE 106
10.1 Summary of Method 106
10.2 Equipment and Supplies 109
10.3 Sampling Procedures 109
10.3.1 Irruptive Species 110
10.3.2 Small Wadeable Streams 113
10.3.3 Large Wadeable Streams 116
10.4 Seining 119
10.5 Processing Fish 123
10.5.1 Identification and Tallying 124
10.5.2 Unknown Specimens 124
10.5.3 Photovouchering 128
10.5.4 Preparing Preserved Voucher Specimen Samples 128
10.5.5 Preserving Voucher Specimen Samples 129
10.5.6 Processing Unknown/Range Extension (UNK/RNG) Voucher Samples 132
10.5.7 Processing QA Voucher Samples 133
11 FISH TISSUE PLUG SAMPLING METHODS 139
11.1 Method Summary 139
11.2 Equipment and Supplies 139
11.3 Sample Collection Procedures 140
12 WHOLE FISH SAMPLING METHOD 143
12.1 Method Summary 143
12.2 Equipment and Supplies 143
12.3 Sampling Procedures 144
13 FINAL SITE ACTIVITIES 148
13.1 Overview of Final Site Activities 148
13.2 General Site Assessment 149
13.2.1 Elevation at Transect K 149
13.2.2 Watershed Activities and Disturbances Observed 149
13.2.3 Site Characteristics 149
13.2.4 General Assessment 149
13.3 Processing the Fecal Indicator (Enterococci), Chlorophyll-a, and Periphyton Samples 152
13.3.1 Equipment and Supplies (Fecal Indicator Filtering) 152
13.3.2 Procedures for Processing the Fecal Indicator (Enterococci) Sample 152
13.3.3 Equipment and Supplies (Chlorophyll-a from Water Sample Filtering) 154
13.3.4 Procedures for Processing the Chlorophyll-a Water Sample 154
13.3.5 Equipment and Supplies (Periphyton Sample) 155
13.3.6 Procedures for Processing the Periphyton Samples 156
£ 13.4 Data Forms and Sample Inspection 160
^ 13.5 Launch Site Cleanup 160
H
g 14 FIELD QUALITY CONTROL 161
u
lj- 14.1 Revisit Sampling Overview 161
O
lu 14.2 Revisit Sampling Sites 162
m 14.3 Field Evaluation and Assistance Visits 163
H 14.3.1 Specifications for QC Assurance Field Assistance Visits 163
vi
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
14.4 Reporting 164
15 REFERENCES 165
APPENDIX A LIST OF EQUIPMENT AND SUPPLIES A-l
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
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 Wadeable Sites 9
Figure 2.2 Overview of Base Site Activities 10
Figure 2.3 Electronic Request Form 20
Figure 3.1 Verification Form (front) 24
Figure 3.2 Verification Form (back) 29
Figure 3.3 Sampling Reach Features for a Wadeable Site 30
Figure 4.1 Field Measurement Form 36
Figure 4.2 Sample Collection Form (front) 39
Figure 6.1 Sample Collection Form (back) 43
Figure 6.2 Benthic Macroinvertebrate Collection at Wadeable Sites 44
Figure 6.3 Transect Sample Design for Collecting Benthic Macroinvertebrates at Wadeable Sites 45
Figure 8.1 Reach Layout for Physical Habitat Measurements for streams 2.5m or greater (plan view) 58
Figure 8.2 Thalweg Profile and Woody Debris Form 62
Figure 8.3 Large Woody Debris Influence Zones (modified from Robison and Beschta, 1990) 66
Figure 8.4 Slope and Bearing Form 69
Figure 8.5 Channel Slope Measurement using a Clinometer 71
Figure 8.6 Measurements of Bearing Between Transects 73
Figure 8.7 Substrate Sampling Cross-Section 74
Figure 8.8 Channel Riparian Cross-Section Form 77
Figure 8.9 Determining Bank Angle Under Different Types of Bank Conditions 79
Figure 8.10 Schematic Showing Relationship Between Bankfull Channel and Incision 81
Figure 8.11 Determining Bankfull and Incision Heights 82
Figure 8.12 Schematic of Modified Convex Spherical Canopy Densiometer 83
Figure 8.13 Riparian Zone and Instream Fish Cover Plots for a Stream Cross-Section Transect 85
Figure 8.14 Proximity Classes for Human Influences in Wadeable Streams 88
Figure 8.15 Riparian and Instream Fish Cover Plots for a Stream with Minor and Major Side Channels .... 90
Figure 8.16 Channel/Riparian Cross-Section Form for an Additional Major Side Channel Transect 91
Figure 8.17 Channel Constraint Form 94
Figure 8.18 Types of Multiple Channel Patterns 95
Figure 8.19 Torrent Evidence Assessment Form 97
Figure 8.20 Layout of Cross-Section for Obtaining Discharge Data by the Velocity-Area Procedure 98
Figure 8.21 Discharge Form 100
Figure 8.22 Use of a Portable Weir and Calibrated Bucket to Obtain an Estimate of Stream Discharge... 101
Figure 10.1 Fish Gear and Sampling Information Form (front) 107 3
Figure 10.2 Fish Collection Form 108 ^
Figure 10.3 Reach Layouts for Fish Sampling at Wadeable Sites 112 ^
Figure 10.4 Seining Information Form 123 |_
Figure 10.5 Unknown/Range Extension Voucher Sample Labels and Voucher Specimen Tags 130 zi
vii
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
Figure 10.6 Fish Gear and Sampling Information Form (back) 131
Figure 10.7 QA Voucher Sample Labels and Voucher Specimen Tags 134
Figure 10.8 Fish Collection Revision Form (Page 1) 136
Figure 10.9 Fish Collection Revision Form (Page 2) 137
Figure 12.1 Whole Fish Tissue Collection Form 147
Figure 13.1 Final Site Activities Summary 148
Figure 13.2 Site Assessment Form (front) 150
Figure 13.3 Site Assessment Form (back) 151
Figure 14.1 Summary of the Repeat Sampling Design 161
Figure 14.2 Summary of Fish Tissue Protocol for Revisit Sites 162
Table 1.1 Summary Table of Indicators for all NRSA 2018/19 Sites 3
Table 1.2 Guidelines for Recording Field Measurements and Tracking Information 5
Table 2.1 Stock Solutions, Uses, and Methods for Preparation 12
Table 2.2 Post-sampling Equipment Care 13
Table 2.3 General Health and Safety Considerations 15
Table 2.4 General Safety Guidelines for Field Operations 17
Table 3.1 Equipment and Supplies: for Site Verification 22
Table 3.2 Procedure: Site Verification 25
Table 3.3 Guidelines to Determine the Influence of Rain Events 26
Table 3.4 Procedure: Laying Out the Sampling Reach at Wadeable Sites 27
Table 3.5 Procedure: Sliding the Sampling Reach 31
Table 3.6 Reach Layout Modifications for Interrupted Streams 32
Table 3.7 Procedure: Modifications for Braided Rivers and Streams 33
Table 4.1 Equipment and Supplies: DO, pH, Temperature, and Conductivity 34
Table 4.2 Procedure: Temperature, pH, Conductivity and Dissolved Oxygen 35
Table 4.3 Equipment and Supplies: Water Chemistry Sample Collection and Preservation 37
Table 4.4 Procedure: Water Chemistry and Chlorophyll-a Sample Collection (Wadeable Sites) 38
Table 5.1 Equipment and Supplies: Microcystin 40
Table 5.2 Procedure: Algal Toxin (Microcystin and Cylindrospermopsin) Collection (Wadeable Sites) 41
Table 6.1 Equipment and Supplies: Benthic Macroinvertebrate Collection at Wadeable Sites 42
Table 6.2 Procedure: Benthic Macroinvertebrates (Wadeable Sites) 46
Table 6.3 Procedure: Compositing Samples for Benthic Macroinvertebrates (Wadeable Sites) 48
Table 7.1 Equipment and Supplies: Periphyton (Wadeable Sites) 50
Table 7.2 Procedure: Collecting Composite Index Samples of Periphyton (Wadeable Sites) 51
Table 8.1 Equipment and Supplies: Physical Habitat 53
Table 8.2 Summary of Components of Physical Habitat Characterization at Wadeable Sites 54
Table 8.3 Procedure: Thalweg Profile 60
Table 8.4 Channel Unit Categories 64
Table 8.5 Procedure: Tallying Large Woody Debris 65
Table 8.6 Procedure: Obtaining Slope Data 67
Table 8.7 Modified Procedure: Obtaining Slope Data (without Surveyor's Level) 70
Table 8.8 Procedure: Obtaining Bearing Data 72
Table 8.9 Procedure: Substrate Measurement 75
Table 8.10 Procedure: Measuring Bank Characteristics 78
^ Table 8.11 Procedure: Canopy Cover Measurements 84
CO
< Table 8.12 Procedure: Characterizing Riparian Vegetation Structure 86
ll. Table 8.13 Procedure: Estimating Instream Fish Cover 87
^ Table 8.14 Procedure: Estimating Human Influence 89
^ Table 8.15 Procedure: Assessing Channel Constraint 92
viii
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
Table 8.16 Procedure: Determining Stream Discharge-Velocity-Area 99
Table 8.17 Procedure: Determining Stream Discharge-Timed Filling 102
Table 8.18 Procedure: Determining Stream Discharge - Neutrally Buoyant Object 103
Table 9.1 Equipment and Supplies: Fecal Indicator Sampling (Wadeable Sites) 104
Table 9.2 Procedure: Fecal Indicator (Enterococci) Sample Collection (Wadeable Sites) 104
Table 10.1 Equipment and Supplies: Fish Collection (Wadeable Sites) 109
Table 10.2 Summary of Wadeable Fishing Protocols Ill
Table 10.3 Procedure: Electrofishing (Small Wadeable Streams) 113
Table 10.4 Procedure: Electrofishing (Large Wadeable Sites) 116
Table 10.5 Procedure: Seining (Wadeable Sites) 120
Table 10.6 Procedure: Processing Fish (Wadeable Sites) 125
Table 10.7 Procedure: Processing Unknown/Range Extension (UNK/RNG) Voucher Samples 135
Table 10.8 Procedure: Processing QA Voucher Samples 138
Table 11.1 Equipment and Supplies: Fish Tissue Plug Sample 140
Table 11.2 Recommended Target and Alternate Species for Fish Tissue Plug Collection 141
Table 11.3 Procedure: Fish Tissue Plug Samples 141
Table 12.1 Equipment and Supplies: Whole Fish Tissue Sample Collection 144
Table 12.2 Recommended Target Species for Whole Fish Tissue Collection 145
Table 12.3 Sampling Procedures for Whole Fish Tissue Samples 145
Table 13.1 Equipment and Supplies: Fecal Indicator (Enterococci) Sample Processing 152
Table 13.2 Procedure: Fecal Indicator (Enterococci) Sample Processing 152
Table 13.3 Equipment and Supplies: Chlorophyll-a Processing 154
Table 13.4 Procedure: Chlorophyll-a Sample Processing 154
Table 13.5 Equipment and Supplies: Periphyton Samples 155
Table 13.6 Procedure: ID/Enumeration Samples of Periphyton 156
Table 13.7 Procedure: Preparing Metagenomic Sample of Periphyton 157
Table 13.8 Procedure: Preparing Chlorophyll Samples of Periphyton 158
Table 13.9 Procedure: Preparing Ash-Free Dry Mass (AFDM) Samples of Periphyton 159
Table 14.1 General Information Noted During Field Evaluation 164
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
<
>
LU
QC
CO
CO
<
O
QC
u
<
AFDM
ANC
Ca
CI
CPR
CWA
DELT
DO
DOC
Dl
EPA
FOM
GIS
GPP
GPS
IBI
IM
K
LIMS
LOM
LWD
Mg
MMI
MSDS
Na
NAD
NARS
NHD
NH3
nh4
NIST
N02
N03
NRSA
O/E
OSHA
OW
PETG
PFD
PP
PPT
Psig
QA
QAPP
QA/QC
QCS
ORG
SEG
Si02
S04
SOP
Ash-Free Dry mass
Acid-neutralizing capacity
Calcium
Chloride
Cardiopulmonary Resuscitation
Clean Water Act
Deformities, Eroded Fins, Lesions and Tumors
Dissolved Oxygen
Dissolved Organic Carbon
Di-ionized Tap Water
Environmental Protection Agency
Field Operations Manual
Geographic Information System
Generator Powered Pulsator
Global Positioning Device
Index of Biotic Integrity
Information Management
Potassium
Laboratory Information Management System
Lab Operations Manual
Large Woody Debris
Magnesium
Multimetric Indicators
Methods Safety Data Sheets
Sodium
North American Datum
National Aquatic Resources Survey
National Hydrology Database
Ammonia
Ammonium
National Institute of Standards and Technology
Nitrite
Nitrate
National Rivers and Streams Assessment
"Observed" over "Expected"
Occupational Safety and Health Administration
Office of Water
polyethylene terephthalate glycol
Personal Flotation Device
Polypropylene
Parts per thousand
Pounds per square inch - gauge
Quality Assurance
Quality Assurance Project Plan
Quality Assurance/Quality Control
Quality Control Solution
Quick Reference Guide
Site Evaluation Guideline
Silica
Sulfate
Standard Operating Procedures
X
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Std
Standard
TL
Total Length
TOC
Total Organic Carbon
TN
Total Nitrogen
TP
Total Phosphorus
TSS
Total Suspended Solids
UNK/RNG
Unknown Range Extension
USGS
United States Geological Survey
WRS
Willamette Research Station
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
DISTRIBUTION LIST
This FOM and associated manuals or guidelines will be distributed to the following U.S.
Environmental Protection Agency (EPA) senior staff participating in the NRSA and to State Water
Quality Agencies or cooperators who will perform the field sampling operations. The Quality
Assurance (QA) Officers will distribute the Quality Assurance Project Plan (QAPP) and associated
documents to participating project staff at their respective facilities and to the project contacts
at participating laboratories, as they are determined.
National Monitoring Coordinators
Richard Mitchell
NRSA Project Leader
mitchell.richard(5) epa.gov
202-564-0064
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Sarah Lehman
NRSA Project QA
Officer
lehmann.sarah (Sena.gov
202-566-1379
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Cynthia N. Johnson
OWOW QA Officer
iohnson.cvnthiaN (Sena.gov
202-566-1679
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Bernice L. Smith
OWOW QA
Coordinator
smith, bernicel (Sena.gov
202-566-1244
U.S. EPA Office of Water
1200 Pennsylvania Ave., NW
Washington, DC 20460
Steven G. Paulsen
EPA ORD Technical
Advisor
Paulsen.steve(6)epa.gov
541-754-4428
Freshwater Ecology Branch Western Ecology Division, NHEERL,
ORD, EPA
200 S.W. 35th St. Corvallis, OR 97330
Marlys Cappaert
NARS Information
Management
Coordinator
cappaert.marlys@epa.gov
541-754-4467
541-754-4799 (fax)
SRA International Inc
200 S.W. 35th Street
Corvallis, OR 9733
Chris Turner
Contract Field Logistics
Coordinator
cturner@glec.com
715-829-3737
Great Lakes Environmental Center, Inc.
739 Hastings Street
Traverse City, Ml 49686
Leanne Stahl
OST Fish Tissue
Coordinator
Stahl.leanne@epa.gov
202-566-0404
U.S. EPA Office of Water
Office of Science and Technology
1200 Pennsylvania Ave., NW
Washington, DC 20460
Regional Monitoring Coordinators
Z>
CO
cc
I—
un
O
XII
Tom Faber
Region 1
Faber.torn (5) epa.gov
617-918-8672
U.S. EPA-Region 1
11 Technology Drive North Chelmsford, MA 01863-2431
Emily Nering
Region 2
nering.emilv@epa.gov
732-321-6764
U.S. EPA-Region II
2890 Woodbridge Ave Edison, NJ 08837-3679
Bill Richardson
Region 3
richardson.william@epa.gov
215-814-5675
U.S. EPA-Region III
1650 Arch Street, Philadelphia, PA 19103-2029
Elizabeth Belk
Region 4
belk.elizabeth@epa.gov
404-562-9377
U.S. EPA - Region IV
61 Forsyth Street, S.W. Atlanta, GA 30303-8960
Mari Nord
Region 5
nord. mari (5) epa.gov
312-353-3017
U.S. EPA - Region V
77 West Jackson Blvd Chicago, IL 60604-3507
Rob Cook
Region 6
cook.robert@epa.gov
214-665-7141
U.S. EPA-Region VI
1445 Ross Ave -Ste 1200 Dallas, TX 75202-2733
Gary Welker
Region 7
welker. ga rv@ epa.gov
913-551-7177
U.S. EPA-Region VII
300 Minnesota Ave, Kansas City, KS 66101
Tom Johnson
Region 8
iohnson.tom@epa.gov
303-312-6226
U.S. EPA-Region VIII
1595 Wynkoop St .Denver, CO 80202-1129
Matthew Bolt
Region 9
bolt.matthew@epa.gov
415-972-3578
U.S. EPA-Region IX
75 Hawthorne Street San Francisco, CA 94105
Lillian Herger
Region 10
Herger.lillian@epa.gov
206-553-1074
U.S. EPA-Region X,
1200 Sixth Avenue Seattle, WA 98101
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
1 BACKGROUND
This manual describes field protocols and daily operations for crews to use in the Wadeable
NRSA 2018/19 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 both the NRSA 2008/09 and NRSA 2013/14.
• Help build State and Tribal capacity for monitoring and assessment and promote
collaboration across jurisdictional boundaries.
This is one of a series of water assessments being conducted by states, tribes, the U.S. EPA, and
other partners. In addition to rivers and streams, the water assessments will also focus on
coastal waters, lakes, and wetlands in a revolving sequence. The purpose of these assessments
is to generate statistically valid reports on the condition of our Nation's water resources and
identify key stressors to these systems.
1.1 Survey Design
The survey design consists of two separate designs to address the dual objectives of: (1)
estimating current status and (2) estimating change in status for all flowing waters:
• Resample design applied to NRSA 2008/09 and NRSA 2013/14 sites.
• New site design for NRSA 2018/19.
The survey design is explicitly stratified by state for both designs. The unequal probability
categories are specific to the survey design used for the NRSA 2008/09, NRSA 2013/14, and
NRSA 2018/19. In all cases the categories are specific combinations of Strahler order categories
and nine National Aquatic Resource Survey (NARS) aggregated ecoregions. In addition, a
minimum of 20 sites (Resample and New) was guaranteed in each state and a maximum of 75
sites was the limit for an individual state. There are 983 unique sites in the Resample Design and
825 unique sites in the New Site Design. Approximately 10% of the total NRSA sites are
scheduled for repeated sampling (revisit sites) in the same year of the two year NRSA field cycle.
The sample frame was derived from the medium 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: 2018/19 documents.
1.2 Target Population and Index Period
The target population consists of all streams and rivers within the 48 contiguous states that
have flowing water during the study index period, including major rivers and small streams. Sites
must have > 50% of the reach length with standing water, and sites with water in less than 50%
of the reach length must be dropped. All sites must be sampled during base flow conditions.
The target population excludes:
• Tidal rivers and streams up to head of salt (defined as < 0.5 ppt for this study).
• Run-of-the-river ponds and reservoirs with greater than seven day residence time.
The study index period extends from:
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
o Beginning of June through end of September for most regions.
o Sites in the select ecoregions or States can be sampled starting in the end of
April with approval from the EPA Project Coordinator.
Please refer to the Site Evaluation Guidelines (EPA-841-B-17-002 and the NRSA Web site
http://www.epa.gov/national-aquatic-resource-surveys/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
unsampleable, it must be replaced by another site within the same state and same panel. The
five panels for NRSA 2018/19 are:
• NRS18_08TS3R2: sites from NRSA 2008/09 that were sampled twice in 2008/09 and
then sampled twice again in 2013/14 (with a few exceptions).
• NRS18_08TS3: sites from NRSA 2008/09 that were sampled once in 2008/09 and then
sampled again in 2013/14.
• NRS18_13TS2R2: sites from NRSA 2013/14 that were sampled twice in 2013/14.
• NRS18_13TS2: sites from NRSA 2013/14 that were sampled once in 2013/14 and will be
sampled again in 2018/19.
• NRS18_18: new sites selected for NRSA 2018/19 that will be sampled once in 2018/19.
If the site is from the New Site Design (panel NRS18_18), then the replacement site must also be
within the same ecoregion and in the same size category. The four general categories are:
• Small streams (SS): 0-2 Strahler order sites.
• Large streams (LS): 3-4 Strahler order sites.
• Major rivers (RM): 5 and above Strahler order sites.
• Other rivers (RO): 5 and above Strahler order sites not considered Major Rivers.
Please refer to the Site Evaluation Guidelines (EPA-841-B-17-002) 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 2018/19
field season. Key evaluation criteria included indicator applicability on a national scale, the
ability of an indicator to reflect various aspects of ecological condition, repeatability, and cost-
effectiveness. The core indicators build upon the work done in the NRSA 2008/09 and NRSA
2013/14. 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
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
purposes, there is no distinction between core and supplemental indicators. Indicators that are
included in the NRSA 2018/19 are briefly described in Table 1.1.
Table 1.1 Summary Table of Indicators for all NRSA 2018/19 Sites
Indicator
Core or Supplemental
Indicator
Specs/Location in Sampling Reach
In Situ measurements (pH, DO,
temperature, conductivity)
Core Indicator
Measurements taken at X site at mid-
channel; readings are taken at 0.5 m
depth, or mid-depth if water depth is less
than 1 meter.
Water chemistry (TP, TN, NH3-
N, NO3-NO2, N03, basic anions
and cations, silica, alkalinity
[Acid-neutralizing capacity
(ANC)], DOC, TOC, TSS,
conductivity, pH, turbidity, true
color)
Core Indicator
Collected at the X site at mid-channel;
from a depth of 0.5 m, or mid-depth if
water depth is less than 1 meter.
Chlorophyll-o
Core Indicator
Collected as part of water chemistry and
periphyton samples
Microcystin and
cylindrospermopsin
Supplemental Indicator
Collected at the X site at mid-channel;
from a depth of 0.5 m, or mid-depth if
water depth is less than 1 meter.
Periphyton composite and
periphyton metagenomic
Supplemental Indicator
Collected from 11 locations
systematically placed at each site and
combined into a single composite sample
Benthic macroinvertebrate
assemblage (Littoral)
Core Indicator
Collected from 11 locations
systematically placed at each site and
combined into a single composite sample
Fish Assemblage
Core Indicator
Sampled throughout the sampling reach
at specified locations
Physical habitat assessment
Core Indicator
Measurements collected throughout the
sampling reach at specified locations
Fecal indicator (Enterococci)
Supplemental Indicator
Collected at the last transect one meter
off the bank at 0.3 m depth
Fish Tissue Plug
Supplemental Indicator
Target species collected throughout the
sampling reach as part offish
assemblage sampling
Whole Fish Tissue
Supplemental Indicator at
select sites
Target species collected throughout the
sampling reach as part offish
assemblage sampling
3
1.5 Supplemental Material to the Field Operations Manual o
The FOM describes wadeable field protocols and daily operations for crews to use in the NRSA. g
Following these detailed protocols will ensure consistency across regions and reproducibility for ^
3
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
future assessments. Before beginning sampling at a site, crews should prepare a packet for each
site containing pertinent information to successfully conduct sampling. This includes a road map
and set of directions to the site, topographic maps, landowner access forms, sampling permits
(if needed), site evaluation forms, and other information necessary to ensure an efficient and
safe sampling day.
Field Crews may request to receive a Quick Reference Guide (QRG) that contains tables and
figures summarizing field activities and protocols from the FOM. This waterproof handbook will
be the primary field reference used by Field Crews after reading the FOM and completing the
required field training session. The QRG will also be made available to all crews in electronic
(Adobe® PDF) format. The Field Crews are also required to keep the FOM available 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
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-017-001).
Related NRSA documents include the following: National Rivers and Streams Assessment:
Quality Assurance Project Plan (EPA 841-B-17-001), National Rivers and Streams Assessment:
Site Evaluation Guidelines (EPA 841-B-017-002), and National Rivers and Streams Assessment:
Laboratory Methods Manual (EPA 841-B-017-004). 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, consistently, and
legibly. The cost of a sampling visit coupled with the short index period severely limits the ability
to resample a site if the initial information recorded was inaccurate or illegible. Guidelines for
recording field measurements are presented in Table 1.2. At the end of each sampling day, the
Field Crew Leader is responsible for reviewing each field form for completeness and legibility.
The Field Crew Leader must initial each field form after reviewed or if using the App, hit the
reviewed button at the top of each electronic form.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 1.2 Guidelines for Recording Field Measurements and Tracking Information
Activity Guidelines
Field Measurements
Data Recording
• Record measurement values and observations within the NRSA 2018/19 app
developed for use on EPA-issued tablet devices.
• If unable to record data within the NRSA 2018/19 app, then record on data
forms preprinted by the IM Team on water-resistant paper.
o Use No. 2 pencil only (fine-point indelible markers can be used if
necessary) to record information on forms.
o Record data and information using correct format as provided on
data forms.
o Be sure to accurately record site IDs and sample numbers. For all
primary sampling visits indicate the event as Visit 1. For revisit sites
use Visit 2 to indicate the second sampling event during the same
year.
o Print legibly (and as large as possible). Clearly distinguish letters
from numbers (e.g., 0 versus 0, 2 versus Z, 7 versus T or F, etc.), but
do not use slashes.
o In cases where information is recorded repeatedly on a series of
lines (e.g., physical habitat characteristics), do not use "ditto marks"
(") or a straight vertical line. Fill in all data even if there is repetition
between subsequent lines.
o When recording comments, print or write legibly. Make notations in
comments field only; avoid marginal notes. Be concise, but avoid
using abbreviations or "shorthand" notations. If you run out of
space, attach a sheet of paper with the additional information,
rather than trying to squeeze everything into the space provided on
the form.
Data Qualifiers
(Flags)
Use only defined flag codes and record on data form in appropriate field.
K = Measurement not attempted or not recorded.
Q = Failed quality control check; re-measurement not possible.
U = Suspect measurement; re-measurement not possible.
Fn = Miscellaneous flags (n = 1, 2, etc.) assigned by a Field Crew during a
particular sampling visit (also used for qualifying samples).
Explain reason for using each flag in comments section on data form.
Sample Labels
• Use adhesive labels with preprinted sample ID numbers and follow the
standard recording format for each type of sample.
• Use a fine point indelible marker to record information on label. Cover the
completed label with clear tape.
• Record sample ID number from label and associated collection information in
the app or on sample collection form preprinted on water-resistant paper.
• Sample IDs from a single label sheet are in sequential order. Do not mix
labels from different sheets.
Q
Z
3
o
cc
u
<
CO
5
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Activity Guidelines
Sample Collection and Tracking
Sample Qualifiers
(Flags)
Use only defined flag codes and record on sample collection form in appropriate
field.
K = Sample not collected or lost before shipment; resampling not
possible.
U = Suspect sample (e.g., possible contamination, does not meet
minimum acceptability requirements, or collected by non-standard
procedure).
Fn = Miscellaneous flags (n=l, 2, etc.) assigned by a Field Crew during a
particular sampling visit (also used for field measurements).
Explain reason for using flags in "Comments" on sample collection form.
Review of Labels
and Data
Collection Forms
• Compare information recorded on labels and sample collection form for
accuracy before leaving site.
• Review labels and data collection forms for accuracy, completeness, and
legibility before leaving site.
• The Field Crew Leader must review all labels and data collection forms for
consistency, correctness, and legibility before transfer to the Information
Management Center.
Q
Z
3
o
QC
u
<
CO
6
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
WATER CHEMISTRY (CHEM)
Site ID: NRS18
Date: /201 Visit #:01 02
ti 990000
WCHL, PCHL, PBIO - OUTER BAG
Site ID: NRS18
Date: / /201 Visit#: 01 02
990001,990003, 990004
T1
PERIPHYTON CHLOROPHYLL (PCHL)
Site ID: NRS18
Date: I /201 Visit #:01 02
Volume Filtered: mL
990003
PERIPHYTON METAGENOMIC (PDNA)
Site ID: NRS18
Date: / /201 Visit#: 01 02
i2 990005
ALGAL TOXIN (MICZ)
(HDPE Bottle (round Nalgene bottle)
Site ID: NRS18
Date: / /201 Visit #:01 02
T2 990007
BENTHIC MACROINVERTEBRATES (BETB)
Site ID: NRS18
Date: / /201 Visit#: 01 02
T3 990009
QA VOUCHER (VERT)
Site ID: NRS18
Date: /201 Visit#: 01 02
T3 990012
CHLOROPHYLL-a (WCHL)
Site ID: NRS18
Date: / /201 Visit#: 01 02
Volume Filtered: mL
ti 990001
PERIPHYTON ASSEMBLAGE ID (PERI)
Site ID: NRS18
Date: / .201 Visit#: 01 02
Composite Volume: mL
990002
PERIPHYTON BIOMASS (PBIO)
Site ID: NRS18
Date: / /201 Visit#: 01 02
Volume Filtered: mL
990004
ALGAL TOXIN (MICX)
(PETG Bottle (clear, square bottle)
Site ID: NRS18_
Date: / /201 Visit#: 01 02
T2 990006
BENTHIC MACROINVERTEBRATES (BERW)
Site ID: NRS18
Date: / /201 Visit#: 01 02
ts 990008
FISH TISSUE PLUG (FPLG)
Site ID: NRS18
Date: / /201 Visit#: 01 02
T2 990011
UNK/RNG VOUCHER (VERT)
Site ID: NRS18
Date: / /201 Visit#: 01 02
Figure 1.1 Example Sample Labels for Sample Tracking and Identification.
Q
Z
3
o
az
e?
(j
<
CO
7
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
2 INTRODUCTION TO WADEABLE SAMPLING
2.1 Daily Operations
Field methods for the NRSA are designed to be completed in one field day for most sites.
Depending on the time needed for both the sampling and travel for the day, an additional day
may be needed to complete sampling or for pre-departure and post-sampling activities (e.g.,
cleaning equipment, repairing gear, shipping samples, and traveling to the next site). Remote
sites with lengthy or difficult approaches may require more time, and Field Crews will need to
plan accordingly.
A Field Crew for a wadeable site will typically consist of four people. Any additional crew
members may either help collect samples, or may remain on the bank to provide logistical
support. Typically, in wadeable sites, two crew members will work on the "habitat" crew, and
two or three will work on the "fish" crew.
A daily field sampling scenario showing how the work load may be split between crew members
is presented in Figure 2.1. The following sections further define the sampling sequence and the
protocols for sampling activities.
Field Crews should define roles and responsibilities for each crew member to organize field
activities efficiently. While crews may choose to allocate resources as they see fit, the sequence
of sampling events presented in the Figure 2.1 cannot be changed and is based on the need to
protect some types of samples from potential contamination and to minimize holding times
once samples are collected. For example, at sites where fish collections are expected to take
longer than physical habitat assessments, crews may choose to task Group A with in situ
measurements and water collections, but these tasks need to be completed before any tasks
near or directly upstream of the water collection are to begin.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Whole Crew
Locate X-site
Verify site as target
Set upstaging area
I
Group A Activities 1
| Group B Activities
Prepare forms, equipment & supplies
Calibrate multi-parameter probe
1
1
Lay out sampling reach (from X-site to Transect A)
Lay out sampling reach (from X-site to Transect K)
i
1
Begin sampling activities at Transect A
Return to transect F (X-site)
Conduct physical habitat
characterizations
Collect benthic samples
Collect periphyton samples
Collect fecal indicator
sample at last transect
Return to staging area
Preserve benthic
samples & prepare for
transport
Inspect and
decontaminate boat
Filter enterococci
sample; prepare for
transport
I
Filter chlorophyll a
samples; prepare for
transport
Clean & organize equipment
Measure in situ temperature,
pH, DO, & conductivity
I
Collect water chemistry,
chlorophyll-a, & algal toxin
samples
UE
Travel to Transect A
I
Conduct fish assessment
I
Collect fish tissue samples
~r
Return to staging area
I£
Prepare fish tissue samples for transport
4
Review data forms
I
Inventory supplies; request more
supplies if needed
Re port sam p I i ng eve nt thro ugh
Site and Sample Status form
Ship samples
<
CO
CD
<
LU
Q
<
Figure 2.1 Field Sampling Scenario for Wadeable Sites
O
H
Z
o
I—
u
Z>
Q
O
QC
9
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
2.2 Base Site Activities
Field Crews conduct a number of activities at their base site (i.e., office or laboratory, camping
site, or motel). These include tasks that must be completed both before departure to the site
and after return from the field (Figure 2.2). Close attention to these activities is required to
ensure that the Field Crews know: (1) where they are going, (2) that access is permissible and
possible, (3) that equipment and supplies are available and in good working order to complete
the sampling effort, and (4) that samples are packed and shipped appropriately.
PREDEPARTURE ACTIVITIES
Crew Leader
• Prepare daily itinerary
Crew Members
• Instrument checks & calibration
• Equipment & supplies preparation
Whole Crew
Site verification
SAMPLE SITE
<
i/i
CO
<
LU
Q
<
U
3
Q
o
QC
POST-SAMPLING ACTIVITIES
Crew Leader
• Review forms & labels
• File status report by email to
IM Team
Crew Members
Filter, preserve & inspect samples
Clean boals/gear with 1-10%
bleach solution
Make any repairs necessary
Charge or replace batteries
Refuel boals, vehicles, etc.
Obtain ice, dry ice and other con-
sumables.
Package and ship samples and
data forms
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.
10
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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.
2.2.1.2 Instrument Checks and Calibration
Each Field Crew must test and calibrate instruments prior to sampling. Calibration can be
conducted prior to departure for the site or at the site, with the exception of dissolved oxygen
(DO) calibration. Because of the potential influence of altitude, DO calibration is to be
performed only at the site. Field instruments include a global positioning system (GPS) receiver,
a multi-probe unit for measuring DO, pH, temperature, and conductivity, and electrofishing
equipment. Field Crews should have access to backup instruments if any instruments fail the
manufacturer performance tests or calibrations. Prior to departure, Field Crews must:
• Turn on the GPS receiver and check the batteries. Replace batteries immediately if a
battery warning is displayed.
• Test and calibrate the multi-probe meter. Each Field Crew should have a copy of the
manufacturer's calibration and maintenance procedures. All meters should be
calibrated according to manufacturer specifications provided along with the meter.
• Turn on the electrofishing unit and check the batteries. Be sure to have fully charged
backup batteries. If using a gas powered electrofishing unit, check the oil and gas supply.
2.2.1.3 Equipment and Supply Preparation
Field Crews must check the inventory of supplies and equipment prior to departure using the
equipment and supplies checklists provided in Appendix A; use of the lists is mandatory. Specific
equipment will be used for wadeable or non-wadeable sites; be sure to bring both sets of
equipment if you are unsure what type of site you will be visiting that day. Pack meters, probes,
and sampling gear in such a way as to minimize physical shock and vibration during transport.
Pack stock solutions as described in Table 2.1. Follow the regulations of the Occupational Safety
and Health Administration (OSHA) when handling chemicals.
Site kits of consumable supplies for each sampling site will be delivered based on the supply
requests each crew submits prior to and during the sampling season. Crews will submit an ^
electronic request form to order site kits, forms, 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 ^
LU
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 £j
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. 5
Store at least one extra site kit in the vehicle in the event replacement items are needed h
immediately. O
i—
u
3
Q
o
QC
11
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 2.1 Stock Solutions, Uses, and Methods for Preparation
Solution
Use
Preparation
Bleach (1%)
Clean nets, other gear, and boat.
Add 40 mL bleach to 4L distilled water.
Bleach (10%)
Clean periphyton sampling
equipment.
Add 40 mL bleach to 400 mL distilled water.
10% Buffered
Formalint
Preservation of periphyton ID sample
and fixing Fish Vouchers
Formaldehyde (37-40%) 100 ml
Distilled water 900 ml
NaH2P04 4.0 g
Na2HP04 (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 is to initial all paper data forms after review or if
submitting data through the app, Field Crew Leader must review the PDFs received after 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, completeness, and legibility. 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 must initial paper field forms to indicate that they
are ready to be sent to the Information Management Center (a similar review process is used for
electronic forms). Each sample label must also be checked for accuracy, completeness, and
legibility. The Field Crew Leader must cross-check the sample numbers on the labels with those
w recorded on the data forms,
z
I
^ 2.2.2.2 Inspect and Prepare Samples
<
1/1 All samples need to be inspected and appropriately preserved and packaged for transport.
^ Check that all samples are labeled, and all labels are filled in completely. Check that each label is
£i covered with clear plastic tape. Check the integrity of each sample container, and be sure there
< are no leaks. Make sure that all sample containers are properly sealed. Make sure that all
^ sample containers are properly preserved for storage or immediate shipment.
I—
g 2.2.2.3 Equipment Cleanup and Check
u All equipment and gear must be cleaned and disinfected between sites to reduce the risk of
q transferring nuisance species and pathogens. Species of primary concern in the U.S. include
§ Eurasian watermilfoil (Myriophyllum spicatum), zebra mussels (Dreissena polymorpha), New
z Zealand mud snails (Potamopyrgus antipodarum), Myxobolus cerebralis (a sporozoan parasite
12
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
that causes salmonid whirling disease), and Batrachochytrium dendrobatidis (a chytrid fungus
that threatens amphibian populations). Field Crews must be aware of regional species of
concern, and take appropriate precautions to avoid transfer of these species. There are several
online resources regarding invasive species, including information on cleaning and disinfecting
gear, such as the Whirling Disease Foundation (www.whirling-disease.org). the USDA Forest
Service (Preventing Accidental Introductions of Freshwater Invasive Species, available from
http://www.fs.fed.us/invasivespecies/documents/Aquatic is prevention.pdf), and the
California Dept. of Fish and Game (Hosea and Finlayson 2005). General information about
freshwater invasive species is available from the U.S. Geological Survey Nonindigenous Aquatic
Species website (http://nas.er.usgs.gov), the Protect Your Waters website that is co-sponsored
by the U.S. Fish and Wildlife Service (http://www.protectyourwaters.net/hitchhikers). and the
Sea Grant Program (http://www.sgnis.org).
Handle and dispose of disinfectant solutions properly, and take care to avoid damage to lawns
or other property. Table 2.2 describes equipment care. Inspect all equipment, including nets,
boat trailer, and waders, and clean off any plant and animal material. Inspect, clean, and
handpick plant and animal remains from vehicle, boat, motor, trailer and waders. Before moving
to the next site, if a commercial car wash facility is available, wash the vehicle, boat, and trailer
and rinse thoroughly (hot water pressurized rinse with no soap). Rinse equipment and boat with
1% -10% bleach solution or other specialized disinfectant to prevent the spread of exotics. Note
that many organizations now recommend against using felt-soled wading boots in affected
areas due to the difficulty in removing myxospores and mudsnails.
2.2.2.4 Supply Inventory
Once a field day is completed, crews should inventory and restock supplies as needed. Ensure
that there is a sufficient quantity of site kits to allow sampling at upcoming sites (for at least the
next 1-2 weeks). Take note of any supplies that are nearing depletion. Also note any items that
may have been lost or damaged during the sampling event. Request additional site kits and/or
supplies as needed via the electronic request form. Requests must be made two weeks before
needed. Note that not all supplies can be replenished by EPA through the Logistics Contractor,
so crews will need to supply some items themselves.
Table 2.2 Post-sampling Equipment Care
Equipment Care after Sampling
z
i
CL
<
l/l
LU
I
CO
<
LU
Q
<
5
o
i—
z
o
I—
u
3
Q
o
QC
1. Clean for biological contaminants.
Prior to departing site, drain all water from live wells and buckets used to hold and process fish.
Inspect sampling gear, waders, boots, etc. for evidence of mud, snails, plant fragments, algae, animal
remains, or debris, and remove using brushes or other tools.
At the base location, inspect and rinse seines, dip nets, kick nets, waders, and boots with water and dry.
Use one of the procedures below to disinfect gear if necessary.
Additional precautions to prevent transfer of Whirling Disease spores, New Zealand mudsnails, and
amphibian chytrid fungus is provided below:
Before visiting the site, consult the site dossier and determine if it is in an area where whirling disease,
New Zealand mud snails, or chytrid fungus are known to exist. Contact the local State fishery biologist to
confirm the existence or absence of these organisms.
13
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Equipment Care after Sampling
If the stream is listed as "positive" for any of the organisms, or no information is available, avoid using
felt-soled wading boots, and, after sampling, disinfect all fish and benthos sampling gear and other
equipment that came into contact with water or sediments (i.e., waders, boots, etc.) by one of the
following procedures:
Option A:
1. Soak gear in a 10% household bleach solution for at least 10 minutes, or wipe or spray on a 50%
household bleach solution and let stand for 5 minutes
2. Rinse with clean water (do not use stream water), and remove remaining debris
3. Place gear in a freezer overnight or soak in a 50% solution of Formula 409® antibacterial cleaner
for at least 10 minutes or soak gear in 120°F (49°C) water for at least 1 minute.
4. Dry gear in direct sunlight (at least 84 °F) for at least 4 hours.
Option B:
1. Soak gear in a solution of Sparquat® (4-6 oz. per gallon of water) for at least 10 minutes
(Sparquat is especially effective at inactivating whirling disease spores).
2. Place gear in a freezer overnight or soak in 120°F (49°C) water for at least 1 minute.
3. Dry gear in direct sunlight (at least 84 °F) for at least 4 hours.
2. Clean and dry other equipment prior to storage.
Rinse coolers with water to clean off any dirt or debris on the outside and inside.
Rinse periphyton sampling equipment with tap water at the base location.
Make sure conductivity meter probes are rinsed with deionized water and stored moist.
Rinse all containers used to collect water chemistry samples three times with deionized water. Place
beakers in a 1-gallon sealable plastic bag with a cube container for use at the next stream.
Check nets for holes and repair or locate replacements.
3. Inventory equipment and supply needs and relay orders to the IM Team via the supply request form.
4. Remove GPS, multi-probe meter, and electrofishing unit from carrying cases and set up for
predeparture checks and calibration. Examine the oxygen membranes for cracks, wrinkles, or
bubbles. Replace if necessary, allowing sufficient time for equilibration.
5. Recharge/replace batteries as necessary.
6. Replenish fuel and oil; if a commercial car wash facility is available, thoroughly clean vehicle and boat
(hot water pressurized rinse and no soap).
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.
CO
<
q 2.3.1 General Considerations
<
5 Important considerations related to field safety are presented in Table 2.3. Please follow your
h own agency's health and safety protocols, or refer to the Health and Safety Guidance for Field
q Sampling: National Rivers and Streams Assessment (available from the EPA Regional
h Coordinator) and Logistics of Ecological Sampling on Large Rivers (Tlotemersch, et al. (editors)
g 2000). Additional sources of information regarding safety-related training include the American
O Red Cross (1979), the National Institute for Occupational Safety and Health (1981), U.S. Coast
t Guard (1987) and Ohio EPA (1990).
14
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Field Crew members should become familiar with the hazards involved with sampling
equipment and establish appropriate safety practices prior to using them. They must make sure
all equipment is in safe working condition. Personnel must consider and prepare for hazards
associated with the operation of motor vehicles, boats, winches, tools, and other incidental
equipment. Boat operators should meet any state requirements for boat operation and be
familiar with U.S. Coast Guard rules and regulations for safe boating contained in a pamphlet,
"Federal Requirements for Recreational Boats," available from a local U.S. Coast Guard Director
or Auxiliary or State Boating Official (U.S. Coast Guard, 1987). A personal floatation device (PFD)
must be worn by crew members at all times on the water. All boats with motors must have fire
extinguishers, boat horns, PFDs or flotation cushions, and flares or communication devices.
Boats should stay in visual contact with each other, and should use 2-way radios to
communicate.
Primary responsibility for safety while electrofishing rests with the Field Crew Leader.
Electrofishing units may deliver a fatal electrical shock, and should only be used by qualified,
experienced operators. Field Crew members using electrofishing equipment must be insulated
from the water, boat, and electrodes via rubber boots and linesman gloves. All personnel should
use chest waders with nonslip soles and linesman gloves. DO NOT wear breathable waders
while electrofishing. If waders become wet inside, stop fishing until they are thoroughly dry or
use a dry pair. Avoid contact with the anode and cathode at all times due to the potential shock
hazard. If you perspire heavily, wear polypropylene or some other wicking and insulating
clothing instead of cotton. If it is necessary for a crew member to reach into the water to pick up
a fish or something that has been dropped, do so only after the electrical current is off and the
anode is removed from the water. Do not resume electrofishing until all individuals are clear of
the electroshock hazard. Ensure that the backpack electrofishing equipment is equipped with a
45° tilt switch that interrupts the current. Do not make any modifications to the electrofishing
unit that would hinder 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.
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
<
i/i
CO
<
LU
Q
<
U
3
Q
o
QC
15
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Personal Safety
• Field clothing and other protective gear including lifejackets for all crew members
• Medical and personal information (allergies, personal health conditions)
• Personal contacts (family, telephone numbers, etc.)
• Physical exams and immunizations
A communications plan to address safety and emergency situations is essential. All field
personnel need to be fully aware of all lines of communication. Field personnel should have a
daily check-in procedure for safety. An emergency communications plan should include contacts
for police, ambulance, fire departments, hospitals, and search and rescue personnel.
Proper field clothing should be worn to prevent hypothermia, heat exhaustion, sunstroke,
drowning, or other dangers. Field personnel must be able to swim, and personal flotation
devices must be used. Chest waders made of rubberized or neoprene material must always be
worn with a belt to prevent them from filling with water in case of a fall. A PFD and suitable
footwear must be worn at all times while on board a boat.
Many hazards lie out of sight in the bottoms of rivers and streams. Broken glass or sharp pieces
of metal embedded in the substrate can cause serious injury if care is not exercised when
walking or working with the hands in such environments. Infectious agents and toxic substances
that can be absorbed through the skin or inhaled may also be present in the water or sediment.
Personnel who may be exposed to water known or suspected to contain human or animal
wastes that carry causative agents or pathogens must be immunized against tetanus, hepatitis,
typhoid fever, and polio. Biological wastes can also be a threat in the form of viruses, bacteria,
rickettsia, fungi, or parasites.
2.3.2 Safety Equipment
Appropriate safety apparel such as waders, linesman gloves, safety glasses, etc. must be
available and used when necessary. First aid kits, fire extinguishers, and blankets must be readily
available in the field. Cellular or satellite telephones and/or portable radios should be provided
to Field Crews working in remote areas in case of an emergency. Supplies (e.g., clean water,
antibacterial soap, ethyl alcohol) must be available for cleaning exposed body parts that may
have been contaminated by pollutants in the water.
2.3.3 Safety Guidelines for Field Operations
General safety guidelines for field operations are presented in Table 2.4. Personnel participating
in field activities should be in sound physical condition and have a physical examination annually
or in accordance with organizational requirements. All surface waters and sediments should be
considered potential health hazards due to potential toxic substances or pathogens. Persons
must become familiar with the health hazards associated with using chemical fixing and/or
preserving agents. Chemical wastes can be hazardous due to flammability, explosiveness,
toxicity, causticity, or chemical reactivity. All chemical wastes must be discarded according to
standardized health and hazards procedures (e.g., National Institute for Occupational Safety and
Health [1981]; 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
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
that the proper actions be taken and that field personnel do not expose themselves to
something harmful. The following guidelines should be applied:
1. First and foremost, protect the health and safety of all personnel. Take necessary steps
to avoid injury or exposure to hazardous materials. If you have been trained to take
action such as cleaning up a minor fuel spill during fueling of a boat, do it. However, you
should always err on the side of personal safety.
2. Field personnel should never disturb or retrieve improperly disposed hazardous
materials from the field to bring back to a facility for "disposal". To do so may worsen
the impact, incur personal liability for the crew members and/or their respective
organizations, cause personal injury, or cause unbudgeted expenditure of time and
money for proper treatment and disposal of the material. Notify the appropriate
authorities so they may properly respond to the incident.
3. For most environmental incidents, the following emergency telephone numbers should
be provided to all Field Crews: State or Tribal department of environmental quality or
protection, U.S. Coast Guard, and the U.S. EPA regional office. In the event of a major
environmental incident, the National Response Center may need to be notified at 1-800-
424-8802.
Table 2.4 General Safety Guidelines for Field Operations
Two crew members must be present during all sample collection activities, and no one should be
left alone while in the field. Boats should proceed together down the river.
Use caution when sampling on foot in swift or deep water. Wear a suitable PFD and consider
using a safety tether held by an assistant.
Use extreme care walking on riprap. Rocks can shift unexpectedly and serious falls are possible.
Field Crew members using electrofishing equipment must be insulated from the water, boat, and
electrodes via non-breathable waders and linesman gloves. Use chest waders with nonslip soles.
Electrofishing units may deliver a fatal electrical shock, and should only be used by qualified,
experienced operators.
Professional quality breathable waders with a belt are recommended for littoral sampling only,
and at a safe distance from the electrofishing sampling. Neoprene boots are an alternative, but
should have sturdy, puncture resistant soles.
Exposure to water and sediments should be minimized as much as possible. Use gloves if
necessary, and clean exposed body parts as soon as possible after contact.
All electrical equipment must bear the approval seal of Underwriters Laboratories and must be
properly grounded to protect against electric shock.
Use heavy gloves when hands are used to agitate the substrate during collection of benthic
macroinvertebrate samples.
Use appropriate protective equipment (e.g., gloves, safety glasses) when handling and using
hazardous chemicals.
Crews working in areas with venomous snakes must check with the local Drug and Poison Control
Center for recommendations on what should be done in case of a bite from a poisonous snake.
Any person allergic to bee stings, other insect bites, or plants (i.e., poison ivy, oak, sumac, etc.)
must take proper precautions and have any needed medications handy.
Field personnel should also protect themselves against deer or wood ticks because of the
potential risk of acquiring pathogens that cause Rocky Mountain spotted fever, Lyme disease,
and other illnesses.
Field personnel should be familiar with the symptoms of hypothermia and know what to do in
case symptoms occur. Hypothermia can kill a person at temperatures much above freezing (up to
10°C or 50°F) if he or she is exposed to wind or becomes wet.
<
i/i
CO
<
LU
Q
<
U
3
Q
o
QC
17
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
¦ Field personnel should be familiar with the symptoms of heat/sun stroke and be prepared to
move a suffering individual into cooler surroundings and hydrate immediately.
¦ Handle and dispose of chemical wastes properly. Do not dispose any chemicals in the field.
2.4 Forms (Paper or Electronic)
Forms are the key to data collection and tracking for the NRSA 2018/19. Electronic forms have
been developed as well as paper forms. These electronic forms should streamline data
collection. Field Crews are encouraged to use electronic forms whenever possible, but will have
the option of using paper forms when necessary.
2.4.1 Field Forms
Field forms are the primary documents where crews record measures, observations, and
collection information during the course of the field day. Additional information regarding
specifics of data entry is contained in Section 1.6.
• Electronic Field Forms: This form of data collection can be collected through an Apple
iOS portable electronic device (tablets). This method of data collection will require a
Field Crew to download or install the developed Application (or "App") onto the device.
The field forms will be optimized for tablet devices. Once downloaded and the App
launched, the field forms will be split into sections or "form-lets" for easier data entry. It
is important for a Field Crew to familiarize themselves with the App prior to field
sampling.
• Paper Field Forms: A paper field form packet (wadeable or non-wadeable) for each site
will be provided by the NARS Information Management (IM) Coordinator if crews have
elected to use paper field forms for data collection. Crews will need to add these forms
to the site packet prior to going in the field. After a site is sampled, the completed NRSA
2018/19 paper field forms are checked for completeness and organized sequentially
into a Data Packet. The data packets from several sites are batched together and sent
every 1-2 weeks to the NARS IM Coordinator and are accompanied by a Tracking form to
track which data packets have been shipped. Extra paper field forms will always be
provided to Field Crews to serve as backup copies in case of lost forms or problems with
electronic field forms.
2.4.2 Tracking Forms
Tracking forms describe the status and location of all samples collected during NRSA 2018/19.
Field Crew Leaders will transmit these forms electronically (through App submissions, by
emailing scans of paper forms or by emailing a fillable PDF form) to the NARS IM Coordinator at
specified times and crews will pack hard copies of the tracking forms in shipping containers with
the samples. See APPENDIX C: SHIPPING GUIDELINES for more information.
• Site and Sample Status/Water Chemistry Lab Tracking: Transmitted within 24 hours of
sampling or visiting a site to report on the status of the site (e.g. sampleable or not), to
record the Sample ID numbers, and to indicate the status of all samples collected at the
site (including immediate shipments and batch shipments).
• Tracking - Packets: Accompany paper data packets that are batched together from
multiple sites and shipped every 1 or 2 weeks. These are sent to the NARS IM
Coordinator.
18
<
i/i
CO
<
LU
Q
<
U
3
Q
o
QC
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
• Tracking -Samples: Submitted at the time of shipping with samples that are batched
together from multiple sites and shipped every 1 or 2 weeks. Whenever batched
samples are shipped to their designated lab for analysis/storage, the appropriate
tracking form, which lists the Sample ID numbers for all samples packed in a shipping
container, is transmitted electronically to the NARS IM Coordinator. Separate forms
exist for the tracking of frozen batched samples, non-chilled batched samples and whole
fish samples.
• Packing Slips: 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.
2.4.3 Equipment and Supplies
2.4.3.1 Request Form
Field Crews will submit requests for field forms, labels and site kits via an electronic request
form (Figure 2.3). This form will be submitted to the NARS IM Coordinator who will ensure that
the request reaches the appropriate entity. Crews should submit the Request Form at least two
weeks prior to their desired sampling date. In addition to typing in specific requests, crews may
select one or more of the pre-populated items listed below:
• Site kit: contains all consumable supplies for one site, sample labels, packing slips, FedEx
shipping label to WRS, cooler (for sending immediate samples to WRS), and cooler liner.
• Whole fish tissue kit (for crews collecting whole fish tissue samples): contains all
consumable supplies for each of the 477 river sites designated for whole fish tissue
sample collection, additional information for whole fish tissue sampling, cooler, Class 9
hazardous label, and FedEx shipping label to the designated laboratory for interim
storage. All whole fish tissue forms and labels will be provided in the paper form
packets and tracking packets.
• Frozen batched cooler: contains cooler, dry ice liner and pad, Class 9 hazardous label,
FedEx label to GLEC.
• Non-chilled batched cooler: contains cooler, cooler liner, FedEx label to GLEC
• Tracking Packets: Crews using the NARS App, may request tracking packets as backups
or replacements to the tracking packets included in the site kit. A tracking packet
contains sample labels and packing slips for one site.
• Paper Form Packets: Crews not using the NARS App will need to request paper form
packets for their use at a site. Paper form packets will contain the appropriate number
of forms for one site and will also contain tracking forms that will need to be completed
by hand and scanned at the time of shipping samples.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
u
Z>:
Q
o
cc
Request Date:
Requester:
State:
Phone:
Email:
/ 2 0 18
i i i i i
NRSA 2018/19 Request Form
Ship to:
Name:
Company:
Address:
j i City:
Zip Code:
Supplies Needed (Mark all that apply):
O Full Site Kit (all bottles and consumables for 1 site, including packing slip, adhesive labels, FedEx shipping label to WRS,
cooler and cooler liner)
How many? Need by: / / 2 0 1 8 Comments:
O Frozen batched cooler (includes cooler, dry ice liner, Class 9 hazardous label, FedEx label to GLEC)
How many? Need by: / / 2 0 1 8 Comments:
O Non-chilled batched cooler (includes cooler, cooler liner, FedEx label to GLEC)
How many? Need by: / / 2 0 1 8 Comments:
O Whole fish tissue kit (materials needed for 1 site whole fish tissue site, including FedEx shipping label to Microbac and cooler)
How many? Need by: / / 2 0 1 8 Comments:
O Full Packet (paper field and tracking forms + adhesive labels (for non-app users only))
How many? Need by: / / 2 0 1 8 Comments:
O Tracking Packet (packing slips + adhesive labels (normally provided in site kits, extras needed)
How many? Need by: / / 2 0 1 8 Comments:
O Foil squares (aluminum) - pack of 25
How many? Need by: /
/ 2 0 1 8 Comments:
O HDPE bottle (1 L, white, wide-mouth)
How many? Need by: /
/ 2 0 1 8 Comments:
O Sodium Thiosulfate Tablets - vial of 25
How many? Need by: /
/ 2 0 1 8 Comments:
O Tape strips - packs of 25
How many?
Need by:
/
/
/ 2 0 1 8 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:
/ 2 0 18
/ 2 0 1
/ 2 0 1 8
/ 2 0 1 8
03/27/2018 NRSA18 Request Form
0835186180
Figure 2.3 Electronic Request Form
20
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
2.4.3.2 Base Kit
The Base Kit is comprised of the subset of durable equipment and supplies needed for NRSA
2018/19 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.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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-17-002). Inspect the selected reach for appropriate access,
safety, and general conditions. Decide whether the site is at base flow condition and not unduly
influenced by rain events which could affect the representativeness of field data and samples. If
you determine that the site can be sampled, lay out a defined reach within which all sampling
and measurement activities are conducted.
3.1 Site Verification Activities
3.1.1 Locating the X-Site
River and stream sampling points were chosen using the medium resolution National
Hydrography Dataset (NHD), in particular NHD-Plus V2, 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. If the provided coordinates are located on dry land near a stream,
move the coordinates to the nearest blue line in NHD-Plus during the desktop reconnaissance.
Note this movement on the site recon tracking form and in the comments section of the Site
Verification Form.
Table 3.1 Equipment and Supplies: for Site Verification
i/i
LU
CC
Z>
Q
LU
U
o
cc
o_
LU
H
LD
I
<
H
For locating and
verifying site
Sampling permit and landowner access (if required)
Field Operations Manual and laminated Quick Reference Guide
Site dossier, including access information, site spreadsheet with map coordinates,
street and/or topographic maps with "X-site" marked
NRSA 2018/19 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
Clipboard
#2 pencils
Verification Form
Fine-tipped indelible markers to write on flagging
22
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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.
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, 150m 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. 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 Site Verification
Form (Figure 3.1). If the site is non-sampleable or inaccessible, no further sampling activities are
conducted. Replace the site with the first available oversample site on the state list within the
same category based on Strahler category, new/resample status, and ecoregion (Section 1.3).
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Site ID:
NRSA 2018/19 VERIFICATION (Front)
Visit: 01 O 2 Date: /
Reviewed by
State of Site Location:
O This is a special State site
STREAM/RIVER VERIFICATION INFORMATION
Stream/River verified by (Mark all that apply): O GPS
O Other (Describe Here):
O Loc a I Co ntact OSigns
O Roads O Topo. Map
Coordinates
Latitude
Longitude
# of Satellites Elevation at transect A
GPS
Decimal Degrees
NAD 83
Os3 0>A
Location: o X-Slte (wadeable) O Transect A (non-wadeable)
Oft Om
DID YOU SAMPLE THIS SITE?
O YES If Yes, check one below
O NO If No, check one below
SAMPLEABLE (Choose method used)
O Wadeable - Continuous water, greater than 50% wadeable
O Beatable
O Partial - Sampled by wading {>50% of reach sampled). Explain below.
O Partial - Sampled by boat (>50% of reach sampled). Explain below.
O Wadeable Interrupted - Not continuous water along reach
O Boatable Interrupted - Not continuous water along reach
O Altered - StreamyRiver Channel Present but differs from Map
ADDITIONAL SITE CHARACTERISTICS
O Tidally Influenced O Blackwater O Not Applicable
NON-SAMPLEABLE-PERMANENT
O Dry - Visited
O Dry - Not visited
O Wetland (No Definable Channel)
O Map Error (No evidence channel/waterbody ever present)
Q Impounded (> 7 day residence time)
0 Tidal (exceeds salinity threshold)
O Other (explain In comments)
NON-SAMPLEABLE-TEMPORARY
O Not boatable - Need a different crew - Reschedule regardless of year
O Not wadeable - Need a different crew - Reschedule regardless of year
O Other (explain In comments)
NO ACCESS
O Access Permission Denied
O Permanently Inaccessible (Unable/Unsafe to Reach Site)
O Temporarily inaccessible-Fire, etc. - Reschedule regardless of year
O Other (explain In comments)
GENERAL COMMENTS
DIRECTIONS TO SITE
7734319701
02/28/2018 NRSA18 Verification
U
O
cc
Q_
Figure 3.1 Verification Form (front)
CO
i
<
24
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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.
Boatable: Continuous water, too deep to sample by wading.
Partial wadeable: Sampled by wading (>50% of reach sampled).
Partial boatable: Sampled by boat (>50% of reach sampled).
Wadeable Interrupted: not continuous water along reach, >50% water in reach.
Boatable Interrupted: not continuous water along reach, >50% water in reach.
Altered Channel: Stream/river channel present but differs from map.
Non-Sampleable Categories
PERMANENT
Dry Channel: Less than 50% water within the reach. Record as "Dry-Visited." If site was determined to be
dry (or otherwise non-perennial) from another source and/or field verified before the actual sampling
visit, record as "Dry-Not visited" 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.
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.
i/i
LU
QC
3
Q
LU
U
o
QC
CL
LU
H
IS)
i
<
I—
25
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
No Access to Site Categories
Access Permission Denied: You are denied access to the site by the landowner.
Permanently Inaccessible: Site is unlikely to be sampled by anyone due to physical barriers that prevent
access to the site (e.g., cliffs).
Temporarily Inaccessible: Site cannot be reached due to barriers that may not be present at a future date
(e.g. forest fire, high water, road temporarily closed, unsafe weather conditions).
6. Do not sample non-target or "Non-sampleable" or "No Access" sites. Fill in the "NO" bubble for
"Did you sample this site?" and mark the appropriate bubble in the "Non-Sampleable" or "No
Access" section of the Verification Form; provide a detailed explanation in comments section.
3.1.3 Elevation at Transect A
Record 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 Site Verification Form. 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.
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
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
camera at a site, date stamp the photograph and include the site ID. Most cell phone cameras
also have the ability to attach geographical location data to a particular picture. Alternatively,
start the sequence with one photograph of an 8.5 x 11 inch piece of paper with the site ID,
waterbody name, and date printed in large, thick letters. After the photo of the site ID
information, take at least two photographs at the X-site, one in the upstream direction and one
downstream. Take any additional photos you find interesting after these first three pictures.
Keep a log of your photographs and briefly describe each one. Photographs can be uploaded to
the NARS SharePoint site.
3.2 Laying out the sampling reach
Many of the biological and habitat structure measures require sampling a certain length of a
stream to get a representative picture of the ecological community. A length of 40 times the
average wetted width is necessary to characterize the habitat and several biotic assemblages
associated with the sampling reach. Establish the sampling reach about the X-site using the
procedures described in Table 3.4 (wadeable sites).
When you arrive at the site, scout the sampling reach to make sure it is clear of obstacles that
would prohibit sampling and data collection activities. Record the channel width used to
determine the reach length, and the sampling reach length upstream and downstream of the X-
site on the Verification Form (back) as shown in Figure 3.2.
Figure 3.3 illustrates the principal features of the established sampling reach for wadeable sites,
including the location of 11 cross-section transects used for collecting samples and physical
habitat measurements. The figures also show the specific sampling stations on each transect for
collection of periphyton, and benthic macroinvertebrate samples.
Before leaving the site, complete a rough sketch map of the reach you sampled on page 2 of the
Verification Form (Figure 3.2). In addition to any other interesting features that should be
marked on the map, note any landmarks/directions that can be used to find the X-site for future
visits.
Table 3.4 Procedure: Laying Out the Sampling Reach at Wadeable Sites
1. Locate the X-site using the coordinates provided for the site.
2. Use a surveyor's rod, tape measure, or laser range finder to determine the wetted width of the
channel at five places of "typical" width within approximately five channel widths upstream and
downstream from the X-site. Average the five readings together and round to the nearest 1 m.
3. Multiply the average wetted width by 40 to determine the reach length If the average width is <4
m, use 150 m as a minimum reach length. If the average width is >100 m, use 4 km as a maximum
reach length. Record both the average channel width and total stream length on page 2 of the
Verification Form.
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.
7. Starting at the X-site (or the new midpoint of the reach if it had to be adjusted as described in Step
i/i
LU
CC
Z>
Q
LU
U
o
en
o_
LU
H
LD
I
<
H
27
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
6), measure a distance of one-half the reach length down one side of the stream using a GPS unit,
laser rangefinder, or tape measure. Be careful not to "cut corners". Enter the channel to make
measurements only when necessary to avoid disturbing the stream channel prior to sampling
activities. This endpoint is the downstream end of the reach, and is flagged as Transect "A".
8. At Transect A, use the seconds display on a digital watch to select the initial sampling station for
standard transect samples: l-3="Left", 4-6="Center", 7-9=Right. Mark "L", "C", or "R" on the
transect flagging; the three potential collection points are roughly equivalent to 25%, 50%, and 75%
of the channel width, respectively. Note that left and right sides of the stream are determined
while you are facing downstream. It is at these locations that you will collect benthic
macroinvertebrate and periphyton samples.
9. Measure 1/10 of the required reach length upstream from transect A. Flag this spot as transect B.
Assign the sampling station systematically after the first random selection following the repeating
pattern L, C, R as you move upstream (Figure 3.3).
10. Proceed upstream with the tape measure and flag the positions of nine additional transects
(labeled "C" through "K" as you move upstream) at intervals equal to 1/10 of the reach length.
Continue to assign the sampling stations systematically.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Channel Width
Used to Define
Reach (m)
DISTANCE (m) FROM X-SITE
Total Reach
Length Intended (m):
Comment:
Upstream Length:
Downstream Length:
1 1 1 1 1
¦ ¦ ¦ ¦ ¦ *
*
V
Site ID:
NRSA 2018/19 VERIFICATION (Back)
Visit: o 1 O 2 Date: /
R*V!«wed by
STREAM/RIVER REACH DETERMINATION
PERSONNEL
Crew Leader:
Fish Taxonomist:
Name:
Name:
Name:
Name:
Name:
Name:
02/28/2018 NR5A18 Verification
2492319707 ^
Figure 3.2 Verification Form (back)
DC
z>
~
LU
u
O
DC
DL
(Ti
I
<
29
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Sampling Points
• L = left; C = center; R = right
• First point (at transect A)
determined randomly
• Subsequent point assigned in
order L, C, R
Upstream endpoint is "Transect K"
Downstream endpoint is "Transect A'
Distance between transects
= 1/10,h of total reach length
X Site
(j>C
| FLOlV^>
Total reach length = 40 x mean wetted width (min = 150m; max = 4km)
Figure 3.3 Sampling Reach Features for a Wadeable Site
3.2.1 Sliding the Reach
There are some conditions that may require sliding the reach about the X-site (i.e., the X-site is
no longer located at the midpoint of the reach) to avoid features we do not wish to or physically
cannot sample across. Reasons for sliding the reach include:
• Lack of landowner permission.
• Confluence with higher order waterbody.
• Impoundment.
• Impassable barrier.
Sliding the reach involves noting the distance of the barrier, confluence, or other restriction
from the X-site, and flagging the restriction as the endpoint of the reach. Add the distance to the
other end of the reach, such that the total reach length remains the same, but it is no longer
centered about the X-site. Table 3.5 describes when you should and should not slide the
sampling reach.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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 4) will be collected at the X-site (even if the
reach has been adjusted by "sliding" it). If the X-site is dry and there is water elsewhere in the
sample reach, collect the sample from a location having water with a surface area >1 m2 and a
depth >10 cm.
Collect data for the physical habitat indicator along the entire sample reach from interrupted
streams, regardless of the amount of water present at each of the transects. Obtain depth
measurements along the deepest part of the channel (the "thalweg") along the entire sampling
reach to provide a record of the "water" status of the stream for future comparisons (e.g., the
percent of length with intermittent pools or no water). Other measurements associated with
characterizing riparian condition, substrate type, etc., are useful to help infer conditions in the
stream when water is flowing.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 3.6 Reach Layout Modifications for Interrupted Streams
Physical Habitat, Periphyton and Benthic Macroinvertebrates
Streams with less than 50% of the reach length containing water (not necessarily continuous) are
considered dry and are not sampled.
If more than 50% of the channel has water and if the X-site is dry but there is flowing water or a pool of
water having a surface area > 1 m2 and a depth > 10 cm somewhere along the defined sampling reach, take
the water sample at the pool or flowing water location that is nearest to the X-site. Note that the sample
wasn't collected at the X-site and where on the reach the sample was collected on the field data form.
Do not collect a water sample if there is no acceptable location within the sampling reach. Record a "K"
flag for the water chemistry sample on the sample collection form and explain why the sample was not
collected in the comments section of the form.
Obtain a complete thalweg profile for the entire reach. At points where the channel is dry, record depth as
0 cm and wetted width as 0 m.
At each of the transects (cross-sections), sample the stream depending on flow status:
DRY CHANNEL: No surface water anywhere in cross-section; collect all physical habitat data. Use the
unvegetated area of the channel to determine the channel width and the subsequent location of substrate
sampling points. Record the wetted width as 0 m. Record substrate data at the sampling points located in
the unvegetated, but dry, channel. Do not collect periphyton or benthic macroinvertebrates from this
transect.
DAMP CHANNEL: No flowing water at transect, only puddles of water < 10 cm deep; collect all physical
habitat data. Do not collect periphyton or benthic macroinvertebrates from this transect.
WATER PRESENT: Transect has flow or pools > 10 cm deep; collect all data and measurements for physical
habitat, periphyton, benthic macroinvertebrate, and fish indicators, using standard procedures.
3.3.2 Braided Rivers and Streams
oe
3
Q
LU
u
o
QC
CL
l/l
I
<
Depending upon the geographic area and/or the time of the sampling visit, you may encounter a
stream having "braided" channels, which are characterized by numerous sub-channels that are
generally small and short, often with no obvious dominant channel. If you encounter a braided
stream, establish the sampling reach using the procedures presented in Table 3.7. Figuring the
mean width of extensively braided rivers and streams for purposes of setting up the sampling
reach length is challenging. For braided channels, measure the mean width and bankfull width
as defined in the physical habitat protocols (Section 8). For relatively small streams (mean
bankfull width <15 m) the sampling reach is defined as 40 times the mean bankfull width. For
larger streams (>15 m), sum the actual wetted width of all the braids and use that as the width
for calculating the 40 channel width reach length. If there is any question regarding an
appropriate reach length for the braided system, it is better to overestimate. Make detailed
notes and sketches on the Verification Form (Figure 3.1 and Figure 3.2) about what you did. If
using the App, sketches on paper can be scanned via CamScanner (software preloaded on iPads)
and submitted to EPA SharePoint. It is important to remember that the purpose of the 40
channel width reach length is to sample enough streams to incorporate the variability in habitat
types. Generally, the objective is to sample a long enough stretch of a stream to include 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
32
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
braids), a 400 m long sample reach length is likely to be sufficient, especially if the system has
fairly homogenous habitat throughout its length.
Table 3.7 Procedure: Modifications for Braided Rivers and Streams
1. Estimate the mean width as the bankfull channel width as defined in the physical habitat protocol.
• If the mean width is <15 m, set up a 40 x channel width sample reach in the normal manner,
using the mean bankfull width for your calculations.
• If >15 m, sum up the actual wetted width of all the braids and use that as the width for
calculating the 40 x channel width reach length. Remember the minimum reach length is
always 150 m.
• If the reach length seems too short for the system in question, set up a longer sample reach,
taking into consideration that the objective is to sample a long enough stretch of a stream to
include at least two to three meander cycles (about six pool riffle habitat sequences).
2. Make detailed notes and sketches on the Verification Form about what you did.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
u
u
4 WATER CHEMISTRY / CHLOROPHYLL-a SAMPLE
COLLECTION AND PRESERVATION
4.1 In Situ Measurements of Dissolved Oxygen, pH, Temperature,
and Conductivity
4.1.1 Summary of Method
Measure in situ DO, pH, water temperature, and conductivity using a calibrated multi-parameter
water quality meter (or sonde). Take the measurements mid-channel at the X-site. Take the
readings at 0.5 m depth. Measure the site depth accurately before taking the measurements. If
the depth at the x-site is less than 1 meter, take the measurements at mid-depth. Take care to
avoid the probe contacting bottom sediments, as the instruments are delicate. Record the
measurements on the Field Measurement Form, as seen in Figure 4.1.
4.1.2 Equipment and Supplies
Table 4.1 provides the equipment and supplies needed to measure dissolved oxygen, pH,
temperature, and conductivity.
Table 4.1 Equipment and Supplies: DO, pH, Temperature, and Conductivity
For taking measurements and Multi-parameter water quality meter with pH, DO, temperature,
calibrating the water quality meter and conductivity probes.
O Extra batteries
< De-ionized (Dl) and tap water
2^ Calibration cups and standards
i/i
£ Barometer or elevation chart to use for calibration
CL
^ For recording measurements Field Measurement Form
^ Pencils (for data forms)
O
^ 4.1,2.1 Multi-Probe Sonde
I
I
O Dissolved Oxygen Meter
^ Calibrate the DO meter prior to each sampling event. It is recommended that the probe be
^ calibrated in the field against an atmospheric standard (e.g., ambient air saturated with water).
^ Note that DO should always be calibrated at the site and should never be calibrated at your base
CD
_j location. Follow your manufacturer's guidelines for calibration of the DO probe.
>
J pH Meter
O
g 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.
I—
to
^ Temperature Meter
Check the accuracy of the sensor against a thermometer that is traceable to the National
oc 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
5 values can be recorded each time for the Field Measurement Form unless the accuracy check is
34
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
conducted again. The entire temperature range encountered in the NRSA should be
incorporated in the testing procedure and a record of test results kept on file.
Conductivity Meter
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.
4.1.3 Sampling Procedure
Table 4.2 presents step by step procedures for measuring dissolved oxygen, pH, temperature,
and conductivity.
Table 4.2 Procedure: Temperature, pH, Conductivity and Dissolved Oxygen
1. Check Meter and probes and calibrate according to manufacturer's specifications.
2. Samples are taken mid-channel, at the X site, at a depth of 0.5 meters or at mid-depth if less than 1
meter deep.
3. Lower the sonde in the water and measure DO, pH, temperature, and conductivity at 0.5 m depth
(or at mid-depth if less than 1 meter deep).
4. Record the measurements on the Field Measurement Form, noting whether the conductivity value
is corrected to 25°C.
5. Flag any measurements that the crew feels needs further comment or when a measurement
cannot be made.
<
>
QC
oe
CL
Q
<
O
b
o
u
<
l/l
CL
o
QC
o
u
£
u
QC
35
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
<
>
DC
DC
Cl
Q
2
<
Z
g
b
o
u
<
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
4.2 Water Chemistry Samples
4.2.1 Summary of Method
The water chemistry samples will be analyzed for total phosphorus (TP), total nitrogen (TN),
total ammonium (NH4), nitrate (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-o) from the X-
site at the midpoint of the stream. After collection, store all samples on ice in a closed cooler.
After you filter the chlorophyll-o sample, the filters must be kept frozen until ready to ship.
4.2.2 Equipment and Supplies
Table 4.3 provides the equipment and supplies needed to collect water samples at the X-site.
Record the water sample collection and preservation data on the Sample Collection Form, as
seen in Figure 4.2.
Table 4.3 Equipment and Supplies: Water Chemistry Sample Collection and Preservation
For collecting samples
Nitrile gloves
4 L cube container
2 L amber Nalgene bottle
3 L Nalgene beaker
Cooler with ice
Dry Ice
Plastic electrical tape
Dl water (for cleaning beaker and 2 L amber bottle between sites)
Field Operations Manual and laminated Quick Reference Guide
For recording
measurements
Sample Collection Form
Water Chemistry sample label with pre-printed Sample ID
Clear tape strips
Sample Collection Form
Pencils (for data forms)
Fine tipped indelible markers
4.2.3 Water Chemistry and Chlorophyll-a Sampling Procedure
Table 4.4 presents step-by-step procedures for collecting water chemistry samples at wadeable
sites.
<
>
QC
QC
CL
Q
<
o
b
o
u
<
l/l
CL
o
QC
o
u
£
u
cc
37
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 4.4 Procedure: Water Chemistry and Chlorophyll-a Sample Collection (Wadeable Sites)
Sampling Procedure
Water Chemistry
1.
2.
4.
5.
7.
8.
Fill out the pertinent information (Site ID and date) on the water chemistry label and affix the label
to the cube container. Completely cover the label with clear tape.
Collect the water samples from the X-site in a flowing portion near the middle of the stream. Be
sure to collect the water samples prior to any disturbance of the stream upstream of the X-site.
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.
Remove the cube container lid and expand the cube container by pulling out the sides if needed
(the weight of the water alone while filling will often open the container sufficiently). NOTE: DO
NOT BLOW into the cube container or place your fingers inside the opening to expand it, because
this will cause contamination.
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.
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.
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" bubble on the sample collection form.
Record the Sample ID on the Sample Collection Form along with the pertinent stream information
(stream name, ID, date, etc.). Note anything that could influence sample chemistry (heavy rain, potential
contaminants) in the Comments section. If the sample was collected at the X-site, darken the X-site
bubble in the "Station Collected" field. If you had to move to another part of the reach to collect the
sample, place the letter of the nearest transect in the "Station Collected" field. Record more detailed
reasons and/or information in the Comments section.
If sample(s) are not collected, fill in the "No Sample Collected" bubble on the data form(s) and indicate
the reason why targeted sample(s) were not collected in the comments section.
u
QC
38
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
Site ID:
NRSA 2018/19 SAMPLE COLLECTION (Front)
Date: /
Renewed by (initial*
CHEMISTRY (CHEM) STATION COLLECTED:
(Target Volum e = 4L) O X-Site (wadeable) O Transect A fnon-wadeahle) O Other Transect:
Sample ID
i_ hilierj fC nmmerits
O
WATER COLUMN CHLOROPHYLL (WCHL) (GF/F Filter)
(Target Volum e = 1000mL; max vol =2000 mL)
Sample ID
Volume Filt*ie>1
{mil
Frozen
O
ALGAL TOXIN (Mcrocyitin) (MCX) (PETG bottle)
(Target Volum e = 500 m L)
Sample ID
Fi^zen Comments
I i i i
O
ALGAL TOXIN (Microcystis (MICZ) (HDPE bottle)
(Target Volume* 500mL)
Sample ID
Frozen Comments
> l i I I I II
o
COMPOSITE PERIPHYTQN
Composite Volume ^Nu ot Ti^ir>sects Comments
PERIPHYTON ASSEMBLAGE ID (PERI) (50-m L tube)
Sample ID
Volume (ml)
O
PERIPHYTON CHLOROPHYLL (PCHL) (GFff Filter)
Preseived Comments
Sample ID
Volume Filtered (ml) Frozen Comments
i > i i 8 i
O
PERIPHYTON BIOMASS (FBIO) (GFiC Filter)
Sample ID Volume Filtered (mi) Frozen Comments
|.. L L J-.
O
PERIPHYTON METAGENOMIC (PDNA) (PETG bottle)
Sample ID
I » i » a I
O
ENTEROCOCCI (ENTE)
(Target Volum e ¦ 250 mL (Filter blank is collected during visit 1 at all revisit sites.)}
Sample ID
i l l k i l
Time
Collected
(hhmrn)
1
Depth | Sample
Collected ^ Volume
(m) j imLj
FiL Start
Time
(hhmrn)
Volume Fiiieied
(Target = 50 ml)
No Sample Collected O
No Sample Collected 0
No Sample Collected 0
No Sample Collected 0
No Sample Collected 0
No Sample Collected 0
No Sample Collected 0
No Sample Collected 0
No Sample Collected 0
Blank Collected 0
Filt. End
T i me
fhhrnm)
Time
Frozen
(hhmm?
Gomme nts
03,CI/2018 NRSA18 Sample Collection
^157226369
<
>
QC
QC
CL
Q
<
o
b
o
u
<
CO
CL
o
ce
O
U
Figure 4.2 Sample Collection Form (front)
£
u
cc
39
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
5 ALGAL TOXINS (MICROCYSTINS and
CYLINDROSPERMOPSIN)
Cyanobacteria naturally occur in surface waters. Under certain conditions, such as in warm
water containing an abundance of nutrients, they can rapidly form harmful algal blooms (HABs).
HABs can produce toxins known as cyanotoxins, which can be harmful to humans and animals.
Microcystin and cylindrospermopsin are two cyanotoxins known to occur in the surface waters
of the United States. Microcystins are the most widespread cyanobacterial toxins and primarily
affect the liver but can also affect the kidney and reproductive system.
Cylindrospermopsin is another commonly identified cyanotoxin found in U.S. waters. The
primary toxic effects of this toxin are damage to the liver and kidney.
5.1 Summary of Method
The algal toxin (microcystin and cylindrospermopsin) samples are grab samples taken from the
X-site. All Field Crews must collect a grab sample using the 3 L beaker to fill two 500 ml bottle.
Collect these samples after the in situ measurements and water chemistry samples are
collected. Store all samples on ice in a closed cooler.
5.2 Equipment and Supplies
Table 5.1 provides the equipment and supplies needed to collect the algal toxin samples at the
index site. Record the water sample collection and preservation data on the Sample Collection
Form, as seen in Figure 4.2.
Table 5.1 Equipment and Supplies: Microcystin
For collecting samples
Nitrile gloves
3 L Nalgene beaker
PETG bottle (500 mL, clear, square) - algal toxins (MICX)
HDPE bottle (500 mL white, round) - algal toxins (MICZ)
Plastic electrical tape
Cooler with ice
Field Operations Manual and laminated Quick Reference Guide
For recording
measurements
Sample Collection Form
Algal toxin sample labels with pre-printed Sample ID
Clear tape strips
Sample Collection Form
Pencils (for data forms) and Fine tipped indelible markers for labels
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
5.3 Sampling Procedure
Table 5.2 presents step-by-step procedures for collecting algal toxin (microcystin and
cylindrospermopsin) samples at wadeable sites.
Table 5.2 Procedure: Algal Toxin (Microcystin and Cylindrospermopsin) Collection (Wadeable Sites)
Microcystin Sample Collection
1. Fill out the pertinent information (Site ID and date) on the algal toxin labels.
2. Affix the MICX label to the 500 mL PETG clear square Nalgene bottle. Completely cover the label
with clear tape.
3. Affix the MICZ label to the 500 mL HDPE white round Nalgene bottle. Completely cover the label
with tape.
4. Collect the algal toxin (microcystin and cylindrospermopsin samples from the X-site in a flowing
portion of the stream near the middle of the transect.
5. 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.
6. Rinse the 3 L Nalgene beaker three times with water, and discard the rinse downstream.
7. Rinse each water sample collection container and lid three times with water, discard the rinse
downstream.
8. Fill the beaker with water and pour into the 500 ml Nalgene bottles to the 500 mL mark (or just
below the shoulder of the bottle), leaving headspace so that the bottles don't burst when frozen.
9. Seal the caps with plastic electrical tape before shipping.
Storage
Place the 500 ml Nalgene bottles in a cooler (on ice or water) and shut the lid. If a cooler is not available,
place the 500 ml bottles in an opaque garbage bag and immerse them in the stream.
Record the Sample IDs on the Sample Collection Form along with the pertinent site information (site
name, ID, date, etc.).
Upon returning to your base site (hotel, lab, office, etc.), freeze both samples and keep frozen until
shipping. Mark the "Frozen" bubbles on the form to verify samples have been frozen.
If sample(s) are not collected, fill in the "No Sample Collected" bubble on the data form(s) and indicate
the reason why targeted sample(s) were not collected in the comments section.
i/i
CL
O
QC
LU
CL
l/l
O
QC
Q
>
U
T3
C
tc
l/l
l/l
>
U
O
QC
u
X
o
<
41
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
6 BENTHIC MACROINVERTEBRATES
6.1 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 3.3). 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.
6.2 Equipment and Supplies
Table 6.1 shows the checklist of equipment and supplies required to complete the collection of
benthic macroinvertebrates. This checklist is similar to the checklist presented in Appendix A,
which is used at the base location to ensure that all of the required equipment is brought to the
site. Record collection data on the back of Sample Collection Form (Figure 6.1).
Table 6.1 Equipment and Supplies: Benthic Macroinvertebrate Collection at 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 or laminated
Quick Reference Guide
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 of
Clear tape strips
jars
Sample Collection Form
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
TRANSECT
A
B
C
D
E
F
e
H
1
J
K
SUBSTRATE
CHAN.
Si®
Cnan
Sub tOan
sob
thm
Su$
Chtn
sma
Ch»n
Sat
Cftan
Sub
CttE>n
Suft.
Chan
S&!«5
Cfc*r»
Sue.
Cton
Sua
Citan
FirWSand
PCK>i
Of
O-
0<=tO"
OF
O"
Of
o-
Of
Op
OF
O'
Of
Of
Of
Of
Of
O"
Of
o-
Of
Op
Gr**H
aiw«
Os
O «¦
O 8 jo «¦
OS
O GL
Os
o«
o«
o«
o«
o®
o-
O oi-
o°
O s"«
OS
Oft
o«
0«-
o =
O GL
Co«rsr
Rffl*
Ot
Om
O c © R>
Oc
O
O:
O w
Oc
o«
Oc
o«
O c
Ow
Ot
o»
Ot
O i»
o =
o»
O =
O w
CTfwr
«=ipM
O OT
o **
O otJo **
Ow
o S»
O OT
O
0"T
O S*
O 07
O w
O w
o»
O"'
O
O OT
O «*
O CT
o **
O w
O
If uttw. Pag »fta
Flog
Flag
fmg
Flag
Fias
Flag
Fif>9
FlJSfl
Flag
f*a
Fhg
Site ID:
NRSA 2018/19 SAMPLE COLLECTION (Back)
Date; I
BENTHiC MACRO INVERTEBRATES (BERW) - WADEABLE
[SeseSSJ fia of
Number of jars t£JOH) Transects Comments
Ho Sample Collected Q
Sample ID
* » «
REACH-WIDE BENTHOS - WADEABLE
BENTHIC MACROINVERTEBRATES {BETB} - BOAT ABLE
Preserved! No. of
Number of jars fETOH) Transects Comments
No Sample Collected Q
Sample ID
TRANSECT BENTHOS - 80ATABL.E
Habitat: C * Coarse Substrate t IWD L * leaf Pack f * Organic fine f Sand M - Macrophyte t?ed$ O-T - Other ^Fiag ar*d e»:psait? m comment see t'on bvlow)
Substrate. F - Fiae /Sand G = GfaveJ C - Goa**** substrate OT - Qthe* 0T> be!;w>
TRANSECT
location Oc
Og Oct
0' o =
o«
Channel;
O* o«*
O w O <31
0OT
Op Os*
0SI oa
Out
O p 0 RA
Qm Qql
OOT
O? O RA
o «l O Gt
0 OT
Q? O
O w O ^
O OT
O p O R*
O RF O Gt
Oct
0 p O «a
0«! 0®-
O of
Of o^
O O c*.
0<"
8;8r
o®'
O' Ox*
O " O
O
8-8"
Oo>
B»9
Flag
F!»B
Flag
Rag
?lsg
Fiag
Fhg
Flip
F!ag
FI«b
jsst.
Comments
O2/2S/201S HRSA18 Sample Collection
8904226360
<
QC
CO
QC
LU
>
Figure 6.1 Sample Collection Form (back)
O
QC
u
<
u
CQ
43
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
6.3 Sampling Procedure
Figure 6.2 summarizes how samples will be collected from wadeable sites. The transect sample
design for collecting benthic macroinvertebrates is shown in Figure 6.3. Collect a sample from 1
m downstream of each of the 11 cross-section transects at the assigned sampling station. The
process for selecting the sample stations is described in the Initial Site Procedures (Section 3). At
transects assigned a "Center" sampling point where the stream width is between one and two
net widths wide, pick either the "Left" or "Right" sampling point instead. If the stream is only
one net wide at a transect, place the net across the entire stream width and consider the
sampling point to be "Center". If a sampling point is located in water that is too deep or unsafe
to wade, select an alternate sampling point on the transect at random.
The procedure for collecting samples at each transect is described in Table 6.2. At each sampling
point, determine if the habitat is a "riffle/run" or a "pool/glide" (any area where there is not
sufficient current to extend the net is operationally defined as a pool/glide habitat). Record the
dominant substrate type (fine/sand, gravel, coarse substrate (coarse gravel or larger) or other
(e.g., bedrock, hardpan, wood, aquatic vegetation, etc.) and the habitat type (pool, glide, riffle,
or rapid) for each sample collected on the Sample Collection Form as shown in Figure 6.1. As
you proceed upstream from transect to transect, combine all samples into a bucket.
WADEABLE
Transfer sample into bucket
Mark bubble for substrate & channel
habitat type on the Sample Collection
Form.
Thoroughly rinse net and proceed to the
next sampling location.
Composite the samples from all stations
to create a single sample for the site.
At Transect A, randomly locate 1
sampling point (left, center, or right
facing downstream)
Collect sample using riffle/run or pool/
glide procedures from 25%, 50%, &
75% width of the channel
Figure 6.2 Benthic Macroinvertebrate Collection at Wadeable Sites
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
FLOW
Combine ALL kick net samples collected from ALL transects
I
TRANSECT SAMPLES (1 per transect)
Sampling point at each transect selected systematically after random start
Sampling points proceed in L, C, R pattern upstream
Modified D-frame kick net
1 square foot quadrat sampled for 30 seconds
I
COMPOSITE SAMPLES FROM ALL TRANSECTS
• Seive bucket or other bucket(s)
SIEVE SAMPLE
• 500 jjm sieve bucket
• Remove and wash large objects
COMPOSITE AND PRESERVE SAMPLE
1 liter bottle(s) (max of 4 bottles if possible)
Fill no more than 50% with sample
Preserve with ~95% ethanol for a final con-
centration of at least 70%
1 L
Figure 6.3 Transect Sample
Design for Collecting Benthic Macroinvertebrates at Wadeable Sites
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 6.2 Procedure: Benthic Macroinvertebrates (Wadeable Sites)
Collecting Benthic Macroinvertebrate Sample
1. At 1 m downstream of each transect, beginning with Transect "A", randomly locate the first sampling
station (Left, Center, or Right as you face downstream) as 25%, 50%, and 75% of the wetted width,
respectively. If you cannot collect a sample at the designated point because of deep water or unsafe
conditions, relocate to another random point on the same transect.
2. Determine if there is sufficient current in the area at the sampling station to fully extend the net. If
so, classify the habitat as "riffle/run" and proceed to Step 3. If not, use the sampling procedure
described for "pool/glide" habitats starting at Step 9.
NOTE: If the net cannot be used, hand-pick a sample for 30 seconds from about 1 ft2 of substrate at the
sampling point. For vegetation choked sampling points, sweep the net through the vegetation within a 1
ft2 quadrat for 30 seconds. Place this hand-picked sample directly into the sample container. Assign a
"U"flag (non-standard sample) to the sample and indicate which transect(s) required the modified
collection procedure in the comments section. Go to Step 13.
Riffle/Run Habitats:
6.
8.
With the net opening facing upstream, quickly position the net securely on the stream bottom to
eliminate gaps under the frame. Avoid large rocks that prevent the net from seating properly on the
stream bottom.
NOTE: If there is too little water to collect the sample with the D-net, randomly pick up 10 rocks from the
riffle and pick and wash the organisms off them into a bucket which is half full of water.
Holding the net in position on the substrate, visually define a quadrat that is one net width wide and
long upstream of the net opening. The area within this quadrat is 1 ft2.
Check the quadrat for heavy organisms, such as mussels and snails. Remove these organisms by hand
and place them into the net. Pick up loose rocks or other larger substrate particles in the quadrat.
Use your hands to dislodge organisms and wash them into the net. Scrub all rocks that are golf ball
sized or larger and which are at least halfway into the quadrat. After scrubbing, place the substrate
particles outside of the quadrat.
Hold the D-net securely in position. Starting at the upstream end of the quadrat, vigorously kick the
remaining finer substrate within the quadrat for 30 seconds (use a stopwatch).
NOTE: For samples located within dense beds of long, filamentous aquatic vegetation (e.g., algae or
moss), kicking within the quadrat may not be sufficient to dislodge organisms in the vegetation. Usually
these types of vegetation are lying flat against the substrate due to current. Use a knife or scissors to
remove only the vegetation that lies within the quadrat (i.e., not entire strands that are rooted within
the quadrat) and place it into the net.
Pull the net up out of the water. Immerse the net in the stream several times to remove fine
sediments and to concentrate organisms at the end of the net. Avoid having any water or material
enter the mouth of the net during this operation.
Go to Step 13.
Pool/Glide Habitats:
9.
10.
Visually define a quadrat that is one net width wide and long at the sampling point. The area within
this quadrat is 1 ft2.
Check the quadrat for heavy organisms, such as mussels and snails. Remove these organisms by hand
and place them into the net. Pick up loose rocks or other larger substrate particles in the quadrat.
Use your hands to dislodge organisms and wash them into the net. Scrub all rocks that are golf ball
sized or larger and which are at least halfway into the quadrat. After scrubbing, place the substrate
particles outside of the quadrat.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
11. Vigorously kick the remaining finer substrate within the quadrat with your feet while dragging the net
repeatedly through the disturbed area just above the bottom. Keep moving the net all the time so
that the organisms trapped in the net will not escape. Continue kicking the substrate and moving the
net for 30 seconds.
NOTE: If there is too little water to use the kick net, stir up the substrate with your gloved hands and use
a sieve with 500 jim mesh size to collect the organisms from the water in the same way the net is used
in larger pools.
12. After 30 seconds, remove the net from the water with a quick upstream motion to wash the
organisms to the bottom of the net.
All samples:
13. Invert the net into a sieve bucket and transfer the sample. Remove as much gravel as possible so that
the organisms do not get damaged. Inspect the net for any residual organisms clinging to the net and
deposit them into the bucket. Use forceps if necessary to remove organisms from the net. Carefully
inspect any large objects (such as rocks, sticks, and leaves) in the bucket and wash any organisms
found off of the objects and into the bucket before discarding the object. Remove as much detritus as
possible without losing organisms.
NOTE: It is recommended that crews carry a sample bottle containing a small amount ofethanol with
them to enable them to immediately preserve larger predaceous invertebrates such as hellgrammites
and water beetles. Doing so will help reduce the chance that other specimens will be consumed or
damaged prior to the end of the field day.
14. Determine the predominant substrate size/type you sampled within the sampling quadrat. Mark the
sampled substrate type on the Sample Collection Form under the pertinent transect heading. The
substrate types are:
• Fine/sand (F): not gritty (silt/clay/muck <0.06 mm diam.) to gritty, up to ladybug sized (2 mm)
• Gravel (G): fine to coarse gravel (ladybug to tennis ball sized; 2 mm to 64 mm)
• Coarse (C): cobble to boulder (tennis ball to car sized; 64 mm to 4000 mm)
• Other (OT): bedrock (larger than car sized; > 4000 mm), hardpan (firm, consolidated fine
substrate), wood of any size, aquatic vegetation, etc.). Note type of "other" substrate in
comments on field form.
15. Identify the channel habitat type where the sampling quadrat was located. Indicate the channel
habitat type on the Sample Collection Form under the pertinent transect heading. The channel
habitat types are:
• Pool (P): Still water; low velocity; smooth, glassy surface; usually deep compared to other parts
of the channel
• Glide (GL): Water moving slowly, with smooth, unbroken surface; low turbulence
• Riffle (Rl): Water moving, with small ripples, waves, and eddies; waves not breaking, and surface
tension is not broken; "babbling" or "gurgling" sound.
• Rapid (RA): Water movement is rapid and turbulent; surface with intermittent "white water"
with breaking waves; continuous rushing sound.
• Other (OT): Note type of "other" channel habitat in comments on field form.
16. Thoroughly rinse the net before proceeding to the next sampling station. Proceed upstream to the
next transect (through Transect K, the upstream end of the reach) and repeat steps 1 -15. Combine
all kick net samples from riffle/run and pool/glide habitats into the bucket.
17. Record the number of transects that were sampled throughout the reach.
<
QC
CO
QC
LU
>
o
QC
u
<
u
CO
47
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
6.4 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
6.3). 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 Sample Collection Form as shown in
Figure 6.1. Place the samples in a cooler or other secure container for transporting and/or
shipping to the laboratory (see Appendix C).
Table 6.3 Procedure: Compositing Samples for Benthic Macroinvertebrates (Wadeable Sites)
Compositing Benthic Macroinvertebrate Sample
<
QC
CO
QC
O
QC
U
<
u
48
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 Ljars will be
required. Try to use no more than four jars per site.
4. Fill in a sample label with the Sample ID, date of collection, and jar # (i.e., Jar 1 of 2). Attach the
completed label to the jar and cover it with a strip of clear tape. Record the sample ID number for
the composite sample on the Sample Collection Form. For each composite sample, make sure the
number on the form matches the number on the label.
5. Wash the contents of the sieve to one side by gently agitating the sieve in the water. Wash the
sample into a jar using as little water from the wash bottle as possible. Use a large bore funnel if
necessary. If the jar is too full pour off some water through the sieve until the jar is not more than Vz
full, or use a second jar if necessary. Carefully examine the sieve for any remaining organisms and
use watchmakers' forceps to place them into the sample jar.
• If a second jar is needed, fill in a sample label that does not have a pre-printed ID number on it. Record
the ID number from the pre-printed label prepared in Step 4 in the "SAMPLE ID" field of the label. Attach
the label to the second jar and cover it with a strip of clear tape. Record the number of jars required for
the sample on the Sample Collection Form. Make sure the number you record matches the actual
number of jars used. Write "Jar N ofX" on each sample label using a waterproof marker ("N" is the
individual jar number, and "X" is the total number of jars for the sample).
6. Place a waterproof label inside each jar with the following information written with a number 2 lead
pencil:
Site ID Collectors initials
Type of sampler Number of stations sampled
Name of site
Date of collection Jar "N" of "X"
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Compositing Benthic Macroinvertebrate Sample
7. Completely fill the jar with 95% ethanol (no headspace). It is very important that sufficient ethanol
be used, or the organisms will not be properly preserved. Existing water in the jar should not dilute
the concentration of ethanol below 70%.
NOTE: Composite samples can be transported back to the vehicle before adding ethanol if necessary. In this
case, fill the jar with stream water, which is then drained using the net (or sieve) across the opening to
prevent loss of organisms, and replace with ethanol.
8. Replace the cap on each jar. Slowly tip the jar to a horizontal position, then gently rotate the jar to
mix the preservative. Do not invert or shake the jar. After mixing, seal each jar with plastic tape.
9. Store labeled composite samples in a container with absorbent material that is suitable for use with
70% ethanol until transport or shipment to the laboratory.
10. If sample(s) are not collected, fill in the "No Sample Collected" bubble on the data form(s) and
indicate the reason why targeted sample(s) were not collected in the comments section.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
7 PERIPHYTON
7.1 Summary of Method
Collect periphyton from the 11 cross-section transects ("A" through "K") established within the
sampling reach. Collect periphyton samples at the same transect location (L, C, or R) as the
benthic macroinvertebrate samples (Section 6) directly after collecting the benthic
macroinvertebrate sample. Prepare one composite sample of periphyton for each reach. At the
completion of the day's sampling activities, but before leaving the site, prepare four types of
laboratory samples (an ID/enumeration sample to determine taxonomic composition and
relative abundances, a metagenomic sample, a chlorophyll-o sample, and a biomass sample (for
ash-free dry mass [AFDM])) from the composite periphyton sample.
7.2 Equipment and Supplies
Table 7.1 is a checklist of equipment and supplies required to conduct periphyton sample
collection and processing activities. This checklist is similar to the checklist presented in
Appendix A, which is used at the base location to ensure that all of the required equipment is
brought to the site.
Table 7.1 Equipment and Supplies: Periphyton (Wadeable Sites)
For collecting samples
Large Funnel (15-20 cm diameter)
12 cm2 area delimiter (3.8 cm diameter pipe, 3 cm tall)
Stiff-bristle toothbrush with handle bent at 90° angle
1 L wash bottle for Dl water
500 mL graduated plastic bottle for the composite sample
60 mL plastic syringe with tip removed, and length of tubing (20 mL)
Timer or stopwatch
Cooler (small soft-sided preferred)
Wet ice
Field Operations Manual and a laminated Quick Reference Guide
For recording measurements
Sample Collection Form
Soft (#2) lead pencils for recording data on field forms
Fine-tipped indelible markers for filling out sample labels
Sample labels (4 per site) with the same Sample ID Number
Clear tape strips for covering labels
For cleaning equipment
10% Bleach solution
7.3 Sampling Procedure
At each of the 11 transects, collect samples from the sampling station assigned during the layout
of the reach (L, C, or R). Collect the substrate selected for sampling from a depth no deeper than
0.5 m. If a sample cannot be collected because the location is too deep, pick another random
spot along the transect. The procedure for collecting samples and preparing a composite sample
is presented in Table 7.2. Collect one sample from each of the transects and composite into one
bottle to produce one composite sample for each site. Record number of transects sampled and
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
the total the volume of the composite sample on the Sample Collection Form as shown in Figure
4.2.
Table 7.2 Procedure: Collecting Composite Index Samples of Periphyton (Wadeable Sites)
Periphyton Composite Sample
Starting with Transect "A", collect a single sample from the assigned sampling station using the
procedure below.
If coarse substrate (cobbles, woody materials, etc.) are present that can be removed from the
water:
(a) Collect a sample of substrate (rock or wood) that is small enough (< 15 cm diameter) and can
be easily removed from the water. Place the substrate in or over a plastic funnel which drains
into a 500 mL plastic bottle with volume graduations marked on it.
(b) Use the area delimiter to define a 12 cm2 area on the upper surface of the substrate. Dislodge
attached periphyton from the substrate within the delimiter into the funnel by brushing with
a stiff-bristled toothbrush for 30 seconds. Take care to ensure that the upper surface of the
substrate is the surface that is being scrubbed, and that the entire surface within the
delimiter is scrubbed.
(c) Fill a wash bottle with 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.
(d) 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).
If large coarse substrate is present that is too large to remove from the water (bedrock, large
woody materials, boulders, etc.):
(a) Use the area delimiter to define a 12 cm2 area on the upper surface of the substrate. Dislodge
attached periphyton from the substrate within the delimiter using the clear tube attached to
the tip of the syringe in a scraping motion.
(b) While dislodging periphyton with the tube, simultaneously pull back to 25 mL on the syringe
plunger to draw the dislodged periphyton into the syringe. The 25 mL in the syringe combined
with the 20 mL in the tube equals the target volume of 45 mL.
(c) Empty the syringe and tube into the same 500 mL plastic bottle as above. If the volume of the
vacuumed sediment is not enough to raise the composite volume to the next graduation on
the bottle (~45 mL), add additional stream water to the bottle to raise the level to the next
graduation.
(d) Put the bottle in a cooler on ice while you travel between transects and collect the
subsequent samples. (The sample needs to be kept cool and dark because a chlorophyll
sample will be filtered from the composite.)
If no coarse sediment (cobbles or larger) are present:
(a) Use the area delimiter to confine a 12 cm2 area of soft sediments.
(b) Vacuum the top 1 cm of sediments from within the delimited area into a de-tipped 60 mL
syringe with attached clear tube up to the 25 mL line of the syringe.
(c) Empty the syringe into the same 500 mL plastic bottle as above. If the volume of the
vacuumed sediment is not enough to raise the composite volume to the next graduation on
the bottle (~45 mL), add additional stream water to the bottle to raise the level to the next
graduation.
(d) Put the bottle in a cooler on ice while you travel between transects and collect the
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
subsequent samples. (The sample needs to be kept cool and dark because a chlorophyll
sample will be filtered from the composite.)
2. Repeat Step 1 for transects "B" through "K". Place the sample collected at each sampling station into
the single 500 mL bottle to produce the composite index sample.
Storage
3. After samples have been collected from all 11 transects (or as many transects as possible), thoroughly
mix the 500 mL bottle regardless of substrate type. Record the total volume of the composite sample
in the periphyton section of the Sample Collection Form.
4. Record the number of transects sampled. If all 11 transects are not sampled, record the reason(s) for
any missed transects on the field form.
5. If sample(s) are not collected at all, fill in the "No Sample Collected" bubble on the data form(s) and
indicate the reason why targeted sample(s) were not collected in the comments section
Clean up
6. After preparing the four types of laboratory samples (see Section 13.3), 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.
7.4 Sample Processing in the Field
You will prepare four different types of laboratory samples from the composite sample: an
ID/enumeration sample (to determine taxonomic composition and relative abundances), a
metagenomic sample, 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 13)
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.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8 PHYSICAL HABITAT CHARACTERIZATION
Field measurements for physical habitat are made at two scales of resolution along the mid-
channel length of the reach, and the results are later aggregated and expressed for the entire
reach. The protocol defines the length of each sampling reach proportional to stream channel
wetted width and then systematically places measurements to statistically represent the entire
reach.
8.1 Equipment and Supplies
Table 8.1 lists the equipment and supplies required to conduct all the activities described for
characterizing physical habitat. This checklist is similar to the checklist presented in Appendix A,
which is used at the base location to ensure that all of the required equipment is brought to the
stream. Use this checklist to ensure that equipment and supplies are organized and available at
the river site in order to conduct the activities efficiently.
Table 8.1 Equipment and Supplies: Physical Habitat
For making
measurements
Convex spherical canopy densiometer (Lemmon Model B), modified with taped "V"
GPS
1 roll each colored surveyor's flagging tape (2 colors)
2 pair chest waders
1 or 2 fisherman's vest with lots of pockets and snap fittings.
Digital camera with extra memory card & battery (optional)
50 m or 100 m measuring tape with reel
Meter stick for bank angle measurements
Laser rangefinder (400 ft. distance range) and clear waterproof bag
Clinometer
Binoculars (optional)
Bearing compass
Current velocity meter, probe, and operating manual
Top-set wading rod for use with current velocity meter
Surveyor's telescoping leveling rod
Sounding rod or wading staff
Level tripod
CST Berger SAL 20 Automatic Level
Field Operations Manual and/or laminated Quick Reference Guide
For recording
data
2 covered clipboards (lightweight, with strap or lanyard)
Soft (#2) lead pencils
11 plus extras Channel/Riparian Transect forms
11 plus extras Thalweg Profile forms
1+ extras field Form: Stream Verification Form
1+ extras field Form: Field Measurement Form
1+ extras field Form: Sample Collection Form
1+ extras field Form: Channel Constraint Form
1+ extras field Form: Visual Assessment Form
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.2 Summary of Methods Approach
Physical habitat in streams includes all those physical attributes that influence or provide
sustenance to organisms within the stream. The physical habitat of a stream varies naturally,
thus expectations differ even in the absence of anthropogenic disturbance. The procedures are
employed on a reach length 40 times its mean wetted width at the time of sampling.
Measurement points are systematically placed to statistically represent the entire reach. Stream
depth and wetted width are measured at very tightly spaced intervals, whereas channel
cross-section profiles, substrate, bank characteristics and riparian vegetation structure are
measured at larger intervals. Woody debris is tallied along the full length of the sampling reach,
and discharge is measured at one location. The tightly spaced depth and width measures allow
calculation of indices of channel structural complexity, objective classification of channel units
such as pools, and quantification of residual pool depth, pool volume, and total stream volume.
8.3 Components of the Habitat Characterization
There are six components of the physical habitat characterization (Table 8.2). Measurements
are recorded using the NARS App or on 11 copies of a two-sided field form, with separate forms
for recording slope and bearing measurements, assessing the degree of channel constraint,
recording evidence of debris torrents or recent major flooding, and for stream discharge
measurements.
Table 8.2 Summary of Components of Physical Habitat Characterization at Wadeable Sites
Component Description
<
M
QC
<
QC
<
I
u
CO
<
<
y
i/i
>
54
Thalweg Profile
• Measure maximum depth, classify habitat and check presence of backwaters,
side channels and loose, soft deposits of sediment particles at 10-15 equally
spaced intervals between each of 11 transects (100 or 150 individual
measurements along entire reach) The number of thalweg measurements is
specified by the stream's mean wetted width.
Wetted Width /
Bar Width
• Measure wetted width and bar width (if present) and evaluate substrate
particle size classes at 11 cross-section transects and midway between them
(21 width measurements and substrate notations along entire reach).
Woody Debris
Tally
• Between each of the channel cross-sections, tally large woody debris numbers
within and above the bankfull channel according to specified length and
diameter classes (10 separate tallies).
Channel and
Riparian
Characterization
• At 11 transects placed at equal intervals along reach:
• Measure channel cross-section dimensions, bank height, bank undercut
distance, bank angle, slope and compass bearing (backsight), and riparian
canopy density (with densiometer).
• Visually estimate3: substrate size class, embeddedness and water depth at
five equidistant points on cross-section; areal cover class and type (e.g.,
woody trees) of riparian vegetation in canopy, understory, and ground
cover; areal cover class of fish concealment features, aquatic macrophytes
and filamentous algae.
• Observe and record3: presence and proximity of human disturbances.
• At 10 cross-sections that are midway between the 11 transects above:
• Visually estimate3 substrate size class at 5 equidistant points on each
cross-section
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Assessment of
Channel
Constraint, Debris
Torrents, and
Major Floods
• After completing thalweg and transect measurements and observations,
identify features causing channel constraint, estimate the percentage of the
channel margin that is constrained for the whole reach, and estimate the
bankfull and valley widths. Check for evidence of recent major floods and
debris torrent scour or deposition.
Discharge
• Measure water depth and velocity at 15 to 20 equally spaced intervals across
one carefully chosen channel cross-section.
• In very small streams, measure discharge by timing the passage of a neutrally
buoyant object through a segment whose cross-sectional area has been
estimated or by timing the filling of a bucket.
a Substrate size class is estimated for a total of 105 particles taken at 5 equally spaced points along each
of 21 cross-sections. Depth is measured and embeddedness estimated for the 55 particles located along
the 11 regular transects A through K. Cross-sections are defined by laying the surveyor's rod or tape to
span the wetted channel. Woody debris is tallied over the distance between each cross-section and the
next cross-section upstream. Riparian vegetation and human disturbances are observed 5m upstream
and 5m downstream from the cross-section transect. They extend shoreward 10m from left and right
banks. Fish cover types, aquatic macrophytes, and algae are observed within the channel 5m upstream
and 5m downstream from the cross-section stations. These boundaries for visual observations are
estimated by eye.
8.4 Work Flow for the Physical Habitat Components
The six components (Table 8.2) of the habitat characterization are organized into four grouped
activities described below in the following sections.
8.4.1 Thalweg Profile and Large Woody Debris Tally
Thalweg spacing is calculated such that either 10 or 15 evenly spaced measurements are made
between each transect (see Section 8.5). Two people proceed upstream from the downstream
end of the sampling reach making observations and measurements at the calculated increment
spacing. One person is in the channel making width and depth measurements and determining
whether soft/small sediment deposits are present under his/her wading staff. The other person
records these measurements, classifies the channel habitat, records presence/absence of side
channels and off-channel habitats (e.g., backwater pools, sloughs, alcoves), and tallies large
woody debris. Each time the crew reaches a flag marking a new cross-section transect, they
start filling out a new copy of the Thalweg Profile Form. They interrupt the thalweg profile and
woody debris tallying activities to complete data collection at each cross-section transect as
they come to it. When the crew member in the water makes a width measurement at channel
locations midway between regular transects (i.e. at the fifth or seventh thalweg measurement in
each sub-reach), she or he also locates and estimates the size class of the substrate particles on
the left channel margin (0%) and at positions 25%, 50%, 75%, and 100% of the distance across
the wetted channel. Procedures for this substrate tally are the same as for those at regular
cross-sections, but data are recorded on the thalweg profile side of the field form.
8.4.2 Channel/Riparian Cross-Sections
At each of the 11 transects, one person proceeds with the channel cross-section dimension
measurements and substrate observations as described above; and also makes measurements
of bank angle (using rod and clinometer) and canopy cover (using densiometer). The second
person records those measurements on the Channel/ Riparian Cross-section Form while making
visual estimates of riparian vegetation structure, instream fish cover, and human disturbance
specified on that form. Slope is measured by measuring the difference in elevation between
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
each transect and bearing is determined by backsighting to the previous transect.
Supplementary points may need to be located and flagged (using a different color) if the stream
is extremely brushy, sinuous, or steep to the point that you cannot sight for slope and bearing
measures between two adjacent transects.
The work flow for the thalweg profile and channel cross-section described above can be
modified by delaying the measurements for slope and bearing and/or the woody debris tally
until after reaching the upstream end of the reach. Backsighting and/or wood tallies can be
done on the way back downstream (note that in this case, the slope and bearing data form
would have to be completed in reverse order). Crews may also elect to return to Transect A and
record slope and bearing measurements on a second trip upstream through the reach.
8.4.3 Channel Constraint and Torrent Evidence
After completing observations and measurements along the thalweg and at all 11 transects, the
field crew completes the overall reach assessments of channel constraint and evidence of debris
torrents and major floods.
8.4.4 Stream Discharge
Discharge measurements can be performed during or at the end of the field sampling, but must
be made after collecting the water chemistry sample so as not to disturb the water and
compromise the sample. The measurements are taken at a chosen optimal cross-section (but
not necessarily at a sampling transect) near the X-site. However, do not use the electromagnetic
current meter close to where electrofishing is taking place. Furthermore, if a lot of channel
disruption is necessary and sediment must be stirred up, wait on this activity until all chemical
and biological sampling has been completed.
8.5 Habitat Sampling Locations within the Reach
Measurements are made at two scales along the length of the reach; the results are later
aggregated for the entire reach using procedures described by Kaufmann et al. (1999). Figure
8.1 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. In addition, they must be sufficiently close together that
they do not miss deep areas and major habitat units.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Follow these guidelines for choosing the increment between thalweg profile measurements:
Channel Width < 2.5 m:
• Minimum reach length of 150 m is used
• Thalweg increment = 1.0 m (resulting in 15 thalweg measurements per subreach)
• A total of 150 evenly spaced thalweg profile measurements will be made, 15 between
each channel cross-section
• Mid-subreach measurements are made at the 7th thalweg location
Channel Width 2.5 to 3.5 m:
• Minimum reach length of 150 m is used
• Thalweg increment = 1.5 m (resulting in 10 thalweg, measurements per subreach)
• A total of 100 evenly spaced thalweg profile measurements will be made, 10 between
each channel cross-section
• Mid-subreach measurements are made at the 5th thalweg location
Channel Width > 3.5 m:
• Reach length is 40 times channel width (maximum of 4 km)
• Thalweg increment = 0.01 x reach length (resulting in 10 thalweg measurements per
subreach)
• A total of 100 evenly spaced thalweg profile measurements will be made, 10 between
each channel cross-section
• Mid-subreach measurements are made at the 5th thalweg location
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
<
y
i/i
>
/
*
*
*
*
r= K, ' 3m | 3m -
NIPAKIAN
PLOT
eu -<>
*
*
*
*
*
M«f—Pi •
<
M
CC
~.^Ch an n el/Riparian
Cross-section,
ran sect
*
«
*
*
*
' V
Thalweg
profile
stations
>, ' -it
*
Intermediate transects (width and
substrate measurements only
Woody
Debris
Tally
r( between
transects)
<
QC
<
T.
U
CO
<
Downstream end
of sampling reach ^
PRK/DVP 8A]6
Figure 8.1 Reach Layout for Physical Habitat Measurements for streams 2.5m or greater (plan view)
58
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.5.1 Thalweg Profile and Large Woody Debris Measurements
8.5.1.1 Thalweg Profile
"Thalweg" refers to the flow path of the deepest water in a stream channel. The thalweg profile
is a longitudinal survey of maximum flow path depth. Data from the thalweg profile allows
calculation of indices of residual pool volume, stream size, channel complexity, and the relative
proportions of habitat types such as riffles and pools.
The procedure for obtaining thalweg profile measurements is presented in Table 8.3. Record
data on the Thalweg Profile and Woody Debris Data Form as shown in Figure 8.2. Use the
surveyor's rod and a metric ruler or calibrated rod or pole to make the required depth and width
measurements at each station, and to measure off the distance between stations as you
proceed upstream. You may need to make minor adjustments to align each 10th measurement
to be one increment short of the next transect. In streams with average widths less than 2.5 m,
make thalweg measurements at 1 meter increments. Because the minimum reach length is set
at 150 meters, there will be 15 measurements on a field data form: Station 0 at the transect plus
14 additional stations between it and the next transect upstream. Use the five extra lines on the
thalweg profile portion of the data form (Figure 8.2) to record measurements 10-14.
Measure thalweg depths at all stations. Flag any missing measurements using a K code and
explain the reason in the comments section of the field data form. At points where a direct
depth measurement cannot be made, make your best estimate of the depth, record it on the
field form, and flag the value using a U code (nonstandard measurement), explaining that it is an
estimated value in the comments section of the field data form. Where the thalweg points are
too deep for wading, measure the depth by extending the surveyor's rod or weighted line at an
angle to reach the thalweg point. On the field form enter a U code in the thalweg depth column
(to indicate a nonstandard technique), and record the water level on the rod or line, and the
rod/line angle, as determined using the external scale on the clinometer (vertical = 90) in the
comment field (i.e. "depth of xx cm taken at an angle of xx degrees"). If a direct measurement of
the thalweg depth is not possible, make the best estimate you can of the depth, record it, and
use a U flag and a comment to note it is an estimated value.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 8.3 Procedure: Thalweg Profile
<
M
QC
<
QC
<
I
u
CO
<
<
y
i/i
>
60
1. Determine the increment distance between measurement stations based on the wetted width used
to determine the length of the reach. Using a laser rangefinder or surveyor's rod:
• For widths < 2.5 m, establish stations every 1 m (150 total).
• For widths > 2.5 and <3.5 m, establish stations every 1.5 m (100 total).
• For widths > 3.5 m, establish stations at increments equal to 0.01 times the reach length (100
total).
2. Complete the header information on the Thalweg Profile and Woody Debris Form, noting the
transect pair (downstream to upstream). Record the increment distance determined in Step 1 in the
Increment field on the field data form used for subreach A-B. Also record the total reach length on
this form. It is not necessary to repeat these calculations on the remainder of the Thalweg Profile
and Woody Debris forms.
3. Begin at the downstream end (station 0) of the first transect (transect A).
4. Measure the wetted width at station 0, and at either station 5 (if the stream width defining the
reach length is > 2.5 m), or station 7 (if the stream width defining the reach length is < 2.5 m).
Wetted width is measured across and over mid-channel bars and boulders. Record the width on the
field data form to the nearest 0.1 m. For streams with interrupted flow, where no water is in the
channel at the station or transect, record zeros for wetted width.
NOTE: If a mid-channel bar is present at a station where wetted width is measured, measure the
wetted width across and including the bar, but also measure the bar width and record it on the field
data form.
5. At station 5 or 7 (see above) classify the size of the bed surface particle at the tip of your depth
measuring rod at the left wetted margin (0%) and at positions 25%, 50%, 75%, and 100% of the
distance across the wetted width of the stream. This procedure is identical to the substrate size
evaluation procedure described for regular channel cross-sections (transects A - K), except that for
these midway supplemental cross-sections, substrate size is entered on the thalweg profile side of
the field form (in the substrate section).
6. At each thalweg profile station, use a calibrated pole or rod to locate the deepest point within the
deepest flow path (the thalweg), which may not always be found at mid-channel (and may not
always be the absolute deepest point in every channel cross-section). Measure the thalweg depth to
the nearest cm from the substrate surface to the water surface, and record it on the thalweg profile
form. Read the depth on the side of the rod to avoid inaccuracies due to the wave formed by the
rod in moving water.
NOTE: For streams with interrupted flow if there is no water at a transect, record zeros for depth.
NOTE: Obtain thalweg depths at all stations. If the thalweg is too deep to measure directly, stand in
shallower water and extend the surveyor's rod or pole at an angle to reach the thalweg. Determine
the angle by resting the clinometer on the upper surface of the rod and reading the angle on the
external scale of the clinometer. Record a Uflag in the thalweg depth column to indicate a non-
standard procedure was used. Record the water level on the rod and the rod angle in the comments
section of the field data form. For deeper depths, use the same procedure with a taut string as the
measuring device. Tie a weight to one end of a length of string or fishing line, and toss the weight
into the deepest channel location. Draw the string up tight and measure the length of the line that is
under water. Measure the string angle with the clinometer exactly as done for the surveyor's rod. If a
direct measurement cannot be obtained, make the best estimate you can of the thalweg depth, and
use a Uflag to identify it as an estimated measurement.
7. At the point where the thalweg depth is determined, observe if unconsolidated, loose (soft)
deposits of small diameter (<16mm) sediments are present directly beneath your ruler, rod, or pole.
Soft/small sediments are defined here as fine gravel, sand, silt, clay or muck readily apparent by
"feeling" the bottom with the rod. Record presence or absence in the Soft/Small Sediment field on
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
the field data form. Note: A thin coating of fine sediment or silty algae coating the surface of cobbles
should not be considered soft/small sediment. However, fine sediment coatings should be identified
in the comments section of the field form when determining substrate size and type.
8. Determine the channel unit code (located at the bottom of the thalweg profile section of the form)
for the station. Record this on the field data form using the standard codes provided. For dry and
intermittent streams, where no water is in the channel, record habitat type as dry channel (DR).
9. If the station cross-section intersects a mid-channel bar, indicate the presence of the bar by filling in
the 'present' bubble in the Bar Width field on the field data form. However, a measurement of the
bar width is only taken if the bar intersects a station at either the endpoint or midpoint of a
subreach (e.g., station 0 and station 5 or 7).
10. Record the presence or absence of a side channel at the station's cross-section in the Side Channel
field on the field data form.
11. Record the presence or absence of quiescent off-channel aquatic habitats, including sloughs, alcoves
and backwater pools in the Backwater column of the field form.
12. Proceed upstream to the next station, and repeat Steps 2 through 11.
13. Repeat Steps 2 through 12 until you reach the next transect. At this point complete Channel/
Riparian measurements at the new transect (Section 8.4.2). Then prepare a new Thalweg Profile
and Woody Debris Form and repeat Steps 2 through 12 for each of the reach segments, until you
reach the upstream end of the sampling reach (Transect K). At Transect K, you will have completed
10 copies of the Thalweg Profile and Woody Debris Form, one for each subreach (A to B, B to C,
etc.).
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
Figure 8.2 Thalweg Profile and Woody Debris Form
<
N
ce
<
cc
<
I
u
CD
<
<
y
00
>
62
3 O O
C
•2* LU
1 q]
! oo
9 —
6 x
o
z
O tJ X
tu GO 6
O
S® 9
< < LL
500
co o
rr w
m £
uj 5
Q DC
>-
O
o
o
5
o3
O
DC
Q.
O
111
<
z
h-
m
<
X
CL
o>
O
CM
<
CO
DC
|E
IP
i* 5
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
o
£ g
GO £
S5
Q i
51
uu $
II
s!
11
° 3
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
At every thalweg increment, determine by sight or feel whether deposits of soft/small
sediments are present on the channel bottom. These particles are defined as substrate equal to
or smaller than fine gravel (< 16 mm diameter). These soft/small sediments are different from
Fines described when determining the substrate particle sizes at the cross-section transects
(Section 8.7). If the channel bottom is not visible, determine if soft/small sediment deposits are
readily obvious by feeling the bottom with your boot, the surveyor's rod, or a calibrated rod or
pole.
Measure wetted width at each transect (station 0), and midway between transects (station 5 for
larger streams having 100 measurement points, or station 7for smaller streams having 150
measurement points). The wetted width boundary is the point at which substrate particles are
no longer surrounded by free water. Estimate substrate size for five locations evenly spaced
across each midway cross-section at regular cross-sections (Figure 8.1), but at the supplemental
cross-sections, only the size class (not distance and depth) data are recorded.
While recording the width and depth measurements and the presence of soft/small sediments,
the second person evaluates and records the habitat class (Table 8.4) applicable to each of the
100 (or 150) measurement points along the length of the reach. Make channel unit scale habitat
classifications at the thalweg of the cross-section. The habitat unit itself must meet a minimum
size criteria in addition to the qualitative criteria listed in Table 8.4. Before being considered
large enough to be identified as a channel unit scale habitat feature, the unit should be at least
as long as the channel is wide. For instance, if there is a small deep (pool like) area at the
thalweg within a large riffle area, do not record it as a pool unless it occupies an area about as
wide or long as the channel is wide. If a backwater pool dominates the channel, record PO
(Pool) as the dominant habitat unit class. If the backwater is a pool that does not dominate the
main channel, or if it is an off channel alcove or slough (large enough to offer refuge to small
fishes), fill in the V bubble to indicate presence of a backwater in the Backwater column of the
field form, but classify the main channel habitat unit type according to characteristics of the
main channel. Sloughs are backwater areas having marsh like characteristics such as vegetation,
and alcoves (or side pools) are deeper areas off the main channel that are typically wide and
shallow (Helm 1985, Bain and Stevenson 1999).
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 8.4 Channel Unit Categories
Channel Unit Habitat Classes1
Class (Code)
Description
Pool (PO)
Still water, low velocity, a smooth, glassy surface, usually deep compared
to other parts of the channel
Glide (GL)
Water moving slowly, with a smooth, unbroken surface. Low turbulence.
Riffle (Rl)
Water moving, with small ripples, waves and eddies waves not breaking,
surface tension not broken. Sound: babbling, gurgling.
Rapid (RA)
Water movement rapid and turbulent, surface with intermittent white-
water with breaking waves. Sound: continuous rushing, but not as loud as
cascade.
Cascade (CA)
Water movement rapid and very turbulent over steep channel bottom.
Much of the water surface is broken in short, irregular plunges, mostly
Whitewater. Sound: roaring.
Falls (FA)
Free falling water over a vertical or near vertical drop into plunge, water
turbulent and white over high falls. Sound: from splash to roar.
Dry Channel (DR)
No water in the channel, or flow is submerged under the substrate
(hyporheic flow).
8.5.1.2 Large Woody Debris Tally
Large Woody Debris (LWD) is defined here as woody material with a small end diameter of at
least 10 cm (4 in.) and a length of at least 1.5 m (5 ft.). This includes any portion of woody
material that meets those minimum size categories (e.g., include any LWD piece with at least 1.5
meters of its length having a diameter of 10 cm or greater). The procedure for tallying LWD is
presented in Table 8.5. The tally includes all pieces of LWD that are at least partially in the
baseflow channel (Zone 1), in the bankfull channel (Zone 2, flood channel up to bankfull stage),
or spanning above the bankfull channel (Zone 3), as shown in Figure 8.3. LWD in zones 1 and 2
will be tallied together (considered all or in-part in the bankfull channel). The bankfull channel is
defined as the channel that is filled by moderate sized flood events that typically recur every
one to two years. LWD in or above the bankfull channel is tallied over the entire length of the
reach, including the area between the channel cross-section transects. Pieces of LWD that are
not at least partially within Zones 1, 2, or 3 are not tallied.
<
M
QC
<
QC
<
T.
U
CO
<
^ 1 Note that in order for a channel habitat unit to be distinguished, it must be at least as wide or long as
the channel is wide (except for off channel backwater pools, which are noted as present regardless of
size)
64
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Table 8.5 Procedure: Tallying Large Woody Debris
Large Woody Debris Tally Form
Field Operations Manual
Wadeable
Note: Tally pieces of large woody debris (LWD) within each segment of stream while the thalweg profile
is being determined. Include all pieces in the tally whose large end is found within the subreach.
1. Scan the stream segment between the two cross-section transects where thalweg profile
measurements are being made.
2. Tally all LWD pieces within the segment that are at least partially within the bankfull channel.
Determine if a piece is LWD (any portion with a small end diameter >10 cm [4 in.], and a length >1.5 m
[5ft.]).
3. For each piece of LWD, determine the class based on the diameter of the large end (0.1 m to < 0.3 m,
0.3 m to <0.6 m, 0.6 m to <0.8 m, or >0.8 m), and the class based on the length of the piece (1.5m to
<5.0m, 5m to <15m, or >15m).
If the piece is not cylindrical, visually estimate what the diameter would be for a piece of wood with
circular cross-section that would have the same volume.
When estimating length, include only the part of the LWD piece that has a diameter >10 cm (4 in).
4. Place a tally mark in the appropriate diameter x length class tally box in the Pieces All/Part in Bankfull
Channel section of the Thalweg Profile and Woody Debris Form.
5. Tally all LWD pieces within the segment that are not actually within the bankfull channel, but are at
least partially spanning (bridging) the bankfull channel. For each piece, determine the class based on
the diameter of the large end (0.1 m to < 0.3 m, 0.3 m to <0.6 m, 0.6 m to <0.8 m, or >0.8 m), and the
class based on the length of the piece (1.5 m to <5.0 m, 5 m to <15 m, or >15 m).
6. When entering data via the NARS App, numbers can be types directly into tally boxes, or the "+" and
" buttons can be used to incrementally change the number.
7. When using paper data forms, Place a tally mark for each piece in the appropriate diameter x length
class tally box in the Pieces Bridge Above Bankfull Channel section of the Thalweg Profile and Woody
Debris Form. After all pieces within the segment have been tallied in the gray box, write the total
number of pieces for each diameter x length class in the white box to the right of the gray tally box.
8. Repeat Steps 1 through 7 for the next subreach, using a new Thalweg Profile and Woody Debris Form.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
bankfull channel width
water surface at
/-baseflow /
ZONE 2
ZONE 1 /
Figure 8.3 Large Woody Debris Influence Zones (modified from Robison and Beschta, 1990).
8.6 Channel and Riparian Measurements at Cross-Section
Transects
8.6.1 Slope and Bearing
Measure slope and bearing by backsighting between transects (e.g., transect B and A, C and B,
etc.). To measure the slope between adjacent transects, follow the procedure presented in
Table 8.6. Measure bearing following the procedure presented in Table 8.8. Record slope and
bearing data on the Slope and Bearing Form as shown in Figure 8.4.
8.6.1.1 Measurement of Slope using Level and Stadia Rod
Slope is typically measured by two people, one holding a surveyor's rod and the second sighting
through the surveyor's level. Be sure that the person is holding the marked pole or rod at the
surface of the water. The intent is to get a measure of the water surface slope, which may not
necessarily be the same as the bottom slope. The surveyor's level is leveled according to the
manufacturer's recommendations, which is generally to adjust the three leveling feet until the
bubble is centered. Level is checked in all planes to be measured. If the level does not "self
level" in all measured planes the user should check the instruction manual for suggested
options. Relative elevation readings are made at each transect and the difference between each
elevation reading is calculated and recorded as the change in elevation. NOTE: Multiple transect
elevations can often be made for each setup of the level, but every time the tripod and level
are moved, a second measurement of the last elevation from the last setup is required. You
cannot use elevations from previous setups because the relative height of the level has
changed.
To calculate sinuosity from bearing measurements, it does not matter whether or not you adjust
your compass bearings for magnetic declination, but it is important that you are consistent in
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
the use of magnetic or true bearings throughout all the measurements you make on a given
reach. Note in the comments section of the Slope and Bearing Form which type of bearings you
are taking, so the measurements can be used to describe reach aspect. Also, guard against
recording reciprocal bearings (erroneous bearings 180 degrees from what they should be). The
best way to do this is to know where the primary (cardinal) directions are in the field: (north [0
degrees], east [90 degrees], south [180 degrees], and west [270 degrees]), and insure that your
bearings "make sense."
As stated earlier, it may be necessary to set up intermediate (supplemental) bearing points
between a pair of cross-section transects if you do not have direct line-of-sight along (and
within) the channel between stations (Figure 8.6). This can happen if brush is too heavy, or if
there are sharp slope breaks or tight meander bends. If you would have to sight across land to
measure bearing between two transects, then you need to make one or more supplemental
measurements (i.e., do not "short-circuit" a meander bend). Mark these supplemental locations
with a different color of plastic flagging than used for the cross-section transects to avoid
confusion. Record these supplemental bearing measurements, along with the proportion of the
stream segment between transects included in each supplemental measurement, in the
appropriate sections of the Slope and Bearing Form (Figure 8.4). Note that the main bearing
observations are always downstream of supplemental observations (i.e., to the downstream
transect). Similarly, first supplemental observations are always downstream of second
supplemental observations.
Because measurements of slope are a calculation of the elevation difference between transects,
you may sight over land for the purposes of slope only (Figure 8.6). You may need to use
supplemental points in your measurement of slope if visibility is severely limited, but
supplemental points are not required for slope as they are for measurements of bearing. As a
result, there may be times when you record bearing data for a supplemental point (or two) on
the Slope and Bearing Form; but only record the total elevation change in the MAIN column.
Table 8.6 Procedure: Obtaining Slope Data
Slope Method with Surveyors Level
Instrument Setup:
1. Extend the tripod legs to approximately eye level and set the legs firmly into the ground; adjust
the legs so that they form a regular triangle and are firmly set with no wobble. Adjust the legs so
that the base plate is approximately level.
2. Hold the instrument on the tripod and start the centering screw. Ensure the adjustable feet are
roughly evenly adjusted. While the centering screw is still loose, slide the instrument on the base
plate until the bubble is approximately centered in the circular level. Tighten the centering screw.
3. Adjust the leveling foot screws until the bubble is exactly level in the center circle.
4. Self-Leveling instruments can now be swiveled gently on the base plate and should maintain level
as long as the tripod remains steady. Check to ensure the bubble indicates the instrument is level
across all planes at which measurements are to be made.
5. Adjust focus, brightness and parallax according to manufacture's specifications. The instrument is
ready to make measurements.
Taking Measurements:
1. Determine a location at transect A to hold a surveyor's rod that will be visible from a point
between transect A and transect B:
a. Set up the instrument at a point approximately halfway between Transects A and B and
where a clear line of sight is possible. If visibility allows, the instrument may be placed farther
<
M
QC
<
QC
<
I
u
CO
<
<
y
i/i
>
67
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
upstream, as long as Transect A is still visible.
Note: In small streams with a clear line of site it may be possible to set the instrument up
once and make readings to several transects from a single set up. Simply record the
readings for each transect and do not skip transects.
b. Position the staff at Transect A, holding the bottom of the staff at the water level and the
staff as vertical as possible and the numbers facing the instrument.
c. Site the staff through the instrument and record the reading to the nearest 0.5 centimeter in
a field notebook or other workspace (this value is not entered on the field form).
d. Move the staff to Transect B and gently swivel the instrument to face the next reading. Hold
the staff as before, vertically, with the bottom at the water level and the numbers facing the
instrument.
e. Site the staff and record the reading to the nearest 0.5 centimeter (again in a field notebook
or elsewhere).
f. Subtract the elevation reading at Transect A from the reading at Transect B.
g. The difference in the readings is the elevation difference that is recorded on the Slope and
Bearing Form. Also be sure to fill in the "cm" bubble to indicate the units on the value
entered.
h. Repeat measurements between each transect.
2. Proceed to the next cross-section transect (or supplementary point), and repeat Steps a - h above.
For each transect pair where the above method is used, mark the "TR" bubble (for surveyors
level/transit). If the above method cannot be used for one or more transect pair(s), see Section
8.6.1.2 for alternative methods3.
NOTE: If you are sighting to a supplemental point (required for bearing measurements in some
streams), you may record the elevation difference in the appropriate Supplemental section(s) of the
Slope and Bearing Form. You may also calculate the total elevation change from one transect to the
next and entered only that value in the form. In this case, values would be entered for bearing in the
supplemental column(s), but not for slope.
a Method codes are: CZ.=clinometer, 77?=surveyor's level / transit, HL=hand level, l/l/T"=Water tube, L4=laser
level, Omf/?=method not listed (describe in comments section of form).
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
z
° a
W I
si a
00
<
LU
G
<
5
i
o
z
DC
<
UJ
CO
G
Z
<
Ui
£
o
_l
c/>
m
<
x
Q.
o>
<
(J)
cc
z
° £
a Z
O O
? P
7
g r
SI
o r
a z
o o
s I
£ £
S a
II
sir
i=!
f- 3 o
ooo
-I _l <
a t 3
OOP
P 5 o
OOO
_J _I <
O X 3
ooo
f5o
OOO
_J —I <
O X 3
ooo
H £ o
OOO
—I _l <
O I j
ooo
'Jo
OOO
—) —I <
o X 5
ooo
P S 5
ooo
_J _i <
o X 3
ooo
H 5 S
ooo
6 t 3
ooo
ooo
d i 5
ooo
K 2 o
ooo
o x _j
ooo
h 3 c
ooo
_J _J c
o x 3
ooo
£
o
s
c
s
if
° i
£ l-
w o
® jg
= II
u
sf
-O ffl
"S *
£ D
>s
9 ?
<0 C
IS
£ 3
1! =
5 «
S|
EC
o
IS
3§
il
£ 3
® C
5;
II D>
*2
Si*
If 1
13 m
" o ?
s S "
£| 2.
•J ?
« = 2
II £T ®
?K»
5 S r-
iEo;
Figure 8.4 Slope and Bearing Form
<
M
cc
<
cc
<
X
u
CQ
<
<
u
LO
>
69
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.6,1.2 Alternate Methods for Obtaining Slope
Because of ease of use, portability, and cost, hand-held clinometers were previously used to
determine slope. In NRSA, the field crews will have access to more sophisticated
instrumentation (e.g., surveyor's level), and have field personnel who are experienced in the use
of these instruments. Clinometers should only be used if the slope is greater than 2.75% or if the
surveyors level malfunctions. However, note that when properly used, a roofers level
(hydrostatic level) can yield slope measurements more precise than even a laser level. The
Slope and Bearing Form (Figure 8.4) is designed to allow for different methods and/or different
units of measuring elevations or direct measurements of slope. Mark the appropriate method
bubble (instead of 77?; method codes are identified in Table 8.6) and mark the % BUBBLE (instead
of the CM bubble) if the method or instrument measures the percent slope rather than the
difference in elevation (Table 8.7).
Table 8.7 Modified Procedure: Obtaining Slope Data (without Surveyor's Level)
Modified Slope Method - Only Use if Surveyor Level is NOT Possible
<
M
QC
<
QC
<
I
u
CO
<
<
y
i/i
>
70
Use this procedure (Figure 8.5) if you are starting at the upstream Transect (K), after completing the
thalweg profile and other cross-section measurements at Transects A through K. It should only be used if
you cannot use the surveyor's level.
1. Stand in the channel at the upstream cross-section transect. Determine if you can see the next
transect downstream. If not, you may have to take supplementary slope measurement(s). While
sighting over land is not prohibited during any slope measurement, the clinometer method is
more difficult to use over larger distances, so supplementary measurements are more likely to be
needed.
a. If the next transect downstream still cannot be seen, you will have more than one
supplemental measurements (e.g., you will measure the second supplemental, then the
first supplemental, and finally the main).
2. Mark a surveyor's rod and a calibrated rod (or meter ruler) at the same height. If a shorter pole
or ruler is used, measure the height from the ground to the opening of the clinometer when it is
resting on top.
3. Have one person take the marked surveyor's rod to the downstream transect. Hold the rod
vertical with the bottom at the same level as the water surface. If no suitable location is available
at the stream margin, position the rod in the water and note the depth.
If you have determined in Step 1 that supplemental measurements are required for this
segment, walk downstream to the furthest point where you can stand in the center of the
channel and still see the center of the channel at the upstream transect. Mark this location
with a different color flagging than that marking the transects.
4. Place the base of the calibrated rod at the same relative height as the surveyor's rod (either at
the water surface or at the same depth in the water).
5. Place the clinometer on the calibrated rod at the height determined in Step 2. With the
clinometer, sight downstream to the flagged height on the surveyor's rod at the downstream
transect (or at the supplementary point).
If you are sighting to the next downstream transect, read and record the percent slope in the
Main section on the Slope and Bearing Form for the downstream transect (e.g., J < K), which is
at the bottom of the form (i.e., you are completing the form in reverse order). Record the
Proportion as 100%.
If you are sighting from a supplemental point, record the slope (%) and proportion (%) of the
stream segment that is included in the measurement in the appropriate Supplemental section
of the Slope and Bearing Form. The last sighting to a downstream transect (from either the
upstream transect or the nearest upstream supplemental point) is always recorded as the
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Main reading. This method requires the form to not only be filled out from the bottom up, but
from right to left when supplemental points are needed. If it was determined in Step 1 that
two supplemental points are needed, you will fill out the third column first.
6. Stand in the middle of the channel at the upstream transect (or supplemental point), and
backsight with your compass to the middle of the channel at the downstream transect (or
supplemental point). Record the bearing (degrees) in the same section of the Slope and Bearing
form (Supplemental or Main) as you recorded the slope in Step 5 (see Table 8.8 for details).
7. Proceed to the next cross-section transect (or to a supplementary point), and repeat Steps 3
through 6 above.
Short pole with
clinometer at
height h
Surveyor rod
with flagging at
h8ighth BacKs"^U°
Upstream
Transect
Both poles must be at water's
surface or at same depth
Downstream
Transect
PRK/DVP 6/06
Figure 8.5 Channel Slope Measurement using a Clinometer
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.6,1.3 Method for Obtaining Bearing
Table 8.8 presents the steps necessary to obtain bearing data with a compass in wadeable
streams.
Table 8.8 Procedure: Obtaining Bearing Data
Obtaining Bearing Data
<
M
QC
<
QC
<
I
u
CO
<
<
y
i/i
>
72
Use this procedure to backsight from one transect to another using a bearing sighting compass.
Readings (in degrees) are taken from the center of one transect to the center of the next transect
downstream (e.g., when using standard workflow and working upstream, you are sighting back to the
transect you just left).
It does not matter whether you use true or magnetic bearings as long as you are consistent for each
reading.
Take care not to accidentally take a reciprocal reading (one that is 180 degrees off).
1. Stand in the center of the channel at the upstream transect. If you cannot see the next transect
downstream without sighting across land, you will have to take supplementary bearing
measurements (i.e., do not "short-circuit" a meander bend).
2. Hold the compass in line with your body and sight down the "lubber line" or through the sight
window to the center of the next transect downstream (or supplemental point). It will help to have
another person at the center of the transect to which you are sighting. Remember that your line of
sight cannot "cross land."
3. While pointing the compass toward the middle of the transect or supplemental point, read the
bearing (in degrees)
• For many navigational compasses, you will rotate the bezel until the index marks are centered
over magnetic north needle. The base of the lubber line or index mark on the compass now
points to the compass heading on the bezel.
• For some direct reading compasses, the bearing will be displayed already.
4. Record the reading on the Slope and Bearing Form.
5. If you are sighting to the next downstream transect, read and record the bearing in the Main
section on the Slope and Bearing Form. Record the Proportion as 100%.
6. If you are sighting from a supplemental point, record the bearing and proportion (%) of the stream
segment that is included in the measurement in the appropriate Supplemental section of the Slope
and Bearing Form. The sighting to a downstream transect (from either the upstream transect or
the nearest upstream supplemental point) is always recorded as the Main reading.
a. The first measurement taken is from the supplemental point to the downstream
transect. This is recorded in the Main column.
b. The second measurement taken is from either the upstream transect to the first
supplemental, or from the second supplemental to the first supplemental. This is
recorded in the first supplemental column.
c. If two supplemental are needed, then the third measurement taken is from the
upstream transect to the second supplemental. This is recorded in the second
supplemental column.
d. It will be extremely rare to need more than two supplemental points, but if this were to
happen, record additional supplemental measurements in the comments section of the
form.
7. Proceed to the next transect (or supplemental point), and repeat steps 1 through 6 above.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Backsight with
compass and
record
main slope
and bearing
measurements
and % of reach
Supplemental slope
and bearing point
Backsight with
compass and record
supplemental slope
and bearing
Measurements and
% of reach
Backsight
with compass
and record
main slope
and bearing
measurements
and % of reach
P**/DVPR*»
Figure 8.6 Measurements of Bearing Between Transects
8.7 Substrate Size and Channel Dimensions
Substrate size and embeddedness are evaluated at 5 points at each of the 11 transects (refer to
Figure 8.7). Substrate size (but not embeddedness) is also evaluated at 10 additional cross-
sections located midway between each of the 11 regular transects [A-K). In the process of
measuring substrate particle sizes at each transect, the water depth at each substrate sample
point is measured (at the 10 midway cross-sections, depth to the substrate point is not
recorded). If the wetted channel is split by a mid-channel bar (Section 8.5.1), the five substrate
points are centered between the wetted width boundaries regardless of the mid-channel bar in
between. Consequently, substrate particles selected in some cross-sections may be "high and
dry." For cross-sections that are entirely dry•, make measurements across the unvegetated
portion of the channel.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
75%
Wetted
Width
50%
Wetted
Width
25%
Wetted
Width
Right
Bank
Left
Bank
calibrated
rod/pole
Surveyor's rod
measuring tape
Figure 8.7 Substrate Sampling Cross-Section
The substrate sampling points along the cross-section are located at 0, 25, 50, 75, and 100
percent of the measured wetted width, with the first and last points located at the water's edge
just within the left and right banks. The procedure for obtaining substrate measurements is
described in Table 8.9 (including all particle size classifications). Record these measurements on
the Channel/Riparian Cross-section side of the field form, as shown in Figure 8.8.
For the supplemental cross-sections midway between regular transects, record substrate size
and wetted width data on the thalweg profile side of the field form. To minimize bias in
selecting a substrate particle for size classification, it is important to concentrate on correct
placement of the measuring stick along the cross-section, and to select the particle right at the
bottom of the stick (not, for example, a more noticeable large particle that is just to the side of
the stick). Classify the particle into one of the size classes listed on the field data form (Figure
8.8) based on the middle dimension of its length, width, and depth. This median dimension
determines the sieve size through which the particle can pass. When you record the size class as
Other, describe the substrate type in the comments section of the field form.
At substrate sampling locations on the 11 regular transects (A-K), examine particles larger than
sand for surface stains, markings, and algal coatings to estimate embeddedness of all particles in
a 10 cm diameter circle around the substrate sampling point. Embeddedness is the fraction of a
particle's volume that is surrounded by (embedded in) sand or finer sediments on the stream
bottom. By definition, record the embeddedness of sand and fines (silt, clay, and muck) as 100
percent, and record the embeddedness of hardpan and bedrock as zero percent.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 8.9 Procedure: Substrate Measurement
1. Fill in the header information on page 1 of a Channel/Riparian Cross-section Form. Indicate the
cross-section transect. At the transect, extend the surveyor's rod or metric tape across the channel
perpendicular to the flow, with the "zero" end at the left bank (facing downstream).
NOTE: If a side channel is present, and contains 16 - 49% of the total flow, establish a secondary
cross-section transect. Use a separate field data form to record data for the side channel,
designating it as a secondary transect by marking both the Extra Side Channel bubble and the
associated primary transect letter (e.g., A, B, etc.). Collect all channel and riparian cross-section
measurements from the side channel as well as the primary channel.
2. Divide the wetted channel width by 4 to locate substrate measurement points on the cross-section.
In the DistLB fields of the form, record the distances corresponding to 0% (Left), 25% (LCtr), 50%
(Ctr), 75% (Rctr), and 100% (Right) of the measured wetted width. Record these distances at
Transects A-K, not at midway cross-sections.
3. Place your sharp ended meter stick or calibrated pole at the Left location (0 m). Measure the depth
and record it on the field data form.
• Depth entries at the left and right banks may be 0 (zero) if the banks are gradual.
• If the bank is nearly vertical, let the base of the measuring stick fall to the bottom (i.e., the
depth at the bank will be > 0 cm), rather than holding it suspended at the water surface.
4. Pick up the substrate particle that is at the base of the meter stick (unless it is bedrock or boulder),
and visually estimate its particle size, according to the following table. Classify the particle according
to its median diameter (the middle dimension of its length, width, and depth). Record the size class
code on the field data form (cross-section side of form for Transects A-K; special entry boxes on
Thalweg Profile side of form for midway cross-sections.)
Code
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
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 comments)
5. Evaluate substrate embeddedness as follows at each of the five points. For particles larger than
sand, examine the surface for stains, markings, and algae. Estimate the average % embeddedness of
particles in a 10 cm circle around the measuring rod. Record this value on the field data form. For
sand and smaller particles, you will not be able to pick up an individual particle, but a "pinch" of fine
particles between your fingers. Determine and record the dominant size of particles in the "pinch."
By definition, sand and fines are embedded 100%; bedrock and hardpan are embedded 0%.
6. Move to the next location on the transect, and repeat Steps 3 - 5 at each location. Repeat Steps 1 - 5
at each transect, including any additional side channel transects established if side channels are
present.
<
M
QC
<
QC
<
I
u
CO
<
<
y
i/i
>
75
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.7.1 Bank Characteristics
The procedure for obtaining bank and channel dimension measurements is presented in Table
8.10. Data are recorded in the Bank Measurements section of the Channel/Riparian Cross-section
Form as shown in Figure 8.8. Bank angle and bank undercut distances are determined on the
left and right banks at each cross-section transect. Figure 8.9 illustrates how bank angle is
determined for several different situations. Measure bank angle on wadeable streams at the
scale of approximately 0.5 m using a short (approx. 1 m long) pole. When measuring the angle,
try to insure that at least half (0.5 m) of the pole length is in contact with the bank. Other
features include the wetted width of the channel, the width of exposed mid-channel bars of
gravel or sand, estimated incision height, and the estimated height and width of the channel at
bankfull stage as described in Figure 8.10. Bankfull height and incised height are both measured
relative to the present water surface (i.e. the level of the wetted edge of the stream). This is
done by placing the base of the small measuring rod at the bankfull elevation and sighting back
to the survey rod placed at the water's edge using the clinometer as a level (i.e., positioned so
the slope reading is 0%.). The height of the clinometer above the base of the smaller rod is
subtracted from the elevation sighted on the surveyor's rod. Bankfull flows are large enough to
erode the stream bottom and banks, but frequent enough (every one to two years) to prevent
substantial growth of upland terrestrial vegetation. Consequently, in many regions, it is these
flows that have determined the width and depth of the channel.
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
1
o
Q>
T5
<75
ctt
a
UL £
o o
LU ^
oo
D -3
oo
o —
oo
00 X
OO
< 13
OO
h
0
UJ
(0
1
cc
il
gj
If
I
S I
5 o
ll
m £
lili
~ O p Q
is
h- v
—I
5
o
o
o
0
01
jr
77
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 8.10 Procedure: Measuring Bank Characteristics
<
M
QC
<
QC
<
I
u
CO
<
<
y
i/i
>
78
1. To measure bank angle, lay a meter ruler or a short (approx. 1 m long) rod down against the left
bank (determined as you face downstream), with one end at the water's edge. At least 0.5 m of
the ruler or rod should be resting comfortably on the ground to determine bank angle. If the
ground adjacent to the water's edge is not indicative of the predominant angle of the 1 meter
shoreline, it may be necessary to move the end of the rod away from the water's edge to
correctly measure the predominant angle of the 1 meter shoreline. Lay the clinometer on the rod,
and read the bank angle in degrees from the external scale on the clinometer. Record the angle in
the field for the left bank in the Bank Measurement section of the Channel/Riparian Cross-section
Form.
• A vertical bank is 90°, overhanging banks have angles >90° approaching 180°, and more
gradually sloped banks have angles <90°. To measure bank angles >90°, turn the clinometer
(which only reads 0 to 90°) over and subtract the angle reading from 180°.
• If there is a large boulder or log present at the transect, measure bank angle at a nearby point
where conditions are more representative.
2. If the bank is undercut, measure the horizontal distance of the undercutting to the nearest 0.01
m. The undercut distance is the distance from the deepest point of the undercut out to the point
where a vertical plumb line from the bank would hit the water's surface. Record the distance on
the field data form. Measure submerged undercuts by thrusting the rod into the undercut and
reading the length of the rod that is hidden by the undercutting.
3. Repeat Steps 1 and 2 on the right bank.
4. Record the wetted width value determined when locating substrate sampling points in the Wetted
Width field in the bank measurement section of the field data form. Also determine the bankfull
channel width and the width of exposed mid-channel bars (if present). Record these values in the
Bank Measurement section of the field data form.
5. While still holding the surveyor's rod as a guide, and sighting with the clinometer as a level,
examine both banks to measure and record the height of bankfull flow above the present water
level. Look for evidence on one or both banks such as:
• An obvious slope break that differentiates the channel from a relatively flat floodplain terrace
higher than the channel.
• A transition from exposed stream sediments to terrestrial vegetation.
• Moss growth on rocks along the banks.
• Presence of drift material caught on overhanging vegetation.
• A transition from flood and scour tolerant vegetation to that which is relatively intolerant of
these conditions.
6. Hold the surveyor's rod vertical, with its base planted at the water's edge. Examine both banks,
then determine the channel incision as the height up from the water surface to elevation of the
first terrace of the valley floodplain (Note, this is at or above the bankfull channel height).
Whenever possible, use the clinometer as a level (positioned so it reads 0% slope) to measure this
height by transferring (backsighting) it onto the surveyor's rod. Record this value in the Incised
Height field of the bank measurement section on the field data form.
7. Repeat Steps 1 through 6 at each cross-section transect, (including any additional side channel
transects established when islands are present). Record data for each transect on a separate field
data form.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
bank angle = clinometer reading
A
I '-"-jft bank angle = clinometer reading
B
pole rests most ^ ^ '\s 'Is ~ V ** 1
bank angle = clinometer reading
c
undercut bank
V above water -'li-f
surface <0.5m !
this is as "comfortable"
\ as pole can get within q
|,\'V1 rn of wetted edge^p
- V i
s bank angle = 180° minus
clinometer reading
41 (e.g. 180° - 30° = 150°)
> > >' 1 ' 'V' |
/ 1 undercut distance
Figure 8.9 Determining Bank Angle Under Different Types of Bank Conditions
(A) typical, (B) incised channel, (C) undercut bank (less than 0.5 m), and (Dj overhanging bank (greater
than 0.5 m).
Unfortunately, we have to depend upon evidence visible during the low flow sampling season. If
available, consult published rating curves relating expected bankfull channel dimensions to
stream drainage area within the region of interest. Graphs of these rating curves can help you
get a rough idea of where to look for field evidence to determine the level of bankfull flows.
Curves such as these are available from the USGS for streams in most regions of the U.S. (e.g.,
Dunne and Leopold 1978; Harrelson et al. 1994, Leopold 1994). To use them, you need to know
the contributing drainage area to your sample site. Interpret the expected bankfull levels from
these curves as a height above the streambed in a riffle, but remember that your field
measurement will be a height above the present water surface of the stream. Useful resources
to aid your determination of bankfull flow levels in streams in the United States are video
presentations produced by the USDA Forest Service for western streams (IJSDA Forest Service
1995) and eastern streams (USDA Forest Service 2002). ^
LU
After consulting rating curves that show where to expect bankfull levels in a given size of £3
stream, estimate the bankfull flow level by looking at the following indicators: 2:
<
First look at the stream and its valley to determine the active floodplain. This is a 6
depositional surface that frequently is flooded and experiences sediment deposition <;
under the current climate and hydrological regime. ^
<
Then look specifically for: n:
I
• An obvious break in the slope of the banks. u
• A change from water loving and scour tolerant vegetation to more drought >
tolerant vegetation. o.
79
<
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
• A change from well sorted stream sediments to unsorted soil materials.
In the absence of clear bankfull indications, consider the previous season's flooding as the best
evidence available (note: you could be wrong if very large floods or prolonged droughts have
occurred in recent years.). Look for:
• Drift debris ("sticky wickets" left by the previous seasons flooding).
• The level where deciduous leaf fall is absent on the ground (carried away by
previous winter flooding).
• Unvegetated sand, gravel or mud deposits from previous years flooding.
In years that have experienced large floods, drift material and other recent high flow markers
may be much higher than other bankfull indicators. In such cases, base your determination on
less transient indicators such as channel form, perennial vegetation, and depositional features.
In these cases, flag your data entry and also record the height of drift material in the comments
section of the field data form.
We use the vertical distance (height) from the observed water surface up to the level of the first
major valley depositional surface (Figure 8.10) as a measure of the degree of incision or
downcutting of the stream below the general level of its valley. This value is recorded in the
Incised Height field. It may not be evident at the time of sampling whether the channel is
downcutting, stable, or aggrading (raising its bed by depositing sediment). However, by
recording incision heights measured in this way and monitoring them over time, we will be able
to tell if streams are incising or aggrading.
If the channel is not greatly incised, bankfull channel height and incision height will be the same
(i.e., the first valley depositional surface is the active floodplain). However, if the channel is
incised greatly, the bankfull level will be below the level of the first terrace of the valley
floodplain, making bankfull channel height less than incision height (Figure 8.11). Bankfull height
is never greater than incision height You may need to look for evidence of recent flows (within
about one year) to distinguish bankfull and incision heights. In cases where the channel is
cutting a valley sideslope and has over-steepened and destabilized that slope, the bare
"cutbank" against the steep hillside at the edge of the valley is not necessarily an indication of
recent incision. In such a case, the opposite bank may be lower, with a more obvious terrace
above bankfull height; choose that bank for your measurement of incised height. Examine both
banks to more accurately determine incision height and bankfull height. Remember that incision
height is measured as the vertical distance to the first major depositional surface above bankfull
(whether or not it is an active floodplain or a terrace. If terrace heights differ on left and right
banks (both are above bankfull), choose the lower of the two terraces. In many cases your
sample reach may be in a "V" shaped valley or gorge formed over eons, and the slope of the
channel banks simply extends uphill indefinitely, not reaching a terrace before reaching the top
of a ridge. In such cases, record incision height values equal to bankfull values and make
appropriate comment that no terrace is evident. Similarly, when the stream has extremely
incised into an ancient terrace, (e.g., the Colorado River in the Grand Canyon), you may crudely
estimate the terrace height if it is the first one above bankfull level. If you cannot estimate the
terrace height, make appropriate comments describing the situation.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
A. Channel not Incised
First terrace on
valley bottom
above bankfull
level
No recent incision— bankfull
level at valley bottom
Downcutting over
geologic time
Active
floodplain at or near
valley bottom elevation
(Record this height)
Second
terrace
Valley Fill
B. Incised Channel
Downcutting over
geologic time
Valley Fill
Former active floodplain
no longer connected—
becomes new first terrace
above bankfull level
(Record this height),
Recent incision—
bankfull level below
first terrace of valley
bottom
Former second
terrace becomes
Former first third terrace
terrace becomes
second terrace
Figure 8.10 Schematic Showing Relationship Between Bankfull Channel and Incision
(A) Not recently incised, and (B) recently incised into valley bottom. Note level of bankfull stage relative to
elevation of first terrace (abandoned floodplain) on valley bottom. (Stick figure included for scale).
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
A) Deeply Incised Channel
From Figure 7-S (B)
Incision Height (Always
equal to or greater than
bankfull height)
Second Terrace
\
First Terrace
Former secwics
terrace becomes
Former flist BW terrace
Recent incsaion-
fconkfu lotsS
first twrace ot wiiey
bottom
ankfull
Height
(When
channel form
is not a good
indicator, use
evidence of
recent
flooding)
B) Small stream constrained in V-shaped valley
Flood-
iitolerant y
vegetation
Bankfull Height
(when channel form is
not a good indicator,
use evidence of recent
flooding, lack of
permanent flood-
intolerant vegetation
Incision Heights
Bankful Height
No incision:
No evidence of
downcutting,
vertical bank
angle, etc.)
Floods
tolerant
r_ jegriatwn
Figure 8.11 Determining Bankfull and Incision Heights
(A) Deeply Incised Channels, and (B) Streams in Deep V Shaped Valleys (Stick figure included for scale).
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.8 Canopy Cover Measurements
Canopy cover over the stream is determined at each of the 11 cross-section transects. A
spherical densiometer (model A convex type) is used (Lemmon 1957) and is provided in the
base kit to each crew. Mark the densiometer with a permanent marker or tape exactly as shown
in Figure 8.12 to limit the number of square grid intersections read to 17. Densiometer readings
can range from 0 (no canopy cover) to 17 (maximum canopy cover). Six measurements are
obtained at each cross-section transect (four measurements in each of four directions at mid-
channel and one at each bank).
Figure 8.12 Schematic of Modified Convex Spherical Canopy Densiometer.
From Mulvey et al. (1992). In this example, 10 of the 17 intersections show canopy cover, giving a
densiometer reading of 10. Note proper positioning with the bubble leveled and face reflected at the apex
of the "V".
The procedure for obtaining canopy cover data is presented in Table 8.11. Hold the densiometer
level (using the bubble level) 0.3 m above the water surface with your face reflected just below
the apex of the taped "V", as shown in Figure 8.12. Concentrate on the 17 points of grid
intersection on the densiometer that lie within the taped "V". If the reflection of a tree or high
branch or leaf overlies any of the intersection points, that particular intersection is counted as
having cover. For each of the six measurement points, record the number of intersection points
(maximum=17) that have vegetation covering them in the Canopy Cover Measurement section of
the Channel/Riparian Cross-section Form as shown in Figure 8.8.
TAPE
i
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 8.11 Procedure: Canopy Cover Measurements
Canopy Cover Measurements with Densiometer
1. At each cross-section transect, stand in the stream at mid-channel and face upstream.
2. Hold the densiometer 0.3 m (1 ft) above the surface of the stream. Level the densiometer using the
bubble level. Move the densiometer in front of you so your face is just below the apex of the taped
"V".
3. Count the number of grid intersection points within the "V" that are covered by either a tree, a leaf,
or a high branch. Record the value (0 to 17) in the CenUp field of the canopy cover measurement
section of the Channel/Riparian Cross-section and Thalweg Profile Form.
4. Face toward the left bank (left as you face downstream). Repeat Steps 2 and 3, recording the value
in the CenL field of the field data form.
5. Repeat Steps 2 and 3 facing downstream, and again while facing the right bank (right as you look
downstream). Record the values in the CenDwn and CenR fields of the field data form.
6. Move to the water's edge (either the left or right bank). Repeat Steps 2 and 3 again, this time facing
the bank. Record the value in the Left or Right fields of the field data form. Move to the opposite
bank and repeat.
7. Repeat Steps 1 through 6 at each cross-section transect (including any additional side channel
transects established when islands are present). Record data for each transect on a separate field
data form.
8.9 Visual Riparian Estimates
8.9.1 Riparian Vegetation Structure
The previous section described methods for quantifying the cover of canopy over the stream
channel. The following visual estimation procedures supplement those measurements with a
semi-quantitative evaluation of the type and amount of various types of riparian vegetation.
Riparian vegetation observations apply to the riparian area upstream five meters and
downstream five meters from each of the 11 cross-section transects. They include the visible
area from the stream back a distance of 10 m (~30 ft.) shoreward from both the left and right
banks, creating a 10 m x 10 m riparian plot on each side of the stream (Figure 8.13). The riparian
plot dimensions are estimated, not measured. On steeply sloping channel margins, the 10 m x
10 m plot boundaries are defined as if they were projected down from an aerial view. Table 8.12
presents the procedure for characterizing riparian vegetation structure and composition. Figure
8.8 illustrates how measurement data are recorded on the Channel/Riparian Cross-section
Form. Conceptually divide the riparian vegetation into 3 layers: the Canopy layer (> 5 m high),
the Understory layer (0.5 to 5 m high), and the Ground cover layer (< 0.5 m high). Note that
several vegetation types (e.g., grasses or woody shrubs) can potentially occur in more than one
layer. Similarly note that some things other than vegetation are possible entries for the Ground
cover layer (e.g., barren ground).
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
10 m
10 m
RIPARIAN
PLOT
(Left Bank)
Cross-section Transect
Flow
10 m
Instream Fish
Cover Plot
RIPARIAN
PLOT
(Right Bank)
10 m
PRK/DVP 8/06
Figure 8.13 Riparian Zone and Instream Fish Cover Plots for a Stream Cross-Section Transect
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 understory layer, record the type as None.
Estimate the areal cover separately in each of the three vegetation layers. Note that the areal
cover can be thought of as the amount of shadow cast by a particular layer alone when the sun
is directly overhead. The maximum cover in each layer is 100%, so the sum of the areal covers for
the combined three layers could add up to 300%. The four areal cover classes are Absent, Sparse
(<10%), Moderate (10 to 40%), Heavy ( 40 to 75%), and Very Heavy (>75%). These cover classes
and their corresponding codes are shown on the field data form (Figure 8.16). When rating
vegetation cover types for a single vegetation layer, mixtures of two or more subdominant
classes might all be given Sparse (1), Moderate (2), or Heavy (3) ratings. One Very Heavy cover
class with no clear subdominant class might be rated 4 with all the remaining classes rated as
either Moderate (2), Sparse (1) or Absent (0). Note that within a given vegetation layer, two
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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 8.12 Procedure: Characterizing Riparian Vegetation Structure
Riparian Vegetative Structure Measurements
1. Standing at a point along the transect where riparian observations can be made effectively, estimate
a 5 m distance upstream and downstream (10 m total length).
2. Facing the left bank (left as you face downstream), estimate a distance of 10 m back into the
riparian vegetation. On steeply sloping channel margins, estimate the distance into the riparian zone
as if it were projected down from an aerial view.
3. Within this 10 m x 10 m area, conceptually divide the riparian vegetation into 3 layers: a Canopy
Layer (>5 m high), an Understory (0.5 to 5 m high), and a Ground Cover layer (<0.5 m high).
4. Within this 10 m x 10 m area, determine the dominant woody vegetation type for the CANOPY
LAYER (vegetation >5 m high) as either Deciduous, Coniferous, broadleaf Evergreen, Mixed, or None.
Consider the layer Mixed if more than 10% of the areal coverage is made up of the alternate
vegetation type. If the canopy layer contains no vegetation or the dominant vegetation type in the
canopy layer is not woody, record the vegetation type as "None". Indicate the appropriate
vegetation type in the Visual Riparian Estimates section of the Channel/Riparian Cross-section Form.
5. Determine separately the areal cover class of large trees (>0.3 m [1 ft] diameter at breast height
[dbh]) and small trees (<0.3 m dbh) within the canopy layer. Estimate areal cover as the amount of
shadow that would be cast by a particular layer alone if the sun were directly overhead. Record the
appropriate cover class on the field data form (Chabsent: zero cover, l=sparse: <10%, 2=moderate:
10-40%, 3=heavy\ 40-75%, or 4=very heavy: >75%).
6. Look at the UNDERSTORY layer (vegetation between 0.5 and 5 m high). Determine the dominant
woody vegetation type for the understory layer as described in Step 4 for the canopy layer. If the
understory layer contains no vegetation or the dominant vegetation type in the understory is not
woody (e.g., herbaceous), record the vegetation type as "None".
7. Determine the areal cover class for woody shrubs and saplings separately from non-woody
vegetation within the understory, as described in Step 5 for the canopy layer.
8. Look at the GROUND COVER layer (vegetation <0.5 m high). Determine the areal cover class for
woody shrubs and seedlings, non-woody vegetation, and the amount of bare ground or duff (dead
organic material present as described in Step 5 for large canopy trees.
9. Repeat Steps 1 through 8 for the right bank.
10. Repeat Steps 1 through 9 for all cross-section transects (including any additional side channel
transects established when islands are present). Uses a separate field data form at each transect.
0 8.10 Instream Fish Cover, Algae, and Aquatic Macrophytes
m Over a defined area upstream and downstream of the sampling transects (Figure 8.13), crews
2j shall estimate by eye and by sounding the proportional cover of fish cover features and trophic
^ level indicators including large woody debris, rootwads and snags, brush, live trees in the wetted
^ channel, undercut banks, overhanging vegetation, rock ledges, aquatic macrophytes,
J filamentous algae, and artificial structures.
The procedure to estimate the types and amounts of instream fish cover is outlined in Table
^ 8.13. Data are recorded on the Channel/Riparian Cross-section Form as shown in Figure 8.8.
1 Estimate the areal cover of all of the fish cover and other listed features that are in the water
I
and on the banks 5 m upstream and downstream of the cross-section (Figure 8.13). The areal
cover classes of fish concealment and other features are the same as those described for
riparian vegetation (Section 8.9.1).
86
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
The entry Filamentous algae refers to long streaming algae that often occur in slow moving
waters. Aquaticmacrophytes are water loving plants, including mosses, in the stream that could
provide cover for fish or macroinvertebrates. If the stream channel contains live wetland
grasses, include these as aquatic macrophytes. WOODY DEBRIS are the larger pieces of wood that
can influence cover and stream morphology (i.e., those pieces that would be included in the
large woody debris tally [Section 8.5.1]). Brush/woody debris refers to smaller wood pieces that
primarily affect cover but not morphology. Live Trees or Roots are living trees that are within the
channel - estimate the areal cover provided by the parts of these trees or roots that are
inundated. Overhanging vegetation includes tree branches, brush, twigs, or other small debris
that is not in the water but is close to the stream (within 1 m of the surface) and provides
potential cover. For ephemeral channels, estimate the proportional cover of these trees that is
inundated during bankfull flows. Boulders are typically basketball- to car-sized particles.
Artificial structures include those designed for fish habitat enhancement, as well as in-channel
structures that have been discarded (e.g., concrete, asphalt, cars, or tires) or deliberately placed
for diversion, impoundment, channel stabilization, or other purposes.
Table 8.13 Procedure: Estimating Instream Fish Cover
Instream Fish Cover Measurements
1. Standing at a point along the transect where riparian observations can be made effectively, estimate
a 5 m distance upstream and downstream (10 m total width).
2. Examine the water and both banks within the 10 m segment of stream for the following features
and types of fish cover: filamentous algae, aquatic macrophytes, large woody debris, brush and
small woody debris, in-channel live trees or roots, overhanging vegetation, undercut banks, boulders,
and artificial structures.
3. For each cover type, estimate the areal cover. Record the appropriate cover class in the Fish
Cover/Other section of the Channel/Riparian Cross-section Form:
Chabsent: zero cover,
l=sparse: <10%,
2=moderate: 10-40%,
3=heavy\ >40-75%, or
4=very heavy: >75%).
4. Repeat Steps 1 through 3 at each cross-section transect (including any additional side channel
transects established when islands are present). Record data from each transect on a separate field
data form.
8.11 Human Influence p
<
For the left and right banks at each of the 11 detailed Channel and Riparian Cross-sections, ^
evaluate the presence/absence and the proximity of 11 categories of human influences with the ^
procedure outlined in Table 8.14. Relate your observations and proximity evaluations to the <
stream and riparian area within 5 m upstream and 5 m downstream from the station (Figure ^
8.14). Four proximity classes are used: In the stream or on the bank within 5 m upstream or ^
downstream of the cross-section transect (B), contained within the 10 m x 10 m riparian plot
but not in the stream or on the bank (C), present outside of the riparian plot (P), and absent (0). ^
Record data on the Channel/Riparian Cross-section Form (Figure 8.8). 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"). S£
87
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
You may mark more than once for the same human influence observed outside of more
than one riparian observation plot (e.g., at both Transects D and E). The rule is that you count
human disturbance items as often as you see them, BUT NOT IF you have to site through
another transect or its 10x10 meter riparian plot.
P—outside plot
(but do not sight through next
transect or pfot)
C—within riparian plot
B—on bank or in stream
Figure 8.14 Proximity Classes for Human Influences in Wadeable Streams
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 8.14 Procedure: Estimating Human Influence
Estimate Human Influence
1. Standing at a point along the transect where riparian observations can be made effectively, look
toward the left bank (left when facing downstream), and estimate a 5 m distance upstream and
downstream (10 m total width). Also, estimate a distance of 10 m back into the riparian zone to
define a riparian plot area.
2. Examine the channel, bank and riparian plot area adjacent to the defined stream segment for the
following human influences: (1) walls, dikes, revetments, riprap, and dams; (2) buildings; (3)
pavement/cleared lots (e.g., paved, graveled, dirt parking lot, foundation); (4) roads or railroads, (5)
inlet or outlet pipes; (6) landfills or trash (e.g., cans, bottles, trash heaps); (7) parks or maintained
lawns; (8) row crops; (9) pastures, rangeland, hay fields, or evidence of livestock; (10) logging; and
(11) mining (including gravel mining).
3. For each type of influence, determine if it is present and what its proximity is to the stream and
riparian plot area. Consider human disturbance items as present if you can see them from the cross-
section transect. Do not include them if you have to sight through another transect or its 10 m xlO
m riparian plot.
4. For each type of influence, record the appropriate proximity class in the Human Influence part of the
Visual Riparian Estimates section of the Channel/Riparian Cross-section Form. Proximity classes are:
B (Bank) Present within the defined 10 m stream segment and located in the stream or on the
stream bank.
C (Contained) Present within the 10 x 10 m riparian plot area.
P (Present) Present, but outside the riparian plot area.
0 (Absent) Not present within or adjacent to the 10 m stream segment or the riparian plot
area at the transect
5. Repeat Steps 1 through 4 for the right bank.
6. Repeat Steps 1 through 5 for each cross-section transect, (including any additional side channel
transects established when islands are present). Record data for each transect on a separate field
form.
8.12 Cross-section Transects on Side Channels
If the wetted channel is split by an island, and the estimated flow in the side channel is less than
or equal to 15% of the total flow, the bank and riparian measurements are made at each side of
the main channel only (the minor side channel is ignored other than to note its presence on the
Thalweg Profile Form), so one riparian plot is established on the island as shown in Figure 8.15
(side A). If an island is present that creates a major side channel containing more than 15% of g
the total flow, an additional cross-section transect is established for the side channel as shown j=
in Figure 8.15 (side B). Separate substrate, bank and riparian measurements are made for side m
channel transects. Data from the additional side channel transect are recorded on a separate lu
Channel/Riparian Cross-section Form as shown in Figure 8.16. Riparian plots established on the ^
island for each transect may overlap (and be < 10 m shoreward) if the island is less than 10 m ^
wide at the transect. Islands are distinguished from mid-channel bars by their relationship to jj
bankfull flow: Islands are not inundated at bankfull stage; bars are part of the main channel and
are inundated at bankfull flow. ^
<
i
<
y
i/i
>
89
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
A) Island and minor side channel
<15% total flow occurs in side channel
No side channel cross section transect
Note presence on field form
Riparian plot established on island
B) Island and major side channel
>15% total flow occurs in side channel
Side channel cross-section must be assessed
Two riparian plotsestablished on island [mayoverlap)
Data is recorded on additional PHab form
10 m
| FISH COVER
RIPARIAN
PLOT
(Left Bank)
10 m
Main cross-section
transect (e.g., E)
10 m
Flow
5 m
Instream Fish
Cover Plot
5 m
PLOT
(Left Bank)
transect (e.g., E)
Flow
RIPARIAN
PLOT
(Right Bank)
10 m
5m
5m
10 m
(Right Bank)
10 m
Figure 8.15 Riparian and Instream Fish Cover Plots for a Stream with Minor and Major Side Channels
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
O •
LU
oo
o -»
• o
u _
OO
co X
OO
< u
oo
z
o
LU
<
UJ
O
<
o
b
o
cc
u
2
<
DC
<
Q_
CC
<
X
VJ
03 o
<
- I
Ov Q
00
O
CM
<
on
oc
2 m
(\|
o
o
i
* §
I I
li
(S) v
^ ;
z I
sa
%Sr
, i
s
gs
p
14
si
14
¦ig
II
a "8
qj W
fE
3
I £
li
i
-e s
*S
"5 °°
II
S3
S s
I
i'
88 <
a
u!I I
gi»
s |f
i/i J -r
tis'
I III,
S = *c
!|i|
it
e £ c r
o £ c
II x |
15 11
= Ssgs
o —. rsi JQ —
** * o
SJ
I
ft
_g> "B
s ?
If
2 s
?
f
tl
|
Hi
3 g
1. ca
5 &
s -
5 3
it " i ii ii ti ii
S3 a & § £8 S bS
5 E-S5
S^E
r 2
$ sp « t
slsi
« "5 = "S
ttlSi
II
II
S I
vO
cc
s
e
Right
fN
vO
ro
Q.
3
s
U
c
91
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.13 Channel Constraint, Debris Torrents, Recent Floods, and
Discharge
8.13.1 Channel Constraint
After completing the thalweg profile and riparian/channel cross-section measurements and
observations, envision the stream at bankfull flow and evaluate the degree, extent and type of
channel constraint, using the procedures presented in Table 8.15. Record data on the Channel
Constraint Assessment Form (Figure 8.17). First, classify the stream reach channel pattern as
predominantly a single channel, an anastomosing channel, or a braided channel (Figure 8.18):
1. Single channels may have occasional in-channel bars or islands with side channels, but
feature a predominant single channel, or a dominant main channel with a subordinate
side channel.
2. Anastomosing channels have relatively long major and minor channels (but no
predominant channel) in a complex network, diverging and converging around many
vegetated islands. Complex channel pattern remains even during major floods.
3. Braided channels also have multiple branching and rejoining channels, (but no
predominant channel) separated by unvegetated bars. Channels are generally smaller,
shorter, and more numerous, often with no obvious dominant channel. During major
floods, a single continuous channel may develop
After classifying the channel pattern, determine whether the channel is constrained within a
narrow valley, constrained by local features within a broad valley, unconstrained and free to
move about within a broad floodplain, or free to move about, but within a relatively narrow
valley floor. Then examine the channel to ascertain the bank and valley features that constrain
the stream. Entry choices for the type of constraining features are bedrock, hillslopes,
terraces/alluvial fans, and human land use (e.g., a road, a dike, landfill, rip-rap, etc.). Estimate
the percent of the channel margin in contact with constraining features (for unconstrained
channels, this is 0%). To aid in this estimate, you may wish to refer to the individual transect
assessments of incision and constraint. Finally, estimate the "typical" bankfull channel width and
estimate the average width of the valley floor either with a topographic map or visually. If you
cannot directly estimate the valley width (e.g., it is further than you can see, or if your view is
blocked by vegetation), record the distance you can see and mark the appropriate bubble on the
field form.
Table 8.15 Procedure: Assessing Channel Constraint
Channel Constraint
NOTE: These activities are conducted after completing the thalweg profile and littoral-riparian
measurements and observations, and represent an evaluation of the entire stream reach.
Channel Constraint: Determine the degree, extent, and type of channel constraint based on envisioning the
stream at bankfull flow.
1. Classify the stream reach channel pattern as predominantly a single channel, an anastomosing
channel, or a braided channel.
• Single channels may have occasional in-channel bars or islands with side channels, but feature a
predominant single channel, or a dominant main channel with a subordinate side channel.
• Anastomosing channels have relatively long major and minor channels branching and rejoining
in a complex network separated by vegetated islands, with no obvious dominant channel.
• Braided channels also have multiple branching and rejoining channels, separated by
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Channel Constraint
unvegetated bars. Subchannels are generally small, short, and numerous, often with no obvious
dominant channel.
2. After classifying the channel pattern, determine whether the channel is constrained within a narrow
valley, constrained by local features within a broad valley, unconstrained and free to move about
within a broad floodplain, or free to move about, but within a relatively narrow valley floor.
3. Then examine the channel to ascertain the bank and valley features that constrain the stream. Entry
choices for the type of constraining features are bedrock, hillslopes, terraces/alluvial fans, and
human land use (e.g., a road, a dike, landfill, rip-rap, etc.).
4. Based on your determinations from Steps 1 through 3, select and record one of the constraint
classes shown on the Channel Constraint Form.
5. Estimate the percent of the channel margin in contact with constraining features (for unconstrained
channels, this is 0%). Record this value on the Channel Constraint Form.
6. Finally, estimate the "typical" bankfull channel width, and visually estimate the average width of the
valley floor. Record these values on the Channel Constraint Form.
NOTE: To aid in this estimate, you may wish to refer to the individual transect assessments of incision and
constraint that were recorded on the Channel/Riparian Cross-Section Forms.
NOTE: If the valley is wider than you can directly estimate, record the distance you can see and mark the
bubble on the field form.
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
<
N
ce
<
en
<
I
u
CD
<
<
y
00
>
Site ID:
NRSA 2018/19 CHANNEL CONSTRAINT
Date: / /
Re Revved by (initial): _
CHANNEL PATTERN (Fill in one):
O One Channel
O Anastomosing (complex) channel - (Relatively long major and minor channels branching and rejoining.)
O Braided channel - (Multiple short channels branching and rejoining - mainly one channel broken up by
numerous mid-channel bars.)
CHANNEL CONSTRAINT(Fill in one):
O Channel very constrained in V-shaped valley (i.e. it is very unlikely to spread out over valley or erode a
new channel during flood)
O Channel is in Broad Valley but channel movement by erosion during floods is constrained by Incision
(Flood flows do not commonly spread over valley floor or into multiple channels.)
O Channel is in Narrow Valley but is not very constrained, but limited in movement by relatively narrow
val ley flo or (< ~ 10 x b ankfull wi dth)
O Channel is Unconstrained in Broad Valley (i.e. during flood it can fill off-channel areas and side channels,
spread out over flood plain, or easily cut new channels by erosion)
O Bedrock (i.e. channel is a bedrock-dominated gorge)
O Hillslope (i.e. channel constrained in narrow V-shaped valley)
O Terrace (i.e. channel is constrained by its own incision into river/stream gravel/soil deposits)
O Human Bank Alterations (i.e. constrained by rip-rap, landfill, dike, road, etc.)
O No constraining features
Percent of channel length with margin
in contact with constraining feature:
%
(0-100%)
Bankfull width:
(m)
Valley width (Visual Estimated Average):
Jm)
Note: Be sure to include distances between both sides of valley border for valley width.
If you cannot seethe valley borders, record the distance
you can see and fill this bubble: ^
Percent of Channel Margin Examples
Y*
COMMENTS
03G8/2018 NRSA18 Channel Constraint
7734623125
Figure 8.17 Channel Constraint Form
94
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
A) Anastomosing channel pattern
FLOW
Vegetated islands above bankfull flow. Multiple
channels remain during major flood events.
B) Braided channel pattern
FLOW
Unvegetated bars below bankfull flow. Multiple
channel pattern disappears during major flood events.
DVP
Figure 8.18 Types of Multiple Channel Patterns
8.13.2 Debris Torrents and Recent Major Floods
Debris torrents, or tahars, 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,
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
including its substrate, banks, and riparian corridor, checking the presence of features described
on the Torrent Evidence Assessment Form (Figure 8.19). 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.
-------
National Rivers and Streams Assessment 2018/19 Field Operations Manual
Version 1.1 June 2018 Wadeable
Site ID:
NRSA 2018/19 TORRENT EVIDENCE ASSESSMENT
Date: / /
Reviewed by (initial):_
Please fill In any of the following that are evident.
EVIDENCE OF TORRENT SCOURING:
01 - Stream channel has a recently devegetated corridor two or more times the width of the low flow channel. This
corridor lacks riparian vegetation with possible exception of fireweed, even-aged alder or cottonwood seedlings, grasses,
or other herbaceous plants.
02 - Stream substrate cobbles or large gravel particles are NOT IMBRICATED. (Imbricated means that they lie with flat
sides horizontal and that they are stacked like roof shingles - imagine the upstream direction as the top of the "roof.") In
a torrent scour or deposition channel, the stones are laying in unorganized patterns, lying "every which way." In addition
many of the substrate particles are angular (not "water-worn.")
03 - Channel has little evidence of pool-riffle structure. (For example, could you ride a mountain bike down the channel?)
04 - The stream channel is scoured down to bedrock for substantial portion of reach.
05 - There are gravel or cobble berms (little levees) above bankfull level.
06 - Downstream of the scoured reach (possibly several miles), there are massive deposits of sediment, logs, and other
debris.
07 - Riparian trees have fresh bark scars at many points along the stream at seemingly unbelievable heights above the
channel bed.
OS - Riparian trees have fallen into the channel as a result of scouring near their roots.
EVIDENCE OF TORRENT DEPOSITS:
o
09 - There are massive deposits of sediment, iogs: and other debris in the reach. They may contain wood and boulders
that, in your judgement, could not have been moved by the stream at even extreme flood stage.
10 - If the stream has begun to erode newly laid deposits, it is evident that these deposits are "MATRIX SUPPORTED."
This means that the large particles, like boulders and cobbles, are often not touching each other, but have silt. sand, and
other fine particles between them (their weight is supported by these fine particles -- in contrast to a normal stream
deposit, where fines, if present, normally "fill-in" the interstices between coarser particles.)
NO EVIDENCE:
11 - No evidence of torrent scouring or torrent deposits.
COMMENTS
10/04/2017 NRSA18 Torrent Evidence
5543621376
<
M
<
cc
<
x
u
Figure 8.19 Torrent Evidence Assessment Form
CQ
<
<
y
LO
>
97
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.14 Stream Discharge
No single method for measuring discharge is applicable to all types of stream channels. The
preferred procedure for obtaining discharge data is based on "velocity-area" methods (e.g.,
Rantz et al., 1982; Linsley et al., 1982). For streams that are too small or too shallow to use the
equipment required for the velocity-area procedure, two alternative procedures are presented.
One procedure is based on timing the filling of a volume of water in a calibrated bucket. The
second procedure is based on timing the movement of a neutrally buoyant object (e.g., an
orange or a small rubber ball) through a measured length of the channel, after measuring one or
more cross-sectional depth profiles within that length.
8.14.1 Velocity-Area Procedure
Because velocity and depth typically vary greatly across a stream, accuracy in field
measurements is achieved by measuring the mean velocity and cross-sectional area of many
increments across a channel (Figure 8.20). Each increment gives a subtotal of the stream
discharge, and the whole is calculated as the sum of these parts. Discharge measurements are
made at only one carefully chosen channel cross-section within the sampling reach and it does
not have to occur at one of the 11 transects. It is important to choose a channel cross-section
that is as much like a canal as possible. A glide area with a "U" shaped channel cross-section that
is free of obstructions provides the best conditions for measuring discharge by the velocity-area
method. You may remove rocks and other obstructions to improve the cross-section before any
measurements are made. However, because removing obstacles from one part of a cross-
section affects adjacent water velocities, you must not change the cross-section once you
commence collecting the set of velocity and depth measurements.
The procedure for obtaining depth and velocity measurements is outlined in Table 8.16. Record
the data from each measurement on the Stream Discharge Form as shown in Figure 8.21. In the
field, data will be recorded using only one of the available procedures.
WATER SURFACE
15 to 20 equally spaced
intervals across stream.
beginning at left margin
Measure stream depth at the midpoint
of each interval, and obtain velocity
measurements at 0.6 depth
Extended surveyor's
rod or tape measure
Record distance
and depth of
right margin
Figure 8.20 Layout of Cross-Section for Obtaining Discharge Data by the Velocity-Area Procedure
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 8.16 Procedure: Determining Stream Discharge - Velocity-Area
Stream Discharge Velocity- Area Option
1. Locate a cross-section of the stream channel for discharge determination that has most of the
following qualities (based on Rantz and others, 1982):
• Segment of stream above and below cross-section is straight
• Depths mostly greater than 15 centimeters, and velocities mostly greater than 0.15
meters/second. Do not measure discharge in a pool.
• "U" shaped, with a uniform streambed free of large boulders, woody debris or brush, and dense
aquatic vegetation.
• Flow is relatively uniform, with no eddies, backwaters, or excessive turbulence.
2. Lay the surveyor's rod (or stretch a measuring tape) across the stream perpendicular to its flow, with
the "zero" end of the rod or tape on the left bank, as viewed when looking downstream. Leave the
tape tightly suspended across the stream, approximately one foot above water level.
3. Attach the velocity meter probe to the calibrated wading rod. Check to ensure the meter is functioning
properly and the correct calibration value is displayed. Calibrate (or check the calibration) the velocity
meter and probe as directed in the meter's operating manual. Darken in the "VELOCITY AREA" bubble
on the Stream Discharge Form.
4. Divide the total wetted stream width into 15 to 20 equal sized intervals. To determine interval width,
divide the width by 20 and round up to a convenient number. Intervals should not be less than 10 cm
wide, even if this results in less than 15 intervals. The first interval is located at the left margin of the
stream (left when looking downstream), and the last interval is located at the right margin of the
stream (right when looking downstream).
5. Stand downstream of the rod or tape and to the side of the first interval point (closest to the left bank
if looking downstream).
6. Place the wading rod in the stream at the interval point and adjust the probe or propeller so that it is
at the water surface. Darken in the appropriate bubbles for "Distance Units" and "Depth Units" on the
Stream Discharge Form. Record the distance from the left bank and the depth indicated on the wading
rod on the Stream Discharge Form.
Note: For the first interval, distance equals 0 cm, and in many cases depth may also equal 0 cm. For
the last interval, distance will equal the wetted width (in cm) and depth may again equal 0 cm.
7. Stand downstream of the probe or propeller to avoid disrupting the stream flow. Adjust the position
of the probe on the wading rod so it is at 0.6 of the measured depth below the surface of the water
(e.g., if the depth of the station was 1 meter, the probe would be placed 0.6 meters under water or
0.4 meters from the bottom). Face the probe upstream at a right angle to the cross-section, even if
local flow eddies hit at oblique angles to the cross-section.
8. Wait 20 seconds to allow the meter to equilibrate, then measure the velocity. Darken the appropriate
"Velocity Units" bubble on the Stream Discharge Form. Record the value on the Stream Discharge
Form. Note for the first interval, velocity may equal 0 because depth will equal 0.
• For the electromagnetic current meter (e.g., Marsh-McBirney), use the lowest time constant scale
setting on the meter that provides stable readings.
• For the impeller type meter (e.g., Swoffer 2100), set the control knob at the mid-position of
"DISPLAY AVERAGING". Press "RESET" then "START" and proceed with the measurements.
9. Move to the next interval point and repeat Steps 6 through 8. Continue until depth and velocity
measurements have been recorded for all intervals. Note for the last interval (right margin), depth and
velocity values may equal 0.
At the last interval (right margin), record a "Z" flag on the field form to denote the last interval
sampled.
If using a meter that computes discharge directly, darken the "Q" bubble on the discharge form, and
record calculated discharge value. In this case, you do not have to record the depth and velocity data
for each interval.
<
M
QC
<
QC
<
I
u
CO
<
<
y
i/i
>
99
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Site ID:
NRSA 2018/19 DISCHARGE - WADEABLE ONLY
Date: /
Revi ev«d by (i niti al): _
OQ Value
If discharge is determined directly
in field, record value here: Q =
O Velocity Area
Distance Units
O ft O cm
Depth Units
Oft Ocm
Velocity Units
Oft'sXX.X Om/sX.XX
Dist from Bank Depth Velocity Flag
1
0
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
O Timed Filling
Repeat
Volume (L)
Time (s)
Flag
O Neutral Buoyant Object
Measure the stream depth using a wading rod or meter stick at
points approximately equal to the following proportions of the total
width: 0.1,0.3, 0.5, 0.7, and 0.9.
Float Dist.
Oft O m
Float Time
(s)
Flag
Float 1
Float 2
Float 3
Cross Sections on Float Reach
Width
Oft O m
Depth 1
Oft O cm
Depth 2
Depth 3
D epth 4
D epth 5
Upper Section Middle Section Lower Section
O cfs
_0
Flag
Flag
Comments
Flag Codes: Z = Last station measured (if not Station 20); F1, F2, etc. = Miscellaneous flags assigned by each field crew.
Explain all flags in comments section.
03/28/2018 NRSA18 Discharge
5312491487
Figure 8.21 Discharge Form
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
8.14.2 Timed Filling Procedure
In channels too "small" for the velocity-area method, discharge can sometimes be measured by
filling a container of known volume and timing the duration to fill the container from natural
spillways or plunges. "Small" channels in this context, are too shallow for the current velocity
probe to be placed in a cross-section of the stream or where the channel is broken up and
irregular due to rocks and debris, and a suitable cross-section for using the velocity area
procedure is not available. The timed filling procedure can be an extremely precise and accurate
method, but requires a natural or constructed spillway of freefalling water. Locate one or more
natural spillways or plunges that collectively include the entire stream flow. A temporary
spillway can also be constructed using a portable V-notch weir, plastic sheeting, or other
materials that are available onsite. If obtaining data by this procedure will result in a lot of
channel disturbance or stir up a lot of sediment, wait until after all biological and chemical
measurements and sampling activities have been completed.
Choose a location within the sampling reach that is narrow and easy to block when using a
portable weir. Position the weir in the channel so that the entire flow of the stream is
completely rerouted through its notch (Figure 8.22). Impound the flow with the weir, making
sure that water is not flowing beneath or around the side of the weir. Use mud or stones and
plastic sheeting to get a good waterproof seal. The notch must be high enough to create a small
spillway as water flows over its sharp crest.
Water Level
Bucket
Weir Crest
Impounded Pool
Weir Crest
<
M
QC
Figure 8.22 Use of a Portable Weir and Calibrated Bucket to Obtain an Estimate of Stream Discharge
<
QC
<
I
u
The timed filling procedure is presented in Table 8.17. Make sure that the entire flow of the
spillway is going into the bucket. Record the time it takes to fill a measured volume on the
Discharge Measurement Form as shown in Figure 8.21. Repeat the procedure 5 times. If the
cross-section contains multiple spillways, you will need to do separate determinations of
discharge for each spillway. Clearly indicate which time and volume data replicates should be
CO
<
<
y
i/i
>
101
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
used for each discharge estimate contribution to the averaged for each spillway. When multiple
spillways are used, the total discharge is the sum of the separate averages for each spillway. Use
additional Stream Discharge Form(s) if necessary.
Table 8.17 Procedure: Determining Stream Discharge - Timed Filling
NOT E: If measuring discharge by this procedure will result in significant channel disturbance or will stir up
sediment, delay determining discharge until all biological and chemical measurement and sampling
activities have been completed.
1. Choose a cross-section that contains one or more natural spillways or plunges, or construct a
temporary one using on-site materials, or install a portable weir using a plastic sheet and on-site
materials.
2. Darken the 'TIMED FILLING" bubble in the stream discharge section of the Stream Discharge Form.
3. Position a calibrated bucket or other container beneath the spillway to capture the entire flow. Use a
stopwatch to determine the time required to collect a known volume of water. Record the volume
collected (in liters) and the time required (in seconds) on the Stream Discharge Form.
4. Repeat Step 3 a total of 5 times for each spillway that occurs in the cross-section. If there is more
than one spillway in a cross-section, you must use the timed filling approach on all of them.
Additional spillways may require additional data forms.
8.14.3 Neutrally Buoyant Object Procedure
In very small, shallow streams with no waterfalls, where the standard velocity-area or timed
filling methods cannot be applied; the neutrally buoyant object method may be the only way to
obtain an estimate of discharge. The required pieces of information are the mean flow velocity
in the channel and the cross-sectional area of the flow. The mean velocity is estimated by
measuring the time it takes for a neutrally buoyant object to flow through a measured length of
the channel. The channel cross-sectional area is determined from a series of depth
measurements along one or more channel cross-sections. Since the discharge is the product of
mean velocity and channel cross-sectional area, this method is conceptually very similar to the
standard velocity-area method.
The neutrally buoyant object procedure is described in Table 8.18. Examples of suitable objects
include plastic golf balls (with holes), small sponge rubber balls, or small sticks. The object must
float, but very low in the water. It should also be small enough that it does not "run aground" or
drag bottom. Choose a stream segment that is roughly uniform in cross-section, and that is long
enough to require 10 to 30 seconds for an object to float through it. Select one to three cross-
sections of this segment (depending on the variability of width and/or depth) to represent the
channel dimensions within the segment. Determine the stream depth at five equally-spaced
points at each cross-section. Three separate times, measure the time required for the object to
pass through the segment that includes all of the selected cross-sections. Record data on the
Stream Discharge Form as shown in Figure 8.21.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 8.18 Procedure: Determining Stream Discharge - Neutrally Buoyant Object
Stream Discharge Neutrally Buoyant Object Option
1. Darken the "NEUTRALLY BUOYANT OBJECT" bubble on the Stream Discharge Form.
2. Select a segment of the sampling reach that is deep enough to float the object freely and long enough
that it will take between 10 and 30 seconds for the object to travel. Mark the units used and record
the length of the segment in the "FLOAT DIST." field of the Stream Discharge Form.
3. If the channel width and/or depth change substantially within the segment, measure widths and
depths at three cross-sections, one near the upstream end of the segment, a second near the middle
of the segment, and a third near the downstream end of the segment.
If there is little change in channel width and/or depth, obtain depths from a single "typical" cross-
section within the segment.
4. At each cross-section, measure the wetted width using a surveyor's rod or tape measure, and record
both the units and the measured width on the Stream Discharge Form. Measure the stream depth
using a wading rod or meter stick at points approximately equal to the following proportions of the
total width: 0.1,0.3, 0.5, 0.7, and 0.9. Record the units and the depth values (not the distances) on the
Stream Discharge Form.
5. Repeat Step 4 for the remaining cross-sections.
6. Use a stopwatch to determine the time required for the object to travel through the segment. Record
the time in the "FloatTime" field of the Stream Discharge Form.
7. Repeat Step 6 two more times. The float time may differ somewhat for the three trials.
8.15 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.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
9 FECAL INDICATOR {ENTEROCOCCI)
9.1 Summary of Method
Collect a fecal indicator sample at the last transect (Transect K) after all other sampling is
completed. Filters must be frozen within 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 (Figure 13.2).
9.2 Equipment and Supplies
Table 9.1 provides the equipment and supplies needed to collect the fecal indicator sample.
Record the sample data on the Sample Collection Form (Figure 4.2).
Table 9.1 Equipment and Supplies: Fecal Indicator Sampling (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
Pencils (for data forms)
9.3 Sampling Procedure
The procedure for collecting the fecal indicator sample is presented in Table 9.2.
Table 9.2 Procedure: Fecal Indicator (Enterococci) Sample Collection (Wadeable Sites)
Enterococci Sample
1. Put on sterile, nitrile gloves.
2. Select a sampling location at transect K that is approximately 1 m from the bank and approximately
0.3 m deep. Approach the sampling location slowly from downstream or downwind.
3. Lower the uncapped, inverted 250 ml sample bottle to a depth of 1 foot (0.3 m) below the water
surface, avoiding surface scum, vegetation, and substrates.
4. Point the mouth of the container away from the body or boat. Right the bottle and raise it through
the water column, allowing bottle to fill completely.
5. If the depth does not reach 0.3 m along the transect at 1 m from the bank, take the sample and flag
it on the field form.
6. After removing the container from the water, discard a small portion of the sample to allow for
proper mixing before filtering (down to the 250 mL mark on the bottle).
7. Add the sodium thiosulfate tablet, cap, and shake bottle 25 times.
Storage
8. Store the sample in a cooler on ice to chill (do not freeze immediately). Chill for at least 15 minutes.
9. Sample must be filtered and all filters frozen within six hours of collection
104
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
9.4 Sample Processing in the Field
You will need to process two separate filters for the Enterococci sample. All the filters required
for an individual site should be sealed in plastic bags until use to avoid external sources of
contamination. Please refer to Section 13.3 for information regarding processing the
Enterococci samples.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
10 FISH ASSEMBLAGE
10.1 Summary of Method
The fish sampling method is designed to provide a representative sample of the fish community,
collecting all but the rarest fish taxa inhabiting the site. It is intended to accurately represent
species richness, species guilds, relative abundance, size, and presence of anomalies. The
intended uses of the fish assemblage data are to calculate predictive models of multimetric
indicators (MMIs; similar to an Index of Biotic Integrity (IBI); Pont et al. 2008, USEPA 2013a) and
possibly Observed/Expected (O/E) taxa richness. In addition, the fish assemblage data provides a
starting point for developing potential indicators of ecosystem services related to fish.
For wadeable streams, collect fish using a bank or towed (e.g., a towed barge, small watercraft,
or float tube) electrofishing unit (2,500-to 5,000-V; 2.5-5.0 Generator Powered Pulsator [GPP]
unit or equivalent) or backpack electrofisher. Use a backpack electrofishing unit in smaller
streams, when conductivity is appropriate, or in larger streams that are inaccessible with a
towed unit. As a last option, use seining as an alternate method when electrofishing is precluded
by high or low conductivity or extreme levels of turbidity.
There are different protocols for collecting fish from wadeable streams of different sizes (see
Section 10.3). For all wadeable sites, conduct sampling in an upstream direction (i.e., from the
downstream end of the reach), allocating effort (button time) within subreaches (areas between
the cross-section transects). At smaller streams (mean channel width [CW] rounded to the
nearest meter and recorded on the stream verification form is less than or equal to 12 m),
sample all available habitats over the entire sampling reach (40 CW; see Section 3.2). At large
wadeable streams (mean channel width is greater than or equal to 13 m), the initial length of
the fish sampling reach is less than the entire sampling reach, and effort is focused on habitats
along the stream margins. At large wadeable streams, the minimum sample reach is 20 CW (five
subreaches). If you have not collected 500 individuals at the end of the 20 CW, sample
additional subreaches in their entirety until you obtain at least 500 individuals, or until you have
sampled the entire sampling reach (40 CW; 10 subreaches). Record information related to
sampling effort on the front of the Fish Gear and Sample Information Form (Figure 10.1). Record
species identification and enumeration data on one or more pages of the Fish Collection Form
(Figure 10.2).
There are several revisions and clarifications to the wadeable sampling protocols from those
used in the NRSA 2013/14 effort. There is now a place on the fish gear form for the crew to note
the sufficiency of the sampling effort and the general response offish to electrofishing. The
primary and secondary electrofishing sections of the fish gear form now allow for users of
certain types of electrofishing systems to record voltage as High or Low, and to record the
percent of power. There is now a place on the collection form for a crew to note that they did
not evaluate the native/introduced status of any fish collected. This clarifies that blank values
for the introduced fish field on the collection form are truly missing, rather than being presumed
to be native.
-------
crq
c
n>
i—4
o
O
n>
QJ
Q.
in
0)
3
T3
3
fU
o
3
Site ID:
NRSA 2018/19 FISH GEAR AND SAMPLING INFORMATION (Front)
Date: I
Re\»ev\ed by (initial): «
I
FISH SAMPLING PROTOCOL (select one):
FISH SAMPLING - NOT CONDUCTED OR SUSPENDED (select one):
O Large Nonwadeable - (> 13m wide)
O Small Nonwadeable - (< 12m wide)
O Large Wadeable - (> 13m wide)
O Small Wadeable - (<12m wide)
O Fished - None Collected
O Not Fished - No Permit
O Fast flowing high gradient site
O Not Fished/Fishing suspended - Can't sample > 50% of ° Not Fished" Site Conditions Prohibit Sampling (Describe in comments)
required reach: O Fishing Suspended - Permit Restriction (Listed species encountered)
- Nonwadeable and Lg. Wadeable (at least 10 CW) O Not Fished - Equipment Failure
- Wadeable (at least 20 CW) O Not Fished - No fish observed (applies to small wadeable streams only)
Did conditions allow for OY ON
sufficient sampling? O Not Sure
Fish Response to q Immobilized O Inhibited Swimming O Escape
Electrofishing
Final Length of Fishing Reach (m):
Sampling Protocol Comments:
FISH GEAR INFORMATION
Water Visibility: OGood O Poor Water Temp (°C):
Specific Conductivity (uS/cm):
Corrected to 25" C ? OY ON
Primary Electrofishing Gear Model
O BOAT (Motor)
O RAFT (No motor)
O BACKPACK
O BANK OR TOWED UNIT Diameter
# of Netters (1):
Anodes
Number:
Wave Form: O AC O DC
O Pulsed DC
Volts: (50-1000)
Watts: likely 400 (bp),
2500 or 5000 (boatf raft) t
O in- % Power
. O CITI (Smith Root GPP only)
Pulse Rate:
pps or Hz (
Amps: (may not be
provided for bp) .
OR O High OLow Pulse Width (ms):
Total Shock (button) Time (s):
Total Fishing Time (min):
Length Sampled (m):
Secondary Electrofishing Gear Model:
O BOAT (Motor)
O RAFT (No motor)
O BACKPACK
O BANK OR TOWED UNIT Diameter
# of Netters (1):
Anodes
Number:
Wave Form: O AC O DC
O Pulsed DC
Volts: (50-1000)
Watts: likely 400 (bp),
2500 or 5000 (boatfraft) t
OR O High OLow Pulse Width (ms):
Total Shock (button) Time (s):
O in. % of Power
O CITI (Smith Root GPP only)
Pulse Rate:
pps or Hz ,
Amps: (may not be
provided for bp) ¦
Total Fishing Time (min):
Length Sampled (m):
Primary Seine Net: O BAG SEINE OMINNOWSEINE No. of crew members:
Height (m): Length (m): Mesh (mm):
Avg. Haul
Lenqth (m):.
No. of
Hauls:
Total Seining
Time (min):
Length Sampled (m):
Secondary Seine Net: O BAG SEINE O MINNOW SEINE No. of crew members:
Height (m): Length (m): Mesh (mm):
Avg. Haul
Length (m): _
No. of
Hauls:
Total Seining
Time (min): ,
Length Sampled (m):
GEAR INFORMATION COMMENTS
03/28/2018 NRSA18 Fish Gear and Sampling Information
0788063181
< 2
O) QJ
-! c+
~ o"
o -J
c
D
ro
NJ
o
<
O)
CD
3
Q.
t/1
n>
QJ
3
3
n>
D
c+
NJ
O
31
0)'
Q_
O
¦a
fD
-1
QJ
o'
s
3
CO
QJ
Q.
2
n>
QJ
SD
3
cr
C
fD
QJ
O FISH ASSEMBLAGE
-------
O FISH ASSEMBLAGE
00
orq
c
-I
n>
i—4
o
fsj
n
o
n>
n
o
3
¦
NRSA 2018/19 FISH COLLE
Site ID: Date: / /
CTI
DN
Page of
Reviewed bv IkiWill:
1 did not evaluate the native/
introduced status for fish collection O Y
¦
Line
#
Common Name
. -D
>» 'C
X JS
£ -o
Tally and Counts
Anom
Count
Mortality
Count
VOUCHER
Voucher
Tag#
Vouchers
Retained
Flag
<150 mm
150-300 mm
300-460 mm
>460 mm
Unk/Rng
QA
Photo
1
O
o
0
0
o
2
O
o
o
o
o
3
o
o
o
o
o
4
o
o
o
o
o
5
o
o
0
0
o
6
o
o
0
0
o
7
o
o
o
o
o
8
o
o
0
0
o
9
o
o
o
o
o
10
o
o
o
o
o
11
o
o
o
0
o
12
o
o
o
o
o
13
o
o
0
0
o
14
o
o
o
o
o
Flag
Comments
Flag
Comments
Flag codes: K = No measurement made, F1.F2, etc. = flags assigned by each field crew. Explain all flags In comments.
A 2194249191 A
^ 09/12/2017 NRSA18 Fish Col lection •
< 2
CD QJ
-J r+
o °
O -j
L_ <
c 3>
D tn
HD
3
Q.
t/>
fD
Q)
3
3
fD
D
r+
NJ
O
31
fD'
Q_
o
¦a
fD
-1
QJ
r+
o'
§
3
to
QJ
Q.
2
ro
QJ
QJ
3
ZT
C
CD
Q)
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
10.2 Equipment and Supplies
Table 10.1 shows the checklist of equipment and supplies required to complete the fish
assessment. This checklist is similar to the one presented in Appendix A, which is used at the
base location to ensure that all of the required equipment is brought to the site.
Table 10.1 Equipment and Supplies: Fish Collection (Wadeable Sites)
For collecting
samples
Bank or towed electrofishing equipment
(e.g., towed barge or mounted on a small
watercraft or float tube), including
variable voltage pulsator unit, wiring
cables, generator, and electrodes
Backpack electrofishing unit with anode
and cathode
Linesman gloves, boots, and other
necessary safety equipment
Dip nets
Extra electrofishing unit batteries
Scientific collection permit(s)
Digital camera with extra memory card &
battery
1 Laser rangefinder (optional)
Polarized sunglasses and hats
10 ft x 6 ft Minnow or Bag Seine with %
inch mesh (additional 4' depth seine may
also be used)
1 Scalpel for slitting open large fish
before preservation.
1 container of 10% buffered formalin
Small mesh bags or several Leak proof
HDPE jars (various sizes from 250 mL- 4L)
for fish voucher specimens
2 non-conducting dip nets with % inch
mesh
1 Minnow net for dipping small fish from
live well
2 measuring boards (3 cm size classes)
1 set Fish ID keys
Field Operations Manual and/or
laminated Quick Reference Guide
20 ft x 6 ft Minnow or Bag Seine with %
inch mesh (additional 4 ft depth seine
may also be used)
Buckets
For recording
measu rements
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 and Information Form
Fish Collection forms (several per site,
depending on expected species richness)
Seining information form
Clear tape strips
Clear tape strips
Soft (#2) lead pencils
Fine-tip indelible markers
10.3 Sampling Procedures
The reach length sampled for fish varies based on the mean width of the stream used to
establish the length of the sampling reach (which has been rounded to the nearest meter as per
the directions in Section 3.2) and on the number of individuals collected (Figure 10.3). For small
wadeable streams (mean CW from the stream verification form <12 meters), follow the protocol ^
presented in Section 10.3.2. Sample the entire reach (150 m to 40 CW, 10 subreaches) and m
move the electrofishing unit (towed r or backpack) within each subreach to sample both ^
shorelines as well as the mid-channel. <%
<
109
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
For large wadeable streams (mean CW from the stream verification form > 13 m), follow the
protocols described in Section 10.3.3. For large wadeable streams the minimum length for fish
sampling is 20 CW (5 of the 10 subreaches). If a minimum of 500 fish are not collected after
sampling this minimum fishing reach, sample additional subreaches in their entirety until at
least 500 fish are collected, or all 10 subreaches have been sampled. Stop sampling when you
reach Transect K (the end of the sampling reach), regardless of the number of individuals
collected. Table 10.2 summarizes the fishing protocols for small and large wadeable streams.
If conditions prohibit any type of electrofishing, collect fish by seining as described in Table 10.3.
The objective of seining is to collect species and relative abundance data that is comparable to
what would have been obtained by electrofishing at the site. If seining is used, record all fish
collected with seining protocols on separate lines of the fish collection form.
It is important that you record the total reach length that was sampled for fish, as this is used
along with the number of fish collected to determine sampling sufficiency. Data from streams
that were not sufficiently sampled for fish cannot be used to assess stream condition based on
the fish assemblage.
10.3.1 Irruptive Species
For the purposes of NRSA, the term irruptive species will be used to describe fish species which
are found in locally abundant "patches" in one or two small places within the sampling reach.
These are distinct from dominant species which are in abundance throughout most of the reach.
As such, irruptive species may artificially skew necessary effort to reach 500 individuals; and, if
included the overall assemblage counts, may artificially skew the calculations of relative
abundance offish species in the reach. To avoid the impact of irruptive species, move quickly
through large isolated schools of a single species (e.g., shad, certain shiners, etc.). Also, when
tallying total fish at the end of the designated fish sampling reach, calculate the percentage of
irruptive species to total individuals captured. If any single irruptive species comprises greater
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%.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 10.2 Summary of Wadeable Fishing Protocols
Small Wadeable (mean channel width [CW1 from stream verification form < 12 meters)
• Fish sampling reach length will be between 150 m (CW < 4 m) and 40 CW.
• Subreaches will be between 15 meters and 4 channel widths long
S Sample all 10 subreaches in their entirety from bank to bank starting at Transect A
S Total button time will range from 500-700 seconds per subreach
• You do not have to expend equal button time among the 10 subreaches—you can devote
more button time to subreaches with more complex habitat.
S No minimum fish number
Large Wadeable (mean channel width [CW1 from stream verification form > 13 meters)
• Initial minimum fish sampling reach will be 20 CW (5 subreaches).
• Subreaches will be 1/10 of the sampling reach in length
S Fish each subreach in a swath 8 meters from bank in pairs of subreaches starting at a
random bank at Transect A
S Button time is roughly 700 seconds per subreach
• Depending upon the habitat complexity, you can vary the distance actively fished to
allocate the available button time throughout the subreach. Put another way, sample for
700 seconds and if you have not reached the next transect, stop fishing and move to that
transect.
S Minimum fish number is 500 or until 10 subreaches have been fished.
> After fishing 5 subreaches, if 500 fish have not been collected, add subreaches one at a
time (but fish them in their entirety) until 500 fish are collected or all 10 subreaches
have been fished.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Small Wadeable Stream: Mean Channel Width < 12 m
40 x Channel Width
FLOW
Large Wadeable Stream: Mean Channel Width > 13 m
< 20 x Channel Width >
8 m
FLOW |
If < 500 individuals have been collected after fishing
20 CW (5 subreaches), continue fishing next subreach
(alternating bank after every two subreaches) until
either 500 individuals are collected, or Transect K is
reached (10 subreaches [40 CW] have been sampled)
Figure 10.3 Reach Layouts for Fish Sampling at Wadeable Sites
Dark shaded areas indicate the minimum length of the fish sampling reach. Light shaded areas are
sampled as needed to meet the required 500 individuals.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
10.3.2 Small Wadeable Streams
Table 10.3 describes the procedure for collecting fish in small wadeable streams. The sampling
crew should consist of one electrofisher operator, one dip-netter (1/4" mesh dip net), and an
optional bucket carrier (who may also have a net to aid in transferring fish to the livewell). An
anode with a net cannot substitute for a netter. For safety, all crew members are required to
wear non-breathable waders and insulated gloves. To aid vision, wear polarized sunglasses and
a hat or visor. See Appendix E for example starting settings for electrofishing using backpack,
towed barge, and boat (Temple 2018). These are only suggestions; the final determination of
settings is decided by the lead fish taxonomist.
Begin sampling at the downstream end of the sampling reach defined for the site (Figure 10.3)
and proceed upstream. Sample the entire reach, which will be between 150 m and 40 channel
widthslO subreaches). Total button time will vary between 500 and 700 seconds per subreach.
Conduct sampling by subreach (area between transects), but you do not have to allocate effort
equally among all 10 subreaches.
Whenever possible, process fish at the end of each subreach to minimize mortality and stress to
fish. You can use multiple lines per species on the Fish Collection Form if necessary (e.g., you
collect a large number of individuals and need additional space for tallying, or collect the same
species at multiple subreaches [e.g., A-B and G-H]).
Table 10.3 Procedure: Electrofishing (Small Wadeable Streams)
Electrofishing Procedures in Small Wadeable Streams
1) Complete the header section of the fish gear form (Site ID and date).
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, mark Not Fished- No Permit.
b) If site conditions prevent barge or backpack electrofishing (e.g., no access, safety concerns,
ambient conductivity is too high or too low to use a barge or backpack electrofishing unit),
determine if you can sample by seining.
i) If yes, follow the procedures presented in Table 10.5.
ii) If not, mark Not Fished - Site Conditions Prohibit Sampling. Note the conditions in the
Sampling Protocol Comments field.
c) If you can determine that > 50% of the required fish sampling reach 75 m or (20 CW; 5 subreaches)
cannot be sampled, mark Not Fished/Fishing Suspended - Can't sample >50% of required reach.
d) If you cannot sample because of equipment problems, mark Not Fished - Equipment Failure.
e) At a very small and very shallow stream, if you cannot attempt to sample, but are very confident
that no fish are present (i.e., you do not observe any at any point along the sampling reach), then
mark Not Fished - No Fish Observed.
f) If you cannot sample for any other reason, note the reason in the Sampling Protocol Comments
field.
3) If you can begin to sample, mark Wadeable in the Fish Sampling Protocol section.
a) Proceed to the downstream end of the sampling reach (Transect A).
b) For safety, everyone must wear personal floatation devices, non-breathable waders, foot
protection, and insulated linesman's gloves.
c) To aid vision while netting fish, wear polarized sunglasses and a hat or visor.
4) Mark the appropriate Water Visibility conditions on the form. Poor implies that your ability to
<
l/l
l/l
<
113
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
electrofish effectively is compromised because of poor visibility. Record the water temperature and
conductivity (note whether the conductivity value is corrected to 25 °C).
5)
Mark either Backpack, Bank or Towed Barge in the Primary Electrofishing Gear section of the fish gear
form. Mark Towed Barge for any electrofishing unit that is towed (e.g., kayak or float tube).
a)
Do not use any secondary electrofishing gear in a small wadeable stream.
6)
Operation of Bank or Towed Electrofisher
a)
Set unit to pulsed DC and mark it in the Wave Form section of the fish gear form.
b)
Select the initial voltage setting based on the conductivity of the stream.
i) See Tables in Appendix E.
ii) If the electrofishing system only lets you select High and Low voltage (rather than a specific
voltage), record the setting used on the fish gear form.
iii) If your conductivity meter cannot measure ambient conductivity, you can "uncorrect" specific
conductance at 25 °C to ambient conductivity using the following equation:
(1) Ambient conductivity=Specific conductance x (l+([water temp-25 °C] x 0.02)).
c)
Select the initial pulse rate and width.
i) In waters with strong swimming fish (length >200 mm), use a pulse rate of 30 Hz with a pulse
width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-120 Hz.
d)
If the electrofishing system only lets you adjust the percent of power, record the value on the fish
gear form.
e)
Turn the electrofisher on, set the timer, and depress the switch to begin fishing. If fishing success
is poor, increase the pulse width first and then the voltage. Increase the pulse rate last to
minimize mortality or injury to large fish. If mortalities occur, first decrease pulse rate, then
voltage, then pulse width.
f)
Once you have determined the appropriate settings, record them on the fish gear form. Start
cleared clocks and resume fishing.
g)
Note: Some electrofishers do not meter all the requested settings; provide what you can.
h)
If button time is not metered, estimate it with a stop watch and flag the data.
7)
Operation of Backpack Electrofisher
a)
Set unit to pulsed DC and mark it in the Wave Form section of the fish gear form.
b)
Select the initial voltage setting based on the conductivity of the stream.
i) See Tables in Appendix E.
ii) If your conductivity meter cannot measure ambient conductivity, you can "uncorrect" specific
conductance at 25 °C to ambient conductivity using the following equation:
(1) Ambient conductivity=Specific conductance x (l+([water temp-25 °C] x 0.02)).
c)
Select the initial pulse rate and width.
i) In waters with strong swimming fish (length >200 mm), use a pulse rate of 30 Hz with a pulse
width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-120 Hz.
d)
Turn the electrofisher on, set the timer, and depress the switch to begin fishing. If fishing success
is poor, increase the pulse width first and then the voltage. Increase the pulse rate last to
minimize mortality or injury to large fish. If mortalities occur, first decrease pulse rate, then
voltage, then pulse width.
e)
Once you have determined the appropriate settings, record them on the fish gear form. Start
cleared clocks and resume fishing.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
f) Note: some electrofishers do not meter all the requested settings; provide what you can.
g) If button time is not metered, estimate it with a stop watch and flag the data.
8) Once the settings on the electrofisher are adjusted properly to sample effectively and minimize injury
and mortality, begin sampling at the downstream end of the reach (Transect A) and fish in an
upstream direction.
a) The minimum reach length is 150 m. The maximum reach length for this protocol is 40 CW.
i) Search for fish even if the stream is extremely small, and it appears that sampling may
produce no specimens.
ii) Button time should range from 500 to 700 sec per subreach. Actual time will depend upon
the water conditions, the diversity and complexity of the available habitat, and on the
number offish present in the reach.
(1) If the electrofisher is highly effective and the fish are staying stunned longer and the
water is clear and flowing slowly then button time will be much lower than it will be in a
system where it is more turbid, flowing faster, and the fish are not being stunned as well.
(2) You do not have to expend equal button time among the 10 subreaches—you can devote
more button time to subreaches with more complex habitat and less time to subreaches
with simple habitat.
b) Depress the switch and slowly sweep the electrode from side to side.
c) Sample all habitat types (deep, shallow, fast, slow, complex, and simple). Avoid the temptation to
focus sampling only in the richest habitat types.
i) For available cut-bank and snag habitats, move the anode wand into cover with the current
off, turn the anode on when in the cover, and then remove the wand quickly to draw fish out.
ii) In fast, shallow water, sweep the anode and fish downstream into a net.
iii) In stretches with deep pools, fish the margins of the pool as much as possible, being
extremely careful not to step or slide into deep water.
d) Keep the cathode near the anode if fish catch is low.
9) The netter, holds the net 1 to 2 ft from the anode, follows the operator, nets stunned individuals, and
places them in a bucket.
a) Irruptive species: If you encounter a large school of a single species (e.g., shad, certain shiners,
etc.), quickly move through it to ensure you can sample the entire subreach within the allotted
button time.
10) Continue upstream until you reach the next transect (end of subreach).
a) Process fish and/or change water after each subreach to reduce mortality and stress.
i) Although not required, you may note amphibians and reptiles captured on the fish collection
form.
b) Release fish in a location that eliminates the likelihood of recapture.
11) Repeat Steps 8-10 until all 10 subreaches are sampled (i.e., you reach transect K).
12) After sampling all 10 subreaches, record the final length of the fish sampling reach in the Primary
Electrofishing Gear section of the fish gear form.
a) If you suspend sampling before completing all 10 subreaches, record the actual length that was
sampled, and mark the reason for the suspension in the Fish sampling - Not Conducted or
Suspended section of the fish gear form.
b) If you did not collect any fish, mark Fished - None Collected in the Fish sampling - Not Conducted
or Suspended section of the fish gear form.
13) In the Primary Electrofishing Gear section of the fish gear form, record the total button time expended
for electrofishing, the total time spent sampling, and the length of the total fish sampling reach
<
l/l
l/l
<
115
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
(recorded in Step 12) sampled by electrofishing.
14) Indicate whether conditions allowed for sufficient sampling on the fish gear form (Yes, No, Not Sure).
If you marked No or Not Sure, explain why in the Sampling Protocol Comments field.
15) Note the general response offish to your final electrofishing settings as either Immobilized, Inhibited
Swimming, or Escape on the fish gear form.
16) 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.
10.3.3 Large Wadeable Streams
Table 10.4 describes the procedure for collecting fish in large wadeable streams. The
electrofishing crew should consist of one electrofishing operator, and one dip netter and an
optional bucket carrier (who may also have a net to aid in transferring fish to the live well). An
anode with a net cannot substitute for a netter. For safety, all crew members are required to
wear non-breathable waders and insulated gloves. Polarized sunglasses and caps to aid vision
are also required. See Appendix E for example starting settings for electrofishing using
backpack, towed barge, and boat (Temple 2018). These are only suggestions; the final
determination of settings is decided by the lead fish taxonomist.
For large wadeable streams with a mean channel width (from the stream verification form) >13
m, the minimum fish sampling reach is 20 channel widths (5 subreaches). As shown in Figure
10.3, begin sampling at Transect A on a randomly determined bank and fish a section of the
subreach that extends approximately 8 m from the bank in an upstream direction. Within each
subreach, fish the near bank habitat as well as midstream habitat within the 8 meter sampling
area for a button time of ~700 seconds. When 700 seconds are reached, stop electrofishing
unless you are "pushing" a large school of fish, in which case continue fishing until you capture
them at a break. To reduce stress and mortality, net immobilized fish immediately and deposit
into a bucket or live-well for processing.
Whenever possible, process fish at the end of each subreach to minimize mortality and stress to
fish. 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.
At the end of the minimum fish sampling reach (20 CW or 5 subreaches), determine if you have
collected at least 500 individuals. If so, stop sampling. If not, sample additional subreaches (one
at a time) until at least 500 individuals are captured. If irruptive species make up > 50% of the
sample, sample one or more additional subreaches to bring the proportion of the irruptive
species below 50%. Stop sampling when you reach Transect K (the end of the entire 40 CW
sampling reach), regardless of the number of individuals collected. Once the decision is made to
fish an additional subreach, it should be completely fished as described above (do not stop
sampling partway through a subreach).
Table 10.4 Procedure: Electrofishing (Large Wadeable Sites)
Electrofishing Procedures in Large Wadeable Streams
1) Complete the header section of the fish gear form (Site ID and date).
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, mark Not Fished- No Permit.
b) If site conditions prevent towed or backpack electrofishing (e.g., no access, safety concerns,
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Electrofishing Procedures in Large Wadeable Streams
conductivity is too high or too low to use a barge or backpack electrofishing unit), determine if
you can sample by seining.
i) If yes, follow the procedures presented in Table 10.5.
ii) If not, mark Not Fished - Site Conditions Prohibit Sampling. Note the conditions in the
Sampling Protocol Comments.
c)
If you can determine that > 50% of the required fish sampling reach cannot be sampled (20 CW; 5
subreaches), mark Not Fished/Fishing Suspended - Can't sample >=50% of required reach.
d)
If you cannot sample because of equipment problems, mark Not Fished-Equipment Failure.
e)
If you cannot sample for any other reason, note the reason in the Sampling Protocol Comments
field.
3)
If you can begin to sample, mark Wadeable in the Fish Sampling Protocol section.
a)
Proceed to the downstream end of the reach (Transect A).
b)
For safety, everyone must wear personal floatation devices, non-breathable waders, foot
protection, and insulated linesman's gloves.
c)
To aid vision while netting fish, wear polarized sunglasses and a hat or visor.
4)
Mark the appropriate Water Visibility conditions on the form. Poor implies that your ability to
electrofish effectively is compromised because of poor visibility. Record the water temperature and
conductivity.
5)
Mark either Backpack or Bank or Towed Unit in the Primary Electrofishing Gear section of the fish gear
form.
a)
Do not use any secondary electrofishing gear in a large wadeable stream.
6)
Operation of Bank or Towed Electrofisher
a)
Set unit to pulsed DC and mark it in the Wave Form section of the fish gear form.
b)
Select the initial voltage setting based on the ambient conductivity of the river (i.e., not corrected
to 25 °C).
i) See Tables in Appendix E.
ii) If the electrofishing system only lets you select High and Low voltage (rather than a specific
voltage), record the setting used on the fish gear form.
c)
Select the initial pulse rate and width.
i) In waters with strong swimming fish (length >200 mm), use a pulse rate of 30 Hz with a pulse
width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-120 Hz.
d)
If the electrofishing system only lets you adjust the percent of power, record the value on the fish
gear form.
e)
Turn the electrofisher on, set the timer, and depress the switch to begin fishing. If fishing success
is poor, increase the pulse width first and then the voltage. Increase the pulse rate last to
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 flag the data.
7)
Operation of Backpack Electrofisher
a)
Set unit to pulsed DC and mark it in the Wave Form section of the fish gear form.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Electrofishing Procedures in Large Wadeable Streams
CD
<
i
CO
CO
CO
<
118
b) Select the initial voltage setting based on the ambient conductivity of the river,
i) See Tables in Appendix E.
c) Select the initial pulse rate and width.
i) In waters with strong swimming fish (length >200 mm), use a pulse rate of 30 Hz with a pulse
width of 2 m/sec.
ii) If you expect mostly small fish, use a pulse rate of 60-120 Hz.
d) Turn the electrofisher on, set the timer, and depress the switch to begin fishing. If fishing success
is poor, increase the pulse width first and then the voltage. Increase the pulse rate last to
minimize mortality or injury to large fish. If mortalities occur, first decrease pulse rate, then
voltage, then pulse width.
e) Once you have determined the appropriate settings, record them on the fish gear form. Start
cleared clocks and resume fishing.
f) Note: some electrofishers do not meter all the requested settings; provide what you can.
g) If button time is not metered, estimate it with a stop watch and flag the data.
8) Once the settings on the electrofisher are adjusted properly to sample effectively and minimize injury
and mortality, begin sampling at the downstream end of the reach (Transect A). Randomly choose a
bank on which to start and fish in an upstream direction within 8 m of the shoreline.
For the large stream protocol, the minimum initial fish sampling reach length is 5 subreaches
9) When using a towed electrofishing unit, the minimum crew size for electrofishing is three.
a) The operator must remain actively at the control box and navigate the towed electrofishing unit.
b) The probe operator will use one probe.
10) When using a backpack electrofishing unit, the minimum crew size is two (one operator with the
probe and one netter). An anode outfitted with a net cannot substitute for a netter.
11) Depress the switch and slowly sweep the electrode from side to side.
a) Search for fish even if it appears that sampling may produce no specimens.
b) Sample all habitat types (deep, shallow, fast, slow, complex, and simple). Avoid the temptation to
focus sampling only in the richest habitat types.
i) In slack water area (e.g., available cut bank and snag habitats), move the anode wand into
cover with the current off, turn the anode on when in the cover, and then remove the wand
quickly to draw fish out.
ii) In fast, shallow water, sweep the anode and fish downstream into a net.
iii) In stretches with deep pools, fish the margins of the pool as much as possible, being
extremely careful not to step or slide into deep water.
c) Keep the cathode near the anode if fish catch is low.
12) The netter, holds the net 1 to 2 ft from the anode, follows the operator, nets stunned individuals, and
places them in a bucket.
a) Use a 6 mm (1/4 inch) mesh dip net to collect stunned fish. Actively capture stunned fish from
the electric field and immediately place them into the live well. Devote special attention to net
small and benthic fishes as well as fishes that may respond differently to the electric current.
b) Irruptive species: If you encounter a large school of a single species (e.g., shad, certain shiners,
etc.), quickly move through it to ensure you can sample the entire subreach within the allotted
button time.
13) Continue upstream until you reach the next transect (end of subreach).
a) The total button time within each subreach should be -700 sec. depending upon the habitat
complexity, you can vary the distance actively fished to allocate the available button time
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Electrofishing Procedures in Large Wadeable Streams
Field Operations Manual
Wadeable
throughout the subreach.
b) Process fish after each subreach to reduce mortality and stress. Release fish in a location that
eliminates the likelihood of recapture.
i) Although not required, you may note amphibians and reptiles captured on the fish collection
form.
14) Move to the opposite bank when necessary (see Figure 10.3). Repeat Steps 11-14 until you have
sampled the required length of stream.
a) If you have to suspend sampling before completing the required fish sampling reach, record the
actual length that was sampled, and mark the reason for the suspension in the Fish sampling -
Not Conducted or Suspended section of the fish gear form.
15) If the minimum reach length has been met (20 X mean stream width) determine the total number of
individuals collected.
a) If the total is < 500, sample one or more additional subreaches in their entirety until at least 500
individuals have been collected and processed, or you sample all 10 subreaches. Go to Step 16.
b) If you collect > 500 individuals, determine if a single irruptive species comprises > 50% of the
total number of individuals.
i) If an irruptive species makes up > 50% of the sample, sample one or more additional
subreaches to bring the proportion of the irruptive species below 50%. Go to Step 16.
ii) If not, go to Step 16.
c) If you have sampled all 10 subreaches (i.e., you have reached Transect K), go to Step 16.
16) After sampling, record the final length of the fish sampling reach in the Primary Electrofishing Gear
section of the fish gear form.
a) If you did not collect any fish, mark Fished - None Collected in the Fish sampling - Not Conducted
or Suspended section of the fish gear form.
17) In the Primary Electrofishing Gear section of the fish gear form, record the total button time expended
for electrofishing (should be -700 sec per subreach sampled), the total time spent sampling, and the
length of the total fish sampling reach (recorded in Step 15 or 16) sampled by electrofishing.
18) Indicate whether conditions allowed for a sufficient sampling effort on the fish gear form (Yes, No, Not
Sure). If you marked either No or Not Sure, explain why in the Sampling Protocol Comments field.
19) Note the general response behavior of fish to your final electrofishing settings on the fish gear form
(source: https://nctc.fws.gOv/courses/CSP/CSP2C01/resources/l EF Effects (Chapter 8).ppt) 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).
20) 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.
10.4 Seining <
CO
In small or large wadeable streams where conditions prohibit electrofishing, use seining only as ^
the last option for collecting fish. Seining is not to be used in concert with electrofishing. If ^
conditions are such that seining is the only method used, provide a justification in the Sampling <
Protocol Comments section of the fish gear form (Figure 10.1). Table 10.5 presents the
119
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
procedure for seining wadeable streams. The intent of the seining effort is to provide
comparable data to electrofishing.
Although electrofishing typically works best in an upstream direction, seining may work best
moving downstream. Allocate seine hauls so that the snag, edge, and mid-channel habitats are
fished thoroughly. In general, edge and snag habitats will be sampled using narrower seines
over shorter distances, while mid-channel habitats will be sampled using longer seines over
longer distances. Generalized habitat seining procedures are presented in Table 10.5.
Depending upon habitat types and complexity, use 2 to 3 crew members. Two crewmembers
move the seine; an optional third person creates and maintains a bag in the seine in area with
higher velocities, or agitates rocks in riffles or snags.
To avoid mortality, process fish after each seine haul. If necessary, you can use additional lines
on the fish collection form (Figure 10.2) to record species collected with different nets or in
different hauls. Record all fish collected with seining protocols on separate lines on the field
collection form from those lines used for fish collected by electrofishing.
If you seine, record information for each seine haul on the Seining Information Form to track
effort (Figure 10.4). Denote the bank as right or left as you face downstream. Restrict each haul
to a single habitat type. After fish sampling is completed for the site, use the information from
the seining information form to complete the information in the Primary Seine Net section of
the fish gear form (Figure 10.1).
Table 10.5 Procedure: Seining (Wadeable Sites)
Procedures for Seining at All Wadeable Sites
CD
<
i
CO
CO
CO
<
120
1) Use seining as a last option only (e.g., when electrofishing is ineffective due to high conductivity or
extremely high turbidity). Do not use seining as a supplementary method to electrofishing. Fish
sampling reach must be safely wadeable.
a) If site conditions are such that only seining is used, note the reason in the Sampling Protocol
Comments section.
b) At the end of each seine haul, immediately place all fish in one or more live wells to minimize
injury and mortality, and so that most fish can be returned to the river alive.
2) Complete the header section of the fish gear form (Site ID and date).
3) Mark the pertinent protocol and size class in the Fish Sampling Protocol section.
a) Proceed to the downstream end of the reach (Transect A).
i) At some sites, seining may be more effective while working downstream (from Transect K)
instead of upstream.
(1) If working downstream in a large wadeable stream, reverse the transects in Figure 10.3
and move to the opposite bank where indicated.
b) For safety, everyone must wear personal floatation devices and foot protection.
c) To aid vision while seining, wear polarized sunglasses and a hat or visor.
4) Mark the appropriate Water Visibility conditions on the form. Poor implies that your ability to seine
effectively is compromised because of poor visibility. Record the water temperature and conductivity.
Note whether the conductivity value is corrected to 25 °C.
5) Mark the type of seine being used (Bag Seine or Minnow Seine) in the Primary Seine Net section of the
fish gear form. This is the seine that will be used for sampling the majority of the fish sampling reach.
a) Record the number of crewmembers (2-3), and the net dimensions (height, total length, and
mesh size) on the fish gear form.
b) If you have to use a second type of seine for parts of the sampling reach, Mark the type and
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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 10.2).
7) To maximize capture efficiency, please do the following:
a) Always use 10 and 20 ft. seines. When necessary, reduce the width by rolling seine poles and
floats into the net.
b) When narrowing seines, always keep lead line outside of the pole.
c) When working edge habitats, only roll the inner side of the seine, while keeping the near bank
pole extended.
d) As a default, use seines that are 2 meters in depth. A 1.25 meter seine may be used in shallow
habitats.
e) Keep the float line above the surface (avoid dragging it below the surface while pulling).
f) Maintain the lead line along the river bottom.
g) Either tie the seine to the poles tightly, or roll the seine into the poles.
h) Always maintain the bag behind the poles.
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 of fish (in terms of species
richness and relative abundance, and allocation of effort throughout the fish sampling reach) to
that obtained if the site was electrofished.
i) Avoid extended seine hauls that collect hundreds of individuals.
ii) Seine as many available habitat types as possible within each subreach (one haul each).
iii) Total time spent seining a site should be comparable to what would have been spent
electrofishing.
b) Riffle Habitats
i) Use two crewmembers, each tending a seine pole. Place the seine perpendicular to the
current across the downstream end of the riffle. Ensure that the lead line is on the bottom.
Tilt the net slightly downstream to form a bag to trap aquatic vertebrates.
ii) Starting no more than 3 m upstream, a third crewmember 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.. Process fish (i.e.,
enumerate, identify and voucher fish) and record tally information on the fish collection form
(Figure 10.2). You may use separate lines on the fish collection form to record species
information from seine hauls.
c) Pool, Backwater, and Bar Habitats (Slack water)
i) Use two crewmembers, each tending a seine pole. Pull the seine across the pool using
shallow riffles or banks as barriers. A third crewmember creates and maintains a bag in the
seine.
ii) In areas with current, pull the net downstream and then sweep toward the bank with one or
both poles, or post one pole on the bank and sweep the other end in a wide arc from
midstream to the same bank.
iii) You can work pools in short to long hauls and use seines of varying width depending on the
complexity and depth of the pool. Keep the seine depth constant at 2 meters.
iv) Pull the bag completely to shore at a predesignated point.
d) Glide or Run Habitats (noticeable current)
<
l/l
l/l
<
121
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
<
I
CO
U~)
LD
<
122
i) Use two crewmembers, each tending a seine pole. Pull the seine diagonally across the glide
towards the bank. If necessary, a third crewmember creates and maintains a bag in the seine.
ii) Pull the net quickly downstream along the glide moving diagonally toward the bank. When
you reach the bank with the outer edge of the seine, post the pole and sweep the other end
in a wide arc from midstream to the same bank.
iii) Because of decreased complexity and shallower depths, seine hauls in glides or runs are
typically longer and use wider nets. You can use a 1.25 m deep seine in shallow glides.
iv) Pull the bag completely to shore at a predesignated point.
e) Edge Habitats
i) Edge habitats may be shallow too deep with complex to uniform habitat, and may include
undercut banks.
ii) Use two crewmembers, each tending a seine pole. Seine along the nearshore area.
iii) The near bank crewmember moves along the shore while jabbing along any undercut or small
structure. The other crewmember stays ahead of the shoreline pole to maintain a "J" in the
seine bag. At a predesignated point, post the near shore pole and sweep the seine towards
and up on the bank.
iv) Depending on edge complexity and depth, seine width and haul length may vary. Use wider
seines and longer hauls in shallower, less complex habitats. As complexity, depths, and flow
increase, shorten the seine width and haul length accordingly. Seine depth may vary
depending on depth.
f) Snag Habitats
i) Snag habitats often require creativity in terms of seine length and approach. You can use a
1.25 meter deep seine to avoid snagging the net on structure, but use a 2 m deep seine in
deeper areas. Narrow seine widths and short hauls are preferred.
ii) Use two crewmembers, each tending a seine pole. Jab seining is often the most effective
method. Quickly jab a shortened seine (< 2 m wide) under the cover and near the river
bottom, then quickly lift the seine to the water surface. You can use a third crewmember to
agitate the snag to move fish out toward the seine.
iii) For small snags along the bank, seining along the edge may work best. The near snag
crewmember moves along the snag, while jabbing along its length. The other crewmember
stays ahead of the shoreline pole to maintain a "J" in the seine bag. At a predesignated point,
quickly pull the seine to the surface.
9) To minimize mortality, process fish (i.e., identify, count, and prepare preserved voucher specimens or
photovoucher images) after each seine haul (rather than at the end of a subreach.
a) Record identifications, tallies, and voucher information on the fish collection form (Figure 10.2).
You may use separate lines on the fish collection form to record species information from
separate seine hauls.
iv) For each seine haul, record seine characteristics and haul length, habitat, and time on the
seining information form (Figure 10.4).
10) 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 sections of the fish
gear form.
11) Indicate whether conditions allowed for sufficient sampling on the fish gear form (Yes, No, Not Sure). If
you marked No or Not Sure, explain why in the Sampling Protocol Comments field.
12) Record the total length of the river that was sampled for fish on the fish gear form. This total length
should coincide with the end of a subreach.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
J NRSA 2018/19 SEINING INFORMATION j
Site ID: Date: / / Page of
a>
LL
BANK: L = Left Bank (facing downstream R = Right Bank (facing dow, earn) CH = In Channel
HABITAT SAMPLED: PL = Pool. GL = GSde/Run. RF = Riffle. ED = Edge. SN = Snag. BW = Backwater or side channel. OT = Other (describe)
FLAG CODES: K = No measurement made. U = Suspect measurement., F1 ,F2, etc. = flags assigned by each field crew. Explain al flags in comments.
Flag Comments
•
If)
CM
o\
U>
CO
o
co
0
CO
|
1
c
o»
c
c
to
CO
Habitat Sampled (choose one) If other, describe:
is
£
6
O
U- Z
C£ tO
oo
a S
OO
si E
oo
s
.C
5
o
u- Z
CC (0
oo
o S
oo
a §
oo
8
£
6
o
u- Z
a; to
oo
o s
oo
_j 3
a 03
oo
01
£
6
O
u. Z
a: to
OO
d s
oo
CL 1
oo
y
£
8
O
u. z
a: to
OO
o S
OO
cl m
OO
c
6
o
u. z
ce to
oo
o S
oo
oo
a
£
6
O
u. z
ce «
OO
o £
OO
a! w
OO
A
¦
£
6
o
u- Z
a: to
OO
(s s
OO
a! E
OO
3
£
6
O
u- Z
a: to
OO
d s
oo
a 1
oo
5
£
6
o
LL Z
ce y>
oo
o 2
oo
~L £
oo
\
¦c
6
o
u- z
ce y>
oo
d s
oo
a :f:
OO
y-
5
o
. z
ce v>
oo
o S
OO
CL
OO
<5
£
6
O
u- Z
a: v>
OO
<* e
OO
a! -j
OO
i
r
If
c
u- Z
Ce tO
00
d s
oo
CL &
00
5
y
c
u- Z
ir
oo
d 2
oo
CL E
oo
8
X
0
o
DC
o
o
X
o
O
DC
o
o
X
o
O
a:
o
_J
o
X
u
o
K
o
o
X
o
O
0£
o
o
X
o
o
ce
o
o
X
u
O
a
o
o
X
u
O
tc
O
o
X
o
o
o
to
CO
31
o
rsi
fl
in
Figure 10.4 Seining Information Form
LU
(£5
10.5 Processing Fish 3
Process the fish at the end of each subreach or pairs of subreaches, as described in Table 10.6. lu
CO
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 x
processing. Always process and release individuals of State or Federally listed threatened or d
123
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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 the information for each haul on a separate line
on the seining information form.
10.5.1 Identification and Tallying
Record species identifications, tallies, and other information for individuals collected on the Fish
Collection Form (Figure 10.2). Use multiple pages of the form as needed to record all species
collected. It is important to record page numbers correctly because page number is one of the
variables used to uniquely identify a species record. You may record separate collections of the
same species on multiple lines of the collection form (e.g., when you encounter a species in non-
adjacent subreaches, or collect a species with a secondary gear type). Do not process individuals
with total length < 25 mm (1 inch), as these are likely young of year individuals that cannot be
identified confidently to species. Only crew members designated as "taxonomic specialists" by
EPA regional coordinators can identify fish species. Tally fish by species and major size class (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 collection
form. If you suspect it represents a potential range extension for the species, prepare one or
more specimens (preserved if possible but photographs if not). Physical specimens are required
in order to publish reports of range extensions. Include specimens to document suspected range
extensions are included as part of the preserved Unknown/Range Extension voucher sample
(UNK/RNG; Section 10.5.6).
10.5.2 Unknown Specimens
If you cannot positively identify individuals to species in the field, record taxonomic information
of the collection form using scientific names rather than common names. If you can identify a
specimen only to family, record the scientific rather than the common family name (e.g.,
UNKNOWN PERCID A, not UNKNOWN PERCH A) on the fish collection form. If you can identify a
specimen to genus, record the scientific name rather than the common name (e.g., UNKNOWN
PERCINA A, not UNKNOWN DARTER) on the fish collection form. Using scientific rather than
common names for unknowns reduces ambiguity, since some common names may in fact refer
to multiple genera (e.g., "darter", "shiner", "sucker", "sunfish", etc.). If you identify an unknown
species to Genus, retain a small number (up to 20 individuals per putative species) as part of the
preserved UNK/RNG voucher sample (see Section 10.5.6) or take good digital photographs
(Section 10.5.3) for laboratory identification. If you are only able to identify an unknown to
Family, retain as many of the individuals as possible for later identification. Use the UNK/RNG
Voucher label on the label sheet to label your jar of unknown to track from which sites the
unknowns originated.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 10.6 Procedure: Processing Fish (Wadeable Sites)
Fish Processing
1)
Complete all header information accurately and completely on the fish collection form. It is important
to paginate the collection forms correctly (e.g., start with page 1, do not duplicate page numbers,
etc.), as page number is part of the unique record identifier for the fish count data.
2)
Process individuals collected at the end of each subreach. You may record a single species on multiple
lines of the collection form (e.g., use separate lines for individuals collected in multiple subreaches,
collect with a secondary gear type, or if you need additional space to record tally marks, etc.).
a) Process species listed as threatened and endangered first as described in Step 4.
i) Photograph specimens for voucher purposes if conditions permit and stress to individuals
will be minimal. Mark as Photo in the Voucher section of the collection form.
ii) If individuals die due to sampling, prepare them as part of the local voucher sample and
preserve them in the field. Comply with the conditions of your collection permit in
regards to mortality of listed species.
iii) Return individuals to the river immediately after processing.
3)
Only identify and process individuals > 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 the NRSA 2008/09 and 2013/14.
(1) Record the complete common name. Avoid using shortened names (e.g., stoneroller,
carp, bass, etc.).
(2) If you use a non-standard name, you must assign a flag to the line and provide the
taxonomic reference for the name in the Comments section of the collection form.
b) If you cannot positively identify an individual to the species level:
i) Identify it to the lowest taxonomic level (i.e., family or genus). Record the putative name
as UNKNOWN plus the scientific name of the family or genus (e.g., UNKNOWN
CATOSTOMID A, UNKNOWN MOXOSTOMA A) in the Common Name column of the
collection form.
ii) If you are permitted to retain the specimen, assign it the next available sequential tag
number (starting with 01) in the Voucher Tag Number column and see Step 9.
c) If you believe the individual is a hybrid:
i) Mark as Hybrid? on the collection form.
ii) If the hybrid has an accepted standard common name (e.g., Tiger muskellunge, Saugeye,
Wiper, etc), record that name. For other hybrids record the common name of both species
(e.g., Green sunfish x Bluegill, Cutthroat trout x Rainbow trout). Avoid using non-specific
terms such as Hybrid sunfish.
iii) If you are unsure of the identification and are permitted to retain the specimen, assign it
the next available sequential tag number (starting with 01) in the Voucher Tag Number
column and see Step 9.
5)
If you know the species is not native to this location, mark as Introduced? If you cannot evaluate the
native/introduced status of fish collected, mark the bubble 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.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
<
I
CO
U~)
LD
<
126
6) Visually estimate the total length of each individual (a measuring board is not necessary). Keep a
running tally in the appropriate Tally and Counts section (< 6 in., 6-12 in., 12-18 in., or > 18 in.) of the
fish collection form.
a) If all individuals of a species appear to be the same size, provide a flag and comment for the line
if you believe the population is stunted.
7) Examine each individual for external anomalies. Readily identify external anomalies including missing
organs (eye, fin), skeletal deformities, shortened operculum, eroded fins, irregular fin rays or scales,
tumors, lesions, ulcerous sores, blisters, cysts, blackening, white spots, bleeding or reddening,
excessive mucus, and fungus. After you process all of the individuals of a species, record the total
number of individuals observed with one or more anomaly in the Anom Count column of the
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. After you process all of the individuals of a species, record the total number observed in the
Mortality/Count column of the collection form.
9) If you are retaining individuals of the species as part of the preserved Unknown/Range Extension
Unk/Rng) voucher sample:
a) Mark as Unk/Rng in the Voucher section of the collection form.
b) Assign the species the next available voucher specimen tag, and record the number in the
Voucher Tog# column of the collection form.
i) If you take one or more photographs of the species instead of preserving specimens,
assign the next available voucher specimen tag number in the Voucher Tag # column of
the collection form. Include the specimen tag in all photos of the species. Mark Photo
in the Voucher section of the collection form.
ii) Ideally, take photos of all species collected at a site that are not being preserved.
c) Record the number of individuals retained for the preserved voucher sample in the Vouchers
Retained column of the collection form.
i) NOTE: Do not keep separate tallies of voucher and non-voucher specimens. Record all
individuals in the appropriate area of the Tally and Counts section. The retained
voucher specimens represent a subsample of the total count.
ii) Place the specimens in a jar which has been labeled with the site ID. You can have
multiple individuals of the same species in the jar, but each species will have a separate
voucher tag number (i.e. one tag number per line on the collection form).
10) If you are retaining specimens as part of a preserved QA voucher sample for the site:
a) Mark as QA in the Voucher section of the fish collection form.
i) NOTE: This should be marked at least once for all species collected at the site (including
unknowns).
b) Use the sheet of labels and tags for the QA voucher sample (the jar label has a preprinted sample
ID number). Assign the species the next available voucher specimen tag number. Record the
specimen tag number in the Voucher Tag # column of the collection form.
i) If you take one or more photographs of the species instead of preserving specimens,
assign the next available voucher specimen tag number, and record the number in the
Voucher Tag # column of the collection form. Include the specimen tag in all photos of
the species. Mark Photo in the Voucher section of the collection form.
c) Record the number of individuals retained for the preserved voucher sample in the Vouchers
Retained column of the collection form.
i) NOTE: Do not keep separate tallies of voucher and non-voucher specimens. Record all
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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).
11) Repeat Steps 2 through 10 for each subreach sampled. Use additional fish collection form sheets as
needed, being careful to paginate each sheet correctly.
12) Record the fish collected with seining methods on a separate line on the field form.
13) At the end of sampling, follow the appropriate procedure to prepare the preserved voucher samples
(UNK/RNG and/or QA) and/or select specimens for tissue samples.
a) For all voucher samples, use a sufficient volume of 10% buffered formalin (the volume of
formalin solution used must exceed the volume of specimens). Use additional jars if necessary.
Slit large individuals (TL > 200 mm [~8 in.]) along the right side in the lower abdominal cavity to
allow penetration of the formalin solution.
14) Complete a sample jar label for the UNK/RNG voucher sample. Attach it to the sample jar and cover it
with clear tape.
15) If you did not prepare a QA voucher sample, mark No Voucher Preserved on the back of the fish gear
form (this is akin to the 'no sample collected' bubble 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.
b) Record QA voucher sample label information on the back of the fish gear form.
16) Record the file names of any photovouchers taken on the back of the fish gear form.
a) Use only one line per voucher tag, even if you took multiple photos (record the beginning and
end of the sequence in the Sequence column). Make sure the page and line numbers you record
match those on the collection form.
b) Name image files as: Site ID + Visit number + tag number + sequence (e.g.,
N RS 18_WY_10001_V l_tagO la).
17) If you did not collect any fish from the entire fish sampling reach, mark Fished - None Collected in the
Fish Sampling - Not Conducted or Suspended section of the fish gear form.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
10.5.3 Photovouchering
Use digital imagery for fish species that cannot be preserved as voucher specimens (e.g., rare,
threatened, and endangered species; very large bodied). Ideally, take photos of all species
collected a site (that are not preserved) to provide a minimal level of documentation of
occurrence. Take photographs of entire specimens and additional specific morphological
features that are appropriate and necessary for an independent taxonomist to accurately
identify the specimen. Additional detail for these guidelines is provided in Stauffer et al. (2001),
which is provided to all field crews in electronic format.
The recommended specifications for digital images to be used for photovouchering include: 16
bit color at a minimum resolution of 1024x768 pixels; macro lens capability allowing for images
to be recorded at a distance of less than 4 cm; and built-in or external flash for use in low light
conditions. Specimens (or morphological features) should occupy as much of the field of view as
possible. Use a fish measuring board, ruler, or some other calibrated device to provide a
reference to scale. Provide an adequate background color for photographs (e.g., fish measuring
board). Include a card with site ID number, site name, and date in each photograph so that
photos can be identified if file names become corrupted. In addition, if the specimen is part of
either the unknown/range extension or QA voucher collection, include the voucher specimen
tag that you assign to the species to provide a link to the line on the fish collection form. For
each photovoucher specimen, include at least a full body photo (preferably of the left side of the
fish), and other macro images of important morphological features (e.g., lateral line, ocular/oral
orientation, fin rays, gill arches, mouth structures, etc.). It may also be necessary to photograph
males, females, or juveniles to depict key identifying features.
Save images in medium to high quality jpeg format. It is important that time and date stamps
are accurate, as this information can also be useful in tracking the origin of photographs.
Transfer images stored in the camera to a personal computer (PC) or external storage device
(e.g., thumb drive or flash memory card) at the first available opportunity. At this time, rename
the original files to include the site ID, visit number, voucher specimen tag number, and photo
sequence (e.g., NRS18_WY_10001_Vl_tag01a.jpg). Record the file names on the back of the fish
gear form (Figure 10.6). 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.
10.5.4 Preparing Preserved Voucher Specimen Samples
There are two different types of samples for preserved voucher specimens. The UNK/RNG
voucher samples are used to identify specimens that cannot be confidently identified in the
field, and to provide physical specimens of suspected range extensions. After submitting the fish
collection form to the NARS IM staff, you will receive an update form that lists only the records
for unknown species recorded on the fish collection form (including phototvouchers) that were
marked as being part of the UNK/RNG voucher sample.
In addition to a UNK/RNG voucher sample (if needed), you will prepare an additional QA voucher
sample (Section 10.5.7). A QA voucher sample will be performed at a pre-designated set of sites
and includes preserved specimens (or photographs) of all species collected at a site (including
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
the unknowns). Use the voucher specimen tags and sample labels designated for QA voucher
samples. QA voucher samples are eventually sent to an independent taxonomist as a check on
the accuracy of each fish taxonomist.
10.5.5 Preserving Voucher Specimen Samples
Preserve UNK/RNG and QA voucher specimens in the field with a 10% buffered formalin
solution. The volume of formalin must be equal to or greater than the total volume of
specimens. Use additional jars if necessary to ensure proper preservation. For individuals having
a total length larger than 200 mm (~8 in.), make a slit along the right side of the fish in the lower
abdominal cavity to allow penetration of the preservative solution. Follow all the precautions for
handling formalin outlined in the MSDS. Formalin is a potential carcinogen. Handle with
extreme caution, as vapors and solution are highly caustic and may cause severe irritation on
contact with skin, eyes, or mucus membranes. Wear vinyl or nitrile gloves and safety glasses,
and always work in a well-ventilated area.
Once you have completed preserving all jars of voucher specimens, complete the appropriate
jar label (Figure 10.5 for UNK/RNG samples, and Figure 10.7 for QA voucher samples). Attach
the completed label to the jar and cover with clear shipping tape. Two jar labels are provided for
each type of voucher collection. If you have > 2 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
labels). 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.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
FISH VOUCHER - UNK/RNG
Site ID: NRS18
I / 201
FISH VOUCHER - UNK. RNG
Site ID: NRS18
/ / 201
FISH - BAG
TAG: 41
FISH - BAG
TAG; -12
FISH - BAG
TAG;43
FISH-BAG
TAG: 44
FISH - BAG
TAG: 37
FISH - BAG
TAG. 38
FISH — BAG
TAG: 39
FISH - BAG
TAG:40
FISH - BAG
TAG: 33
FISH - BAG
TAS: 34
FISH - BAG
TAG: 35
FISH - BAG
TAG: 36
FISH-BAG
TAG; 29
FISH-BAG
TAG: 30
FISH - BAG
TAG: 31
FISH - BAG
TAG:32
FISH - BAG
TAG; 25
FISH-HAS
TAG; 26
FISH - BAG
TAG;27
FISH - BAG
TAG: 28
FISH - BAG
TAG*. 21
FISH - BAG
TAS; 22
FISH - BAG
TAG: 23
FISH - BAG
TAG: 24
FISH-BAG
TAG;1?
FISH-BAG
TAG: 18
FISH - BAG
TAG:19
FISH - BAG
TAG: 20
FISH - BAG
TAG: 13
FISH - BAG
TAG; 14
FISH - BAG
TAG: 15
FISH - BAG
TAG: 16
FISH-BAG
TAG; 09
FISH - BAG
TAG: 10
FISH - BAG
TAG.11
FtSH - BAG
TAG: 12
FISH - BAG
TAG; §5
F ISH - BAG
TAG: 06
FISH - BAG
TAG: 07
FtSH - BAG
TAG: 08
FISH-BAG
TAG: 01
FISH - BAG
TAG: 02
FISH - BAG
TAG: 03
FtSH - BAG
TAG: 04
^2 Figure 10.5 Unknown/Range Extension Voucher Sample Labels and Voucher Specimen Tags
i/i
i/i
<
130
-------
crq
c
fD
I—4
O
QJ
Q.
en
QJ
3
T3
3
QJ
a-
QJ
¦ NRSA 2018/19 FISH GEAR AND SAMPLING INFORMATION (Back) g
Site ID: Date: / /
QA VOUCHER SAMPLE INFORMATION (VERT)
NO VOUCHERS PRESERVED O
Sample ID
# of Jars
Preserved
Comments
o
PHOTO VOUCHER FILE INFORMATION
Page
Line
Photo File Name (SitelD V(Visit#) Tag#)
(e.g.: NRS18_OR_11158_V1_Tag01)
Sequence
(e.g. a-c)
Comments
Page
Line
Photo File Name (SitelD V(Visit#) Tag#)
(e.g.: NRS18_0R_11158_V1_Tag01)
Sequence
(e.g. a-c)
Comments
FISH TISSUE PLUG SAMPLES (FPLG)
NO SAMPLE COLLECTED O
Collection Method: O ELECTROFISHING O HOOK & LINE O SEINING O OTHER:
Sample ID
Common Name
Length(mm)
Weight(g)
Comments
03/28£018 NRSA18 Fish Gear and Sampling Information
1604063189 0
< 2
fD QJ
-! r+
~ o'
O -j
c
D
ro
NJ
o
<
0)
qj
3
Q.
t/>
fD
QJ
3
3
CD
D
r+
NJ
O
31
fD'
Q_
O
¦a
fD
-1
QJ
r+
O'
£
3
CO
QJ
Q.
2
fD
QJ
QJ
3
cr
C
fD
QJ
M
CO
I-1
FISH ASSEMBLAGE
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
10.5.6 Processing Unknown/Range Extension (UNK/RNG) Voucher Samples
Table 10.7 outlines the procedure for determining the identification of unknown specimens
from each UNK/RNG sample. A more detailed procedure for conducting the laboratory
identifications is presented in the NRSA laboratory operations manual (USEPA 2013b). Identify
unknown specimens using whatever resources are necessary (magnification, literature,
reference collections/specimens, including dissected anatomical features or in-house
colleagues).
Following positive laboratory identification, use the fish collection revision form for the sample
(Figure 10.8) to reconcile the unknown records to reflect revisions to the actual species
identifications, counts, and any other information recorded on the original collection form. It is
important to update counts and identifications by voucher tag—do not combine multiple
samples of the same unknown before updating.
The fish collection revision form for a site will be provided with all of the original information
that was recorded on the original collection form. The form is a fillable Portable Document
Format (pdf) form, and corrections can be made using the Adobe Reader® software. For each
line, make any corrections on page 1 of the revision form, and provide an explanation of the
changes made on the corresponding line number on the page 2 of the revision form (Figure
10.9), and mark the YES button under Changes Made?. If no corrections are necessary, mark
the NO button on the back side of the form under Changes Made? Provide your contact
information in the space provided on the back side of the form.
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 column, and
enter any changes to the original counts in the appropriate Counts column. If you determine
that a single unknown record is actually >1 species, replace the original UNKNOWN record with
the information for the most abundant species. Record the information for additional species
from this original unknown as new data records (use blank lines on the revision form), but retain
the page, line number, and 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), enter the page, line
number, tag number, and taxonomic reference for the name in the Comments section on page 2
of the revision form (Figure 10.9). Submit your completed revision forms to the NARS IM staff 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@glec.com, 715-829-3737). There are provisions under which fish can be identified by a
contracted lab and the results returned to you.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
10.5.7 Processing QA Voucher Samples
Prepare the QA voucher sample as outlined in Table 10.8. Prepare the QA voucher sample
separately from the UNK/RNG voucher sample. Processing involves ensuring that the sample
jar(s) and photovoucher files include 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 10.7).
Record information about the preserved QA voucher sample on the back side of the fish gear
form (Figure 10.6).
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
whether the samples contain formalin or ethanol, the size of individual bottles, and on the
particular shipping service used.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
QA FISH VOUCHER (VERT)
Site ID: NRS18
/ / 201
999012
QA FISH VOUCHER (VERT)
Site ID: NRS18
/ / 201
999012
FISH — BAG
TAG:
41
999012
FISH - BAG
TAG:
42
999012
FISH-BAG
TAG:
43
999012
FISH-BAG
TAG:
44
999012
FISH - BAG
TAG:
37
999012
FtSH - BAG
TAG:
36
999012
FISH - BAG
TAG:
38
999012
FISH - BAG
TAG:
40
999012
FISH - BAG
TAG:
33
899012
FISH ¦ BAG
TAG;
34
999012
FISH - BAG
TAG:
35
999012
FISH - BAG
TAG:
36
999012
FISH — BAG
TAG:
29
999012
FISH - BAG
TAG;
30
999012
FISH — BAG
TAG:
31
939012
FISH - BAG
TAG:
32
999012
FISH - BAG
TAG:
25
999012
FISH - BAG
TAG:
26
999012
FISH-BAG
TAG;
27
999012
FISH-BAG
TAG:
28
999012
FISH - BAG
TAG:
21
999012
FISH - BAG
TAG;
22
999012
FISH - BAG
TAG:
23
999012
FISH - BAG
TAG:
24
999012
FISH - BAG
TAG:
17
999012
FISH - BAG
TAG:
18
999012
FISH - BAG
TAG:
19
999012
FISH-BAG
TAG:
20
999012
FISH - BAG
TAG:
13
999012
FISH - BAG
TAG:
14
999012
FISH - BAG
TAG:
15
999012
FISH-BAG
TAG:
16
999012
FISH - BAG
TAG:
09
999012
FISH - BAG
TAG:
10
999012
FISH - BAG
TAG:
11
999012
FISH - BAG
TAG:
12
999012
FISH - BAG
TAG:
06
999012
FISH ¦¦ BAG
TAG:
08
999012
FISH - BAG
TAG:
07
999012
FISH - BAG
TAG:
08
999012
FISH - BAG
TAG:
01
§98012
FISH - BAG
TAG:
02
§99012
FISH - BAG
TAG:
03
999012
FISH - BAG
TAG:
04
999012
^2 Figure 10.7 QA Voucher Sample Labels and Voucher Specimen Tags
i/i
i/i
<
134
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 10.7 Procedure: Processing Unknown/Range Extension (UNK/RNG) Voucher Samples
Processing UNK/RNG Voucher Samples
1) Following fixation for 5 to 7 days, decant and properly discard the formalin solution. Formalin is a
potential carcinogen and should be used with extreme caution, as vapors and solution are highly
caustic and may cause severe irritation on contact with skin, eyes, or mucus membranes. Wear vinyl or
nitrile gloves and safety glasses, and always work in a well-ventilated area.
a) Formalin must be disposed of properly. Contact your regional EPA coordinator if your laboratory
does not have the capability of handling waste formalin.
2) Replace the formalin with tap water and soak specimens over a 4-5 day period. Soaking may require
periodic water changes and should continue until the odor of formalin is barely detectable.
3) Decant the tap water. Use 45%-50% isopropyl alcohol or 70% ethanol as a final preservative for
specimens.
4) The NARS IM staff will send you a Fish Collection Revision Form for each fish collection form you
completed at a site e . This form lists all records from the original collection form, including any that
were marked as being part of the UNK/RNG voucher sample. It will be provided as a fillable form in a
portable document format (pdf) file. Identify unknown fish to species in the laboratory, using the
procedure described in the NRSA laboratory operations manual, which is briefly described below.
a) Process unknowns by tag number—do not combine multiple bags (or jars) of the same unknown
before determining the final identifications. Corrections and updates need to be linked back to
the original page and line number, as well as the voucher specimen tag number you recorded on
the collection form.
5) Make any corrections to the original collection form on the revision form using software that can edit
a fillable pdf file (e.g., Adobe Reader®'. For every line where you make a correction on page 1 of the
revision form, provide an explanation on the corresponding line on page 2 of the form.
a) Use common names from the standard list (Appendix D) as revised names.
b) If you must use a non-standard name, provide the page, line number, specimen tag number, and
the taxonomic reference in the Comments section on page 2 of the revision form.
6) If an unknown turns out to include > 1 species, correct the final counts based on the proportion of
each species found in the original unknown bag.
a) Record the revised name and count for one species (the most abundant) on the line of the original
unknown.
b) Record the revised name and count for the second species as a new record on the revision form.
i) Record the page, line number, and specimen tag number from the original unknown record
on the next available blank line of the update form.
ii) Fill in the appropriate button under the Changes Made? section on page 2 of the revision
form to indicate whether or not any changes were made. Provide your contact information
on page 2 of the revision form.
7) After reconciling all of the unknowns, and correcting any other information from the original collection
forms, submit the completed revision forms to the NARS IM staff in Corvallis. Retain the preserved
UNK/RNG voucher samples. Contact your regional EPA Regional Coordinator if you cannot store the
samples at your facility.
-------
UU FISH ASSEMBLAGE
cn
era
c
n>
i—4
o
bo
n
o
n>
o
73
n>
<
"O
0)
era
fD
^ NRSA 2018/19 FISH COLLECTION REVISIONS
Site ID, Visit, Date Collected (Chem), Crew Fish Collection Date (Actual Date) Page
I did not evaluate the native/
introduced status for fish collection. O Y
Lin©
#
Common Name
c-
. TJ
>. -c
X £
(*•
el
I 1
Counts
Anom
Count
Mortality
Vouchers
Vouchers
Retained
Flag
<150
mm
150-
300 mm
300-
460 mm
>460
mm
Count
Unk/Rng
QA
Photo
Tag#
1
o
o
o
o
o
2
o
o
o
o
o
3
o
o
o
o
o
4
o
0
o
o
o
5
o
0
o
o
o
6
o
o
o
o
o
7
o
o
o
o
o
8
o
o
o
o
o
9
o
0
o
o
o
10
o
o
o
o
o
11
o
o
o
o
o
12
o
o
o
o
o
13
o
o
o
o
o
14
o
o
o
o
o
The above filled data represents your fish file when these records were pulled from the database. We need you to confirm that we have accurate information regarding all aspects of your fishing event.
Please make updates here as needed to correctly identify species, update counts and other information. If you make any changes to fish data on this side, please provide a reason for change on the back
of this form corresponding to the line number. Your reasons for change are very helpful when reviewing the data and documenting changes in the database for OA purposes.
For instance, if you change unknown darter on line 12 to Johnny Darter - please tell us that on the backside of this page, line 12. If an unknown species turns out be >1 species, record the most abundant
species on the line replacing the unknown, and revise the count and any other information. Add the name and information for any new species to an empty line (or use the blank form provided to you). If
the flag field is filled, please review your flag and comment data provided to you separately in csv format. If your comment is no longer applicable, remove the flag here, open the csv file and delete
the flag and comment there. Fill in reason for change on both the revision form and the csv file.
Thisform isyourfinal attempt at correcting your fish data. Do not make corrections to fish collection via app submissions. Please return all pages of record - whether or not there were
changes. Indicate if changes were made on the back and give us your contact information should we have any follow-up questions. Thank you in advance for your attention to detail.
^ 6859616124 A
02/15/2018 NRSA18 Fish Collection Revisions
< 2
fD Qj
O
3
L_ <
c 3>
D cn
HD a)
3
Q.
t/>
fD
QJ
3
3
fD
D
c+
NJ
O
ro_
Q_
o
¦a
fD
o
3
Q_
ro
ro
-------
era
c
n>
M
o
Id
n
o
73
n>
<
o
£
¦O
QJ
era
fD
¦
NRSA 2018/19 FISH COLLECTION REVISIONS J
Site ID, Visit, Date Collected (Chem), Crew Fish Collection Date (Actual Date) Page
If you made any changes to the front of this form, please provide a reason for those changes here corresponding by line.
1
2
3
4
5
6
7
8
9
10
11
12
13
14
Changes Made? Please provide contact information in case we have a question.
Name:
O YES - Chanaefs) were necessary and reasonfsl noted above.
Email address:
Phone:
•
02/15/2018 NRSA18 Fish Collection Revisions
3067616129 0
<
ro
o
3
C
D
ro
NJ
o
<
fD
CD
3
Q.
LT>
a>
QJ
3
3
fD
D
r+
NJ
O
ro_
Q_
o
¦a
fD
o
3
Q.
ro
CX
fD
LU FISH ASSEMBLAGE
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 10.8 Procedure: Processing QA Voucher Samples
1) Ensure that all species collected at a site are represented by either preserved voucher specimens or
photovouchers. There should be a unique QA voucher specimen tag 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(5)glec.com,
715-829-3737).
3) After preparing the preserved QA voucher sample, check that the sample ID number recorded on the
fish gear form matches the preprinted label attached to each sample jar, and that the number of jars
recorded on the fish gear form is correct.
4) Retain the QA voucher samples in appropriate storage space for formalin until you receive information
regarding where to send them from the NRSA staff at EPA Office of Water, or EPA Regional
Coordinator.
5) If you are storing the preserved QA voucher samples for an extended period, you may need to replace
the formalin fixative with ethanol.
a) Following fixation for 5 to 7 days, decant and properly discard the formalin solution. Formalin is a
potential carcinogen handle with extreme caution, as vapors and solution are highly caustic and
may cause severe irritation on contact with skin, eyes, or mucus membranes. Wear vinyl or nitrile
gloves and safety glasses, and always work in a well-ventilated area.
b) Formalin must be disposed of properly. Contact your regional EPA Regional Coordinator if your
laboratory does not have the capability of handling waste formalin.
6) Replace the formalin with tap water and soak specimens over a 4-5 day period. Soaking may require
periodic water changes and should continue until the odor of formalin is barely detectable.
7) Decant the tap water. Use 45%-50% isopropyl alcohol or 70% ethanol as a final preservative for
specimens.
8) When ready to ship all of the QA voucher samples, complete a sample tracking form as described in
Appendix C.
9) Package the preserved samples properly for either formalin or ethanol and prepare all required
documentation and safety measures for the shipment.
10) Post all photovoucher files for each QA voucher sample to SharePoint. Use the file names that are
recorded on the fish gear form.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
11 FISH TISSUE PLUG SAMPLING METHODS
11.1 Method Summary
Because many fish spend their entire life in a particular water body they can be important
indicators of water quality, especially for toxic pollutants (e.g., pesticides and trace elements).
Toxic pollutants, which may be present in the water column or sediments at concentrations
below our analytical detection limits, can be found in fish tissue due to bioaccumulation.
Typical fish tissue collection methods require the fish to be sacrificed, whether it be a whole fish
or a skin-on fillet tissue sample. This can be problematic when there is a need to collect large
trophy-sized fish for contaminant analysis or when a large sample size is necessary for statistical
analysis. The following describes an alternative method for the collection of fish tissue samples
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 except during any site visit where whole fish tissue
samples are collected (see Section 12). These fish are collected during the fish assemblage
sample collection effort (Section 10). 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. Fish tissue plugs will not be
removed in the field from whole fish tissue samples collected at the 477 designated river sites
(Section 12). Instead, those plug samples will be extracted in the laboratory by the fish sample
preparation lab personnel.
11.2 Equipment and Supplies
Table 11.1 lists the equipment and supplies necessary for Field Crews to collect fish tissue plug
samples. This list is comparable to the checklist presented in Appendix A, which provides
information to ensure that Field Crews bring all of the required equipment to the site. Record
the fish tissue plug sampling data on the Fish Gear and Voucher/Tissue Sample Information
Form (Figure 10.6).
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 11.1 Equipment and Supplies: Fish Tissue Plug Sample
For fish tissue plug samples
Fish measuring board
Fish weigh scale
Plastic bags
Sterile 20 mLglass scintillation vial
Coolers with ice
Cooler with dry ice
Nitrile gloves
8 millimeter disposable biopsy punch (Acuderm brand Acu-Punch or
equivalent)
Sterile disposable scalpel
Sterile forceps
Laboratory pipette bulb.
Antibiotic salve.
Fish collection gear (electrofisher, nets, livewell, etc.)
Dip net
Field Operations Manual and laminated Quick Reference Guide
For recording measurements
Fish tissue plug sample labels
Fish Gear and Sampling Form
Soft (#2) lead pencils for recording data on field forms
Fine-tipped indelible markers for filling out sample labels
Clear tape strips for covering labels
11.3 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 11.2. If the target species are unavailable, the fisheries biologist will
select an alternative species (i.e., a species that is commonly consumed in the study area, with
specimens of harvestable or consumable size) to obtain a plug sample. Recommended and
alternate target species are given in Table 11.2. The procedures for collecting and processing
fish plug samples are presented in Table 11.3.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 11.2 Recommended Target and Alternate Species for Fish Tissue Plug Collection
Family name
Common name
Scientific name
Length Guideline
(Estimated
Minimum)
Centrarchidae
Spotted bass
Micropterus punctulatus
~280 mm
Largemouth bass
Micropterus salmoides
~280 mm
Smallmouth bass
Micropterus dolomieu
~300 mm
Black crappie
Pomoxis nigromaculatus
~330 mm
White crappie
Pomoxis annularis
~330 mm
Ictaluridae
Channel catfish
Ictalurus punctatus
~300 mm
Blue catfish
Ictalurus furcatus
~300 mm
Flathead catfish
Pylodictis olivaris
~300 mm
Percidae
Sauger
Sander canadensis
~380 mm
Walleye
Sander vitreus
~380 mm
Yellow perch
Perca flavescens
~330 mm
Moronidae
White bass
Morone chrysops
~330 mm
Esocidae
Northern pike
Esoxlucius
~430 mm
Chain pickerel
Esox niger
~430 mm
Salmonidae
Brown trout
Salmo trutta
~300 mm
Cutthroat trout
Oncorhynchus clarkii
~300 mm
Rainbow trout
Oncorhynchus mykiss
~300 mm
Brook trout
Salvelin us fon tin alis
~330 mm
Cyprinidae Northern pikeminnow Ptychocheilus oregonensis ~300 mm
Bluegill Lepomis macrochirus ~200 mm
Centrarchidae Rock bass Ambloplites rupestris ~200 mm
Redbreast sunfish Lepomis auritus ~200 mm
a>
a.
to
+-»
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Fish Tissue Plug Methods
i/i
Q
o
<
i/i
3
3
i/l
i/l
142
• are of similar size, so that the smaller individual is no less than 75% of the total length of
the larger individual.
4. Remove one fish retained for analysis from the clean holding container(s) (e.g., livewell) using clean
nitrile gloves.
5. Measure the fish to determine total body length. Measure total length of the specimen in
millimeters, from the anterior-most part of the fish to the tip of the longest caudal fin ray (when
the lobes of the caudal fin are depressed dorsoventrally).
6. Weigh the fish in grams using the fish weigh scale.
7. Note any anomalies (e.g., lesions, cuts, sores, tumors, fin erosion) observed on the fish.
8. Record site ID, date, sample ID, species, and specimen length and weight on the back of the Fish
Gear and Sampling Form in the Fish Tissue Plug section (Figure 10.6). Make sure the sample ID
numbers and specimen numbers/lengths that are recorded on the collection form match those on
the sample tracking form and labels where applicable.
9. Prepare a Sample Identification Label for the sample, ensuring that the label information matches
the information recorded on the Fish Tissue Plug section of the Fish Gear and Sampling Form. Affix
label to a sterile 20 milliliter scintillation vial and cover with clear tape.
10. On a meaty portion of the left side dorsal area of the fish between the dorsal fin and the lateral
line, clear a small area of scales with a sterile disposable scalpel.
11. Wearing clean nitrile gloves, insert the 8 millimeter biopsy punch into the dorsal muscle of the fish
through the scale-free area. The punch is inserted with a slight twisting motion cutting the skin and
muscle tissue. Once full depth of the punch is achieved a slight bending or tilting of the punch is
needed to break off the end of the sample. Remove biopsy punch taking care to ensure sample
remains in the punch. Note: The full depth of the punch should be filled with muscle tissue, which
should result in collecting a minimum of 0.25 to 0.35 grams 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.
15. Place the sample immediately on dry ice for shipment.
16. Dispose of gloves, scalpel and biopsy punch.
17. Keep the samples frozen on dry ice or in a freezer at <-20°C until shipment.
18. Frozen samples will subsequently be packed on dry ice and shipped to the batched sample
laboratory via priority overnight delivery service within 1 week.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
12 WHOLE FISH SAMPLING METHOD
12.1 Method Summary
Fish are important integrators of toxic contaminants that are bioavailable in the water column
and in sediment. EPA monitors the occurrence of toxic chemicals in fish fillet samples to assess
the potential health impacts to people who consume fish. Results from the NRSA 2008/09
provided the first statistically representative national data for fish contamination in U.S. rivers.
Collecting whole fish tissue samples and submitting them to the laboratory for filleting and
homogenization during the NRSA 2018/19 allows consistency with fish tissue methods of
previous NRSAs (2008/09 and 2013/14) and provides sufficient tissue for analysis of multiple
chemical contaminants of concern (e.g., mercury, polychlorinated biphenyls or PCBs, and
perfluorinated compounds of PFCs). Continued analysis of fillet tissue also allows for temporal
analysis of probability-based national fish contamination trends in U.S. rivers. Collecting fish at
locations sampled during previous NRSAs will reduce the variability in data for trends analysis.
Whole fish tissue sampling procedures are described in detail in Table 12.3. The objective is to
collect one whole fish sample from each of the 477 designated target river sites. The focus is on
obtaining fish species that are commonly consumed by humans, that satisfy legal requirements
of harvestable size for each river site (or at least consumable size if no legal harvest
requirements exist), and that are sufficiently abundant within a sampling reach. Each whole fish
tissue sample will consist of five adult fish of the same species that are similar in size (i.e., the
smallest individual in the sample is no less than 75% of the total length of the largest individual).
Collection occurs anywhere in the fish assemblage sampling reach (Section 10). Whole fish
samples are shipped to the laboratory designated for interim storage of the samples. Fish
sample preparation laboratory staff fillet the fish and homogenize the fillet tissue for analysis of
mercury and other contaminants (e.g., PCBs and PFCs).
12.2 Equipment and Supplies
Table 12.1 lists the equipment and supplies necessary for Field Crews to collect whole fish tissue
samples. This list is comparable to the checklist presented in Appendix A, which provides
information to ensure that Field Crews bring all of the required equipment to the site. Record
the fish tissue sampling data on the Whole Fish Tissue Collection Form (Figure 12.1).
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 12.1 Equipment and Supplies: Whole Fish Tissue Sample Collection
For collecting whole
fish tissue sample
Electrofishing equipment (including Coast Guard approved personal
variable voltage pulsator unit, wiring floatation devices
cables, generator, electrodes, dip nets, Maps of target sites & access
protective gloves, boots, and necessary routes G|oba, positioning System
safety equipment) (GPS) unit
Scientific collection permit Livewe|| and/or buckets
Sampling vessel (including boat, motor, Measuring board (millimeter
trailer, oars, gas, and all required safety scale)
equipment) Clean nitrile gloves
For storing and
preserving whole
fish tissue sample
Aluminum foil (solvent rinsed and baked) „
Knife or scissors
Heavy-duty food grade polyethylene ^
tubing
. . , Plastic cable ties
Large plastic (composite) bags
Coolers
Composite (Tyvek) tag
For documenting the
whole fish tissue
sample
Whole Fish Tissue Collection Form Sample Identification Labels
Clipboard Black ink pen
Clear tape strips Fine tipped indelible markers
Tracking Form
For shinnine the Chain-of-custody labels
hor snipping tne Preaddressed FedEx airbill
whole fish tissue Packing/strapping tape
„m„. Coolers
samples Hazard Class 9 shipping labels
Dry ice
12.3 Sampling Procedures
The whole fish tissue samples will be collected with the same gear used to collect the fish
assemblage samples. Collection of individual specimens for whole fish samples occurs anywhere
in the sample reach during the fish assemblage sampling. Ideally, each fish sample will contain 5
fish of the same species that are similar in size. Depending on the size of the fish, fewer than 5
fish may be acceptable or more than 5 fish will be necessary to meet the 500-gram fillet tissue
requirement for chemical analysis and archived tissue (refer to Frequently Asked Questions in
the whole fish tissue kits). Recommended target species are given in Table 12.2. If the target
species are unavailable, the fisheries biologist will select an alternative species to obtain a whole
fish sample (i.e., a species that is commonly consumed by humans, with specimens that are of
harvestable or consumable size and are in sufficient numbers to yield a fish sample with
adequate tissue for analysis). If sufficient fish are not collected during the fish assemblage
sampling, sample for up to one additional hour (collections can occur in areas/subreaches not
otherwise sampled if desired). If no fish can be collected, record "no sample collected" on the
whole fish tissue collection form, along with the reason in the comments section of the form.
The procedures for collecting and processing whole fish tissue samples are presented in Table
12.3.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 12.2 Recommended Target Species for Whole Fish Tissue Collection
Family name
Common name
Scientific name
Length
Guideline
(Estimated
Minimum)
Spotted bass
Micropterus punctulatus
~280 mm
Largemouth bass
Micropterus salmoides
~280 mm
Centrarchidae
Smallmouth bass
Micropterus dolomieu
~300 mm
Black crappie
Pomoxis nigromaculatus
~330 mm
White crappie
Pomoxis annularis
~330 mm
Channel catfish
Ictalurus punctatus
~300 mm
0J
"
Ictaluridae
Blue catfish
Ictalurus furcatus
~300 mm
a.
to
a!
Flathead catfish
Pylodictis olivaris
~300 mm
£?
ro
l-
Sauger
Sander canadensis
~380 mm
Percidae
Walleye
Sander vitreus
~380 mm
Yellow perch
Perca flavescens
~330 mm
Moronidae
White bass
Moron e chrysops
~330 mm
Esocidae
Northern pike
Esoxlucius
~430 mm
Chain pickerel
Esox niger
~430 mm
Brown trout
Salmo trutta
~300 mm
Salmonidae
Cutthroat trout
Oncorhynchus clarkia
~300 mm
Rainbow trout
Oncorhynchus mykiss
~300 mm
Brook trout
Salvelin us font in alis
~330 mm
Table 12.3 Sampling Procedures for Whole Fish Tissue Samples
Whole Fish Tissue Method
Q
0
1
I—
LU
z
i
CL
<
l/l
I
Ll_
LU
I
0
1
5
145
1. Put on clean nitrile gloves before handling the fish. Do not handle any food, drink, sunscreen, or
insect repellant until after the whole fish sample has been collected, measured, and wrapped.
Rinse potential target species/individuals in ambient water to remove any foreign material from the
external surface and place in clean holding containers (e.g., livewells, buckets). Return non-target
fishes or small specimens to the river or stream.
3. Collect one target species sample from each designated site. The sample should consist of 5 fish of
adequate size to provide a total of 500 grams of edible tissue for analysis (refer to Table 12.2 for
minimum species length guidelines). Select fish for each sample based on the following criteria:
• all are of the same species,
• all satisfy legal requirements of harvestable size for the sampled river, or at least be of
consumable size if no legal harvest requirements are in effect,
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Whole Fish Tissue Method
• all are of similar size, so that the smallest individual in a composite is no less than 75% of
the total length of the largest individual, and
• all are collected as close to the same time as possible, but no more than one week apart
(Note: Individual fish may have to be frozen until all fish to be included in the sample are
available for delivery to the designated laboratory).
Accurate taxonomic identification is essential in assuring and defining the organisms that have
been collected and submitted for analysis. Do not mix species in a single whole fish sample.
4. Measure each individual fish to determine total body length. Measure total length of each
specimen in millimeters, from the anterior most part of the fish to the tip of the longest caudal fin
ray (when the lobes of the caudal fin are depressed dorsoventrally).
5. Record site ID, date, sample ID, species (common name), and specimen length on the Whole Fish
Tissue Collection Form (Figure 12.1) in black ink. Fill in site type ("Wadeable" or "Boatable") at the
top of the form. Address the two sample criteria in the space above the fish specimen data to
confirm compliance. All samples must meet these two criteria (i.e., fish are all the same species and
fish lengths are all within 75% of the largest specimen length). Make sure the sample ID numbers
and specimen numbers/lengths that are recorded on the collection form match the corresponding
information on each individual specimen label.
6. Remove each fish selected for analysis from the clean holding container(s) (e.g., livewell) using
clean nitrile gloves. Dispatch each fish using a clean wooden bat (or equivalent wooden device).
7. Wrap each fish in extra heavy-duty aluminum foil with the dull side in (foil provided by EPA as
solvent-rinsed, oven-baked sheets).
8. Prepare a Sample Identification Label for each sample, ensuring that the label information matches
the information recorded on the Whole Fish Tissue Collection Form.
9. Cut a length of food grade tubing (provided by EPA) that is long enough to contain each individual
fish and to allow extra length on each end to secure with cable ties. Place each foil wrapped
specimen separately into an appropriate length of tubing. Seal each end of the tubing with a plastic
cable tie. Attach the fish sample label to the outside of the food grade tubing with clear tape and
secure the label by taping around the entire fish (so that tape sticks to tape).
10. Place all the wrapped fish in the whole fish composite sample from each river site in a large plastic
composite sample bag, complete the bag label, and tape it to the Tyvek composite tag, then seal
the composite bag with a cable tie with the composite sample tag attached.
11. After each sample is packaged, place it immediately on dry ice for shipment. If samples will be
carried back to a laboratory or other facility to be frozen before shipment, wet ice can be used to
transport wrapped and bagged fish samples in the coolers to a laboratory or other interim facility.
12. If possible, keep all (five) specimens designated for a particular sample in the same shipping
container (ice chest) for transport.
13. Samples may be stored temporarily on dry ice (replenishing the dry ice daily). You have the option,
depending on site logistics, of:
• shipping the samples packed on dry ice in sufficient quantities to keep samples frozen for
up to 48 hours (50 pounds are recommended), via priority overnight delivery service (e.g.,
Federal Express), so that they arrive at Microbac Laboratories (Baltimore, MD) within less
than 24 hours from the time of sample collection, or
• freezing the samples within 24 hours of collection at <-20°C, and storing the frozen
samples until shipment within 2 weeks of sample collection (frozen samples will
subsequently be packed on dry ice and shipped to Microbac Laboratories (Baltimore, MD)
via priority overnight delivery service).
14. Ship fish tissue samples to the designated laboratory for interim sample storage on Monday
through Thursday (no Saturday delivery to the laboratory).
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
¦
NRSA 2018/19 WHOLE FISH TISSUE COLLECTION |
o WADEABLE Q BOATABLE
Site ID: Date: /
/ PAGE: OF
WHOLE FISH TISSUE FILLET SAMPLE (FTIS) SAMPLE ID:
1.
NO SAMPLE COLLECTEDO
i I i 1 1 »
O FISH ARE ALL THE SAME SPECIES
O FISH ALL WITHIN 75% OF LARGEST SPECIMEN
Total
Common Name
Length (mm)
Frozen
Comments
.01
o
.02
o
.03
G
.04
O
.05
O
.06*
G
.07*
G
,0B*
G
.09*
G
.10*
G
'Additional specimens for smaller fish species to ensure sufficient tissue is available for chemical analysis of fillet tissue.
¦
10/26/2017 NRSA18 Fish Tissue Collection
9371531329 |
Figure 12.1 Whole Fish Tissue Collection Form
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
13 FINAL SITE ACTIVITIES
13.1 Overview of Final Site Activities
Prior to leaving the site, make a general visual assessment of the site and its surrounding
catchment. The objective of the site assessment is to record observations of catchment and site
characteristics that are useful for future data interpretation, ecological value assessment,
development of associations, and verification of stressor data. Your observations and
impressions are extremely valuable.
You will filter and process the fecal indicator, chlorophyll-a, and periphyton samples, as well as.
Conduct a final check of the data forms, labels and samples. The purpose of the second check of
data forms, labels and samples is to assure completeness of all sampling activities. Finally, clean
and pack all equipment and supplies, and clean the launch site and staging areas. After you
leave the site, report the sampling event to the IM Coordinator, and ship or store the samples.
Activities described in this section are summarized in Figure 13.1.
LEAVE SITE
COMMUNICATIONS
SHIP SAMPLES
COMPLETE SITE
ASSESSMENT
(4 People)
PACK EQUIPMENT AND
SUPPLIES FOR TRANSPORT
(2 People)
LOAD BOAT ONTO TRAILER;
CLEAN UP LAUNCH SITE
AND STAGING AREA
(2 People)
REVIEW DATA FORMS
(Crew Leader)
Completeness
Accuracy
Legibility
Flags/Comments
INSPECT BOAT, MOTOR,
TRAILER, AND NETS FOR
PRESENCE OF PLANT AND
ANIMAL MATERIAL, AND
CLEAN THOROUGHLY
(3 People)
REVIEW SAMPLE LABELS
(Crew Leader)
Completeness
Accuracy
Legibility
Cross-check with forms
FILTER, PRESERVE, &
INSPECT SAMPLES
(3 People)
Complete
Sealed
Ice packs
Packed for transport
Figure 13.1 Final Site Activities Summary
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
13.2 General Site Assessment
Complete the Site Assessment Form (Figure 13.2) after sampling, recording all observations
from the site that were noted during the course of the visit. This Site Assessment Form is
designed as a template for recording pertinent field observations. It is by no means
comprehensive, and any additional observations should be recorded in the General Assessment
section.
13.2.1 Elevation at Transect K
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.
13.2.2 Watershed Activities and Disturbances Observed
Record any of the sources of potential stressors listed in the "Watershed Activities and
Disturbances Observed" section on the Site Assessment Form (Figure 13.2). Include those that
were observed while on the site, while driving or walking through the site catchment, or while
flying over the site and catchment. For activities and stressors that you observe, rate their
abundance or influence as low (L), moderate (M), or heavy (H) on the line next to the listed
disturbance. Leave the line blank for any disturbance not observed 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, fill in "L" for low next to "Residences." If the site is ringed with houses, rate it
as heavy (H). Similarly, a small patch of clear-cut logging on a hill overlooking the site would rate
a low ranking. Logging activity right on the site shore, however, would get a heavy disturbance
ranking. This section includes residential, recreational, agricultural, industrial, and stream
management categories.
13.2.3 Site Characteristics
Record observations regarding the general characteristics of the site on the Site Assessment
Form (Figure 13.2). When assessing these characteristics, look at a 200 m riparian distance on
both banks. Rank the site between "pristine" and "highly disturbed", and between "appealing"
and "unappealing." Document any signs of beaver activity and flow modifications. Record the
dominant land use and forest age class. Document the weather conditions on the day of
sampling and any extreme weather conditions in the days prior to sampling.
13.2.4 General Assessment
Record any additional information and observations in this narrative section. Information to
include could be observations on biotic integrity, vegetation diversity, presence of wildlife, local
anecdotal information, or any other pertinent information about the site or its catchment.
Record any observations that may be useful for future data interpretation.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Site ID:
NRSA 2018/19 ASSESSMENT (Front)
Date: /
Reviewed by (initial):.
Elevation at transect K:
Oft Om
WATERSHED ACTIVITIES AND DISTURBANCES OBSERVED
(Intensity: Blank=Not observed, L=Low, M=Moderate, H=Heavy)
BLANK FIELD INDICATES ABSENCE Q
Residential
Recreational
Agricultural
Industrial
Stream Management
O
©
O Residences
0
©
0 Hiking Traits
©
©
0 Cropland
©
©
0 Industrial Plaits
©
©
0 Liming
O
O
0 Maintained Laws
O
©
O Parks, Campgrounds
o
o
0 Pasture
o
0
0 Minss'Quarries
o
0
0 Chemical Treatment
O
o
O Const met wn
O
©
O Primitive Parks, Camping
o
0
0 Livestock Use
o
©
0 Oil/Gas Wells
o
©
0 Angling Pressure
0
©
0 Pipes, Drains
O
©
0 Trash'Littei
0
o
0 Orchards
0
©
0 Power Plants
o
©
0 Dredgoog
0
o
0 Dumping
o
©
0 Surface Fims, Scums,
©
©
0 Poultry
©
©
© Logging
o
©
0 Channelization
o
o
0 Roads
or Slicks
O
©
0 Feedot
o
©
0 Evidence of Fire
©
o
0 Water Level Fluctuations
O
o
0 Bridges/Causeway
o
©
0 Watef Withdrawa
o
0
0 Odors
©
o
0 Fish Stocking
0
o
0 Sewage Treatment
o
©
0 Commercial
o
©
0 Dams
SITE CHARACTERISTICS (200m radius)
WATERBODY CHARACTER
PRISTINE: 05 04 03 02 01 Highly Disturbed
APPEALING: 05 04 03 02 Ol Unappealing
BEAVER
Beaver Signs: O Absent O Rare O Common
Beaver Flow Modifications: O None O Minor O Major
DOMINANT LAND USE
Dominant Land Use Around 'X' O Forest O Agriculture O Range O Urban O Suburban/Town
If Forest, Dominant Age Class O 0 - 25 yrs. O 26 - 75 yrs O > 75 yrs.
WEATHER
CONDITIONS AND LOCAL CONTACTS
OBSERVATIONS (e.g. accessibility, boating, fishing, swimming, health concerns):
09/13/2017 NRSA18 Assessment
4803036903
>
b
Figure 13.2 Site Assessment Form (front)
IS)
I
<
150
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
hevwweq pviinioai*:
• NRSA 2018/19 ASSESSMENT (Back) #
Site ID: Date: / /
GENERAL ASSESSMENT AND COMMENTS
INVASIVE OR NUISANCE SPECIES OF LOCAL INTEREST
Record species of plants and animals that were observed but are not on the invasive plant form. Examples would be Zebra Mussel or
New Zealand Mud Snail, or invasive plants or animals of concern to a particular state. Indicate your level of confidence in your
identification, and provide some idea of how prevalent it is in the sampling reach or adjacent riparian area.
Species (Common Name) Confidence Prevalence Comments
O LOW
O high
O DOMINANT o SPARSE
O COMMON
O LOW
O high
O DOMINANT O SPARSE
O COMMON
O LOW
O high
O DOMINANT O SPARSE
O COMMON
O LOW
O HIGH
O DOMINANT O SPARSE
O COMMON
O LOW
O high
O DOMINANT O SPARSE
O COMMON
O LOW
O high
O DOMINANT O SPARSE
O COMMON
O LOW
O high
O DOMINANT O SPARSE
O COMMON
O LOW
O high
O DOMINANT O SPARSE
O COMMON
# 4aeoo3S9oe #
w 09/13/2017 NRSA18 Assessment
Figure 13.3 Site Assessment Form (back)
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
13.3 Processing the Fecal Indicator [Enterococci), Chlorophyll-a,
and Periphyton Samples
13.3.1 Equipment and Supplies (Fecal Indicator Filtering)
Table 13.1 provides the equipment and supplies needed for Field Crews to collect the fecal
indicator sample.
Table 13.1 Equipment and Supplies: Fecal Indicator (Enterococci) Sample Processing
For processing samples
Nitrile gloves
sterile screw-cap 50 mL PP tube
Filtration apparatus with collection flask
Sterile filter holder, Nalgene 145/147
Vacuum pump (electric pump may be used if available)
Sterile phosphate buffered saline (PBS)
Osmotics 47 mm polycarbonate sterile filters
Sterile disposable forceps
Petri dishes (60 x 15, disposable)
2 sterile microcentrifuge tubes containing sterile glass beads
1 additional sterile microcentrifuge tube if collecting filter blank
Bubble bag
Zip-top bag
Dry ice
Cooler
Field Operations Manual and laminated Quick Reference Guide
For recording measurements
Sample Collection Form
Soft (#2) lead pencils for recording data on field forms
Fine-tipped indelible markers for filling out sample labels
Fecal Indicator sample labels (2 vial labels and 1 bag label)
Filter blank label if collecting filter blank
13.3.2 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 13.2. The sample must be filtered and frozen
within six hours of collection.
Table 13.2 Procedure: Fecal Indicator (Enterococci) Sample Processing
Filtering for the fecal indicator (Enterococci) sample
1. Put on nitrile gloves.
2. Set up sample filtration apparatus on flat surface and attach vacuum pump. Set out 50 mL
sterile PP tube, sterile 60 mm Petri dish, two bottles of chilled phosphate buffered saline
(PBS), Osmotics 47 mm polycarbonate sterile filter box, and two filter forceps.
3. Chill Filter Extraction tubes with beads on dry ice.
4. Aseptically transfer two polycarbonate filters from filter box to base of opened Petri dish.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Close filter box and set aside.
5. Remove the pre-loaded cellulose nitrate (CN) filter (the filter with grid design on it) from
funnel and discard. Be sure to leave the support pad in the filter funnel.
6. Load filtration funnel with sterile polycarbonate filter on support pad (shiny side up).
7. Shake sample bottle(s) 25 times to mix well.
8. Measure 25 mL of the mixed water sample in the sterile graduated 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 mL and 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 and the volume of buffer
rinsate each filter was rinsed with on the Sample Collection Form, Side 2. Record the
filtration start time and finish time for the sample as well as the time the filters were frozen.
19. Repeat steps 6 to 15 for the remaining 50 mL sub-sample volume to be filtered. Make every
effort to filter the same volume of sample through each of the two filters.
Processing Procedure—fecal indicator (Enterococci) filter blank
Enterococci filter blanks will be prepared at all revisit sites during the first visit. Prepare the filter
blanks before filtering the river sample.
1. Set up sample filtration apparatus using same procedure as used for the river sample. Chill Filter
Extraction tubes with beads on dry ice.
2. Aseptically transfer 1 polycarbonate filter from filter box to base of opened Petri dish. Close filter
box and set aside.
3. Remove the pre-loaded cellulose nitrate (CN) filter (the filter with grid design on it) from funnel
and discard. Be sure to leave the support pad in the filter funnel.
4. Load filtration funnel with sterile polycarbonate filter on support pad (shiny side up).
5. Measure 10 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.
>
b
CO
i
<
153
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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(s) 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 filling in the "Blank Collected" button on the
Sample Collection Form.
13.3.3 Equipment and Supplies (Chlorophyll-a from Water Sample Filtering)
Table 13.3 provides the equipment and supplies needed to process the chlorophyll-a water
sample.
Table 13.3 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
For recording measurements
Sample Collection Form
Sample labels
#2 pencils
Fine-tipped indelible markers
Clear tape strips
>
b
<
LU
H
l/l
i
<
13.3.4 Procedures for Processing the Chlorophyll-a Water Sample
The procedures for processing chlorophyll-a water samples are presented in Table 13.4.
Whenever possible, sample processing should be done in subdued light, out of direct sunlight.
Table 13.4 Procedure: Chlorophyll-a Sample Processing
Filtering for the chlorophyll a water sample
1. 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 shake the bottle to
154
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
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 visible color, proceed; if not, repeat steps 3 & 4 until
color is visible on the filter or until a maximum of 2,000 mL have been filtered. Record the actual
sample volume filtered on the Sample Collection Form.
6. Rinse the upper portion of the filtration apparatus and graduated cylinder thoroughly with Dl
water to include any remaining cells adhering to the sides and pump through the filter. Monitor
the level of water in the lower chamber to ensure that it does not contact the filter or flow into
the pump. Remove the bottom portion of the apparatus and pour off the water from the bottom
as often as needed.
7. Remove filter funnel from base without disturbing filter.
8. Remove the filter from the holder with clean forceps. Avoid touching the colored portion of the
filter. Fold the filter in half, with the colored side folded in on itself.
9. Place the folded filter into a 50 mL screw-top centrifuge tube and cap. Tighten the cap as tightly
as possible. The cap will seal tightly after an additional % turn past the point at which initial
resistance is met. Failure to tighten the lid completely could allow water to infiltrate into the
sample and may compromise its integrity.
10. Record the sample volume filtered on a chlorophyll label and attach it to the centrifuge tube (do
not cover the volume markings on the tube). Ensure that all written information is complete and
legible. Cover with a strip of clear tape.
11. Wrap the tube in aluminum foil and place in a self-sealing plastic bag labelled with the completed
chlorophyll outer bag label. Cover the outer label with clear tape. Place this bag immediately on
dry ice to freeze.
13.3.5 Equipment and Supplies (Periphyton Sample)
Table 13.5 lists the equipment and supplies needed to process the periphyton sample.
Table 13.5 Equipment and Supplies: Periphyton Samples
For preparing
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 mLgraduated cylinder
Pipette and pipette bulb (2 mL)
3 50 mL screw-top centrifuge tubes
125 mL sterile PETG bottle
60 mL syringe with tip removed
Aluminum foil squares
Forceps
Surgical gloves
deionized water in wash bottle
plastic electrical tape
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
dry ice
wet ice
coolers
formalin
For data recording
Sample Collection Form
Sample labels
Pencils
Fine-tipped indelible markers
Clear tape strips
For cleaning
10% Bleach solution
equipment
13.3.6 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.
13.3.6.1 ID/Enumeration Sample
Prepare the ID/Enumeration sample as a 50 mL aliquot from the composite periphyton sample,
following the procedure presented in Table 13.6. Preserve each sample with 2 mL of formalin.
Record the sample ID number from the container label and the total volume of the periphyton
composite sample in the appropriate fields on the Sample Collection Form as shown in Figure
4.2. Store the preserved samples upright in a container containing absorbent material.
Table 13.6 Procedure: ID/Enumeration Samples of Periphyton
Periphyton ID Sample Processing Procedure
1. Prepare a sample label (with pre-printed sample ID number sample type "PERI"). Record the
volume of the subsample (typically 50 mL) and the volume of the composite index sample on the
label. Attach completed label to a 50 mL centrifuge tube; avoid covering the volume graduations
and markings. Cover the label completely with a clear tape strip.
2. Record the sample ID number of the label and the total volume of the composite index sample on
the Sample Collection Form.
3. 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.
7. Record the volume of the sample in the centrifuge tube (excluding the volume of preservative) on
the Sample Collection Form under the Periphyton Assemblage ID in the Volume Field.
Field Operations Manual
Wadeable
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
13.3.6,2 Periphyton Metagenomic Sample
Prepare the periphyton metagenomic sample as a 100 mL aliquot from the composite index
sample, following the procedure presented in Table 13.7.
Table 13.7 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
volume of the subsample (100 mL) and the volume of the composite index sample 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 sample ID number of the label and the total volume of the composite index sample
on the Sample Collection Form.
3.
Put on surgical gloves (non-powdered).
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.
13.3.6.3 Periphyton Chlorophyll a Sample
Prepare the periphyton chlorophyll a sample by filtering a 25 mL aliquot of the composite index
sample through a 47 mm 0.7 micron GF/F glass fiber filter. The procedure for preparing
periphyton chlorophyll a samples is presented in Table 13.8. Chlorophyll a can degrade rapidly
when exposed to bright light. If possible, prepare the samples in subdued light (or shade),
filtering as quickly as possible after collection to minimize degradation. If using the same
filtration chamber that was used for Enterococci and index site chlorophyll-o samples, rinse it
with deionized water prior to filtering the periphyton chlorophyll-o sample. If you are reusing a
filtration chamber from a previous site, you should rinse it with Dl water each day before use at
the base site and then seal in a plastic bag until use at the stream (be sure to use a new chamber
at each site for the Enterococci sample as it needs to be filtered in a sterile chamber). Keep the
glass fiber filters in a dispenser inside a sealed plastic bag until use.
It is important to measure the volume of the sample being filtered accurately (±1 mL) with a
graduated cylinder. During filtration, do not exceed 7 inches of Hg (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.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 13.8 Procedure: Preparing Chlorophyll Samples of Periphyton
Periphyton Chlorophyll a Sample Processing Procedure
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 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.
3. Rinse a 25 mL or 50 mL graduated cylinder three times with small volumes of deionized water
and discard.
4. Mix the composite sample bottle thoroughly.
5. Measure 25 mL (±1 mL) of sample into the graduated cylinder.
NOTE: For a composite sample containing fine sediment, allow grit to settle for 10 - 20 seconds
before pouring the sample into the graduated cylinder.
6. Pour the 25 mL aliquot into the filter funnel, replace the cap, and pull the sample through the
filter using the vacuum pump. Vacuum pressure from the pump should not exceed 7 inches of Hg
(3.44 psig) to avoid rupture of fragile algal cells.
NOTE: If 25 mL of sample will not pass through the filter, discard the filter and rinse the chamber
thoroughly with deionized water. Collect a new sample using a smaller volume of sample,
measured to ±1 mL. Be sure to record the actual volume sampled on the sample label and the
Sample Collection Form.
7. Monitor the level of water in the lower chamber to ensure that it does not contact the filter or
flow into the pump. Remove the bottom portion of the apparatus and pour off the water from
the bottom as often as needed.
8. Remove the filter chamber from the filter holder being careful not to disturb the filter. Remove
the filter from the holder with clean forceps. Avoid touching the colored portion of the filter.
Fold the filter in half, with the colored sample (filtrate) side folded in on itself. Place the folded
filter in a 50 mL centrifuge tube.
9. Tighten the cap as tightly as possible. The cap will seal tightly after an additional % turn past the
point at which initial resistance is met. Seal the cap with plastic electrical tape.
10. Prepare a sample label (with pre-printed sample ID number sample type "PCHL") including the
volume filtered, and attach it to the centrifuge tube. Cover the label completely with a strip of
clear tape.
11. Place the centrifuge tube into the self-sealing plastic bag with the water column chlorophyll
sample.
12. Record the sample ID number of the label and the total volume of the composite index sample
on the Sample Collection Form. Record the volume filtered in the "Periphyton Chlorophyll" field
on the Sample Collection Form. Double check that the volume recorded on the collection form
matches the total volume recorded on the sample label.
13. Place the centrifuge tube containing the filter on dry ice.
13.3.6.4 Periphyton Biomass Sample (AFDM)
p Prepare the ash-free dry mass (AFDM) sample by filtering a 25 mL aliquot of the composite
^ index sample through a 47 mm 1.2 micron GF/C glass fiber filter. The procedure for preparing
£! AFDM samples is presented in Table 13.9. Using the same filtration chamber that was used for
1/1 Enterococci and chlorophyll-a samples, rinse it with deionized water prior to filtering the
^ periphyton biomass sample. If you are reusing a filtration chamber from a previous site, you
^ should rinse it with Dl water each day before use at the base site and then seal in a plastic bag
158
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
until use at the stream (be sure to use a new chamber at each site for the Enterococci sample as
it needs to be filtered in a sterile chamber). Keep the glass fiber filters in a dispenser inside a
sealed plastic bag until use.
It is important to measure the volume of the sample being filtered accurately (±1 mL) with a
graduated cylinder. During filtration, do not exceed 7 inches of Hg (3.44 psig) to avoid rupturing
cells. If the vacuum pressure exceeds 7 inches of Hg prepare a new sample. If the filter clogs
completely before all the sample in the chamber has been filtered, discard the sample and filter,
and prepare a new sample using a smaller volume of sample.
Table 13.9 Procedure: Preparing Ash-Free Dry Mass (AFDM) Samples of Periphyton
Periphyton AFDM Sample Processing Procedures
1. Rinse the sides of the filter funnel and the filter with a small volume of deionized water to prevent
contamination from the previously filtered sample.
2. Using clean forceps, place a Whatman 47 mm 1.2 micron GF/C glass fiber filters on the filter holder
gridded side down. If needed, use a small amount of deionized water from a wash bottle to help
settle the filter properly. Attach the filter funnel to the filter holder and filter chamber, then attach
the hand vacuum pump to the filter flask.
3. Rinse a 25 mL or 50 mL graduated cylinder three times with small volumes of deionized water and
discard.
4. Mix the composite sample bottle thoroughly.
5. Measure 25 mL (±1 mL) of sample into the graduated cylinder.
NOTE: For a composite sample containing fine sediment, allow grit to settle for 10 - 20 seconds
before pouring the sample into the graduated cylinder.
6. Pour the 25 mL aliquot into the filter funnel, replace the cap, and pull the sample through the filter
using the vacuum pump. Vacuum pressure from the pump should not exceed 7 inches of Hg (3.44
psig) to avoid rupture of fragile algal cells.
NOTE: If 25 mL of sample will not pass through the filter, discard the filter and rinse the chamber
thoroughly with deionized water. Collect a new sample using a smaller volume of sample, measured
to ±1 mL. Be sure to record the actual volume sampled on the sample label and the Sample
Collection Form.
7. Monitor the level of water in the lower chamber to ensure that it does not contact the filter or flow
into the pump. Remove the bottom portion of the apparatus and pour off the water from the
bottom as often as needed.
8. Remove the filter chamber from the filter holder being careful not to disturb the filter. Remove the
filter from the holder with clean forceps. Avoid touching the colored portion of the filter. Fold the
filter in half, with the colored sample (filtrate) side folded in on itself. Place the folded filter in a 50
mL centrifuge tube.
9. Tighten the cap as tightly as possible. The cap will seal tightly after an additional % turn past the
point at which initial resistance is met. Seal the cap with plastic electrical tape.
10. Prepare a sample label (with pre-printed sample ID number sample type "PBIO"), including the
volume filtered, and attach it to the centrifuge tube. Cover the label completely with a strip of clear
tape. Place the centrifuge tube into the self-sealing plastic bag with the water column and
periphyton chlorophyll samples.
11. Record the sample ID number of the label and the total volume of the composite index sample on
the Sample Collection Form. Record the volume filtered in the "Periphyton Biomass" field on the
Sample Collection Form. Double check that the volume recorded on the collection form matches the
total volume recorded on the sample label.
12. Place the centrifuge tube containing the filter on dry ice.
>
b
CO
i
<
159
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
13.3.6.5 Cleaning o/Periphyton Equipment
Once all four laboratory samples have been prepared, discard any remaining sample and
thoroughly clean all periphyton sampling equipment (including brush, delimiter, composite
bottle, funnel, and syringe) with a 10% bleach solution to disinfect the equipment and limit the
possible spread of periphyton DNA to future samples. After cleaning, thoroughly rinse all the
equipment with tap or Dl water. Store the equipment in a clean plastic bag.
13.4 Data Forms and Sample Inspection
After the Site Assessment Form is completed, the Field Crew Leader reviews all of the data
forms and sample labels for accuracy, completeness, and legibility. The other crew members
inspect all sample containers and package them in preparation for transport, storage, or
shipment. Refer to Appendix C for details on preparing samples for shipping.
Ensure that all required data forms for the site have been completed. Confirm that the SITE-ID,
the visit number, and date of visit are correct on all forms. On each form, verify that all
information has been recorded accurately, the recorded information is legible, and any flags are
explained in the comments section. Ensure that written comments are legible, with no
"shorthand" or abbreviations. Make sure there are no markings in the scan code boxes. Make
sure the header information is completed on all pages of each form. After reviewing each form
initial the upper right corner of each page of the form.
Ensure that all samples are labeled, all labels are completely filled in, and each label is covered
with clear plastic tape (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.
13.5 Launch Site Cleanup
Inspect all nets for pieces of macrophyte or other organisms and remove as much as possible
before packing the nets for transport. Pack all equipment and supplies in the vehicle and trailer
for transport. Keep equipment and supplies organized so they can be inventoried using the
equipment and supply checklists presented in Appendix A. Lastly, be sure to clean up all waste
material at the launch site and dispose of or transport it out of the site if a trash can is not
available.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
14 FIELD QUALITY CONTROL
Standardized training and data forms provide the foundation to help assure that data quality
standards for field sampling are met. This section for field sampling and data collection are the
primary guidelines for all cooperators and Field Crews. In addition, repeat sampling and field
evaluation and assistance visits will address specific aspects of the data quality standards for the
NRSA.
14.1 Revisit Sampling Overview
Revisit sampling will provide data to make variance estimates (for measurement variation and
index period variation) that can be used to evaluate the NRSA design for its potential to
estimate status and detect trends in the target population of sites. A summary of the repeat
sampling design is provided in Figure 14.1.
Space revisits minimum of two
weeks to one month apart
Revisits = Measurement
Variation + Index period
variation
Revisit Sites (4 per State)
Filter Blank
Enterococci
BEFORE filtering
other samples
Col lect all
Samples
In situ measures
Water chemistry
Chlorophyll-a
Fteriphyton
Benthos
Enterococci
Fish
Fish plugs
Physical habitat
Col lect all
samples
In situ measures
Water chemistry
Chlorophyll-a
Fteriphyton
Benthos
Enterococci
Fish
Fish plugs
Fish tissue
(at select sites)
Physical habitat
Figure 14.1 Summary of the Repeat Sampling Design
As described in Sections 11 and 12, 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) at all sites where
suitable fish species and lengths are available except during any site visit where whole fish tissue
samples are collected. Additionally, whole fish tissue samples are to be collected at only one of
the two visits to a revisit site (ideally visit 1). Figure 14.2 describes the decision making process §
regarding the collection of fish plugs versus whole fish tissue. z
O
u
>
H
i
<
3
a
Q
i
LU
161
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Revisit Sites
Second Visit
Whole Fish
Tissue Site
No whole fish
sample collected
If
—
then
If
No fish plug
sample collected
then
Figure 14.2 Summary of Fish Tissue Protocol for Revisit Sites
14.2 Revisit Sampling Sites
A total of 200 (approximately 10%) of the target sites visited will be revisited during the same
sample year by the same Field Crew that initially sampled the site. Revisit samples and
measurements are taken from the same reach as the first visit. Each state has four revisit sites;
two wadeable and two non-wadeable sites. For each state these sites are:
Wadeable Revisit sites:
• The two wadeable revisit sites are re-samples from the NRSA 2008/09 and NRSA
2013/14 (l-4th order; labeled as Strahler categories Large Stream (LS) or Small Stream
(SS)). The base sites are labeled as _08TS3R2 and _13TS2R2, respectively, and are
located in the Base/Oversample panel of the Resampled Streams tab within the site
evaluation spreadsheet.
Non-Wadeable Revisit sites:
• The two non-wadeable revisit sites are resamples from the NRSA 2008/09 and NRSA
2013/14 (5th order and above, labeled as Strahler categories RiversOther (RO) or
RiversMajor (RM)). The base sites are labeled as _08TS3R2 and _13TS2R2, respectively,
and are located in the Base/Oversample panel of the Resampled Rivers tab within the
site evaluation spreadsheet.
Fish Tissue
Plug Site
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
If a site selected for revisit sampling is dropped, then the oversample site (which will also carry
the _R2 designation) assigned to replace it should be revisited. If there are no _R2 oversample
sites remaining in the panel, re-designate the next _R2 Base site in the panel as a Revisit site
AND replace the base site with an appropriate oversample site. The primary purpose of this
"revisit" set of sites is to collect temporal replicate samples to provide variance estimates for
both measurement variation and index period variation. The revisit will include the full set of
indicators and associated parameters. The time period between the initial and repeat visit to a
site is, not less than 2 weeks and not more than one month. Label the samples Visit 2 to indicate
that they are samples from the second sampling event at a revisit site. We will not be collecting
replicate data on whole fish tissue. Whole fish tissue samples will only be collected on the first
visit.
At each revisit site, a filter blank will be collected for Enterococci during the first sampling visit
(Visit 1). The crews will filter a small amount (10 mL) of sterile buffer through 1 filter, label them
and write "blank" on the label and field form, and package and submit these samples to the lab.
The filter blanks should be run before the sample is filtered. (Figure 14.1). Detailed description
of the filter blanks is found in Table 13.2.
14.3 Field Evaluation and Assistance Visits
A rigorous program of field and laboratory evaluation and assistance visits has been developed
to support the National Rivers and Streams Assessment Program. These evaluation and
assistance visits are explained in detail in the 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).
14.3.1 Specifications for QC Assurance Field Assistance Visits
Field evaluation and assistance personnel are trained in the specific data collection methods
detailed in 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 14.1 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.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Table 14.1 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.
14.4 Reporting
When the sampling operation has been completed, the Evaluator will review the results of the
evaluation with the Field Crew before leaving the site (if practicable), noting positive practices
and problems (i.e., weaknesses [might affect data quality] or deficiencies [would adversely
affect data quality]). The Evaluator will ensure that the Field Crew understands the findings and
will be able to perform the procedures properly in the future. The Evaluator will record
responses or concerns, if any, on the Field Evaluation and Assistance Check List. After the
Evaluator completes the Field Evaluation and Assistance Check List, including a brief summary of
findings, all Field Crew members must read and sign off on the evaluation.
If the Evaluator's findings indicate that the Field Crew is not performing the procedures
correctly, safely, or thoroughly, the Evaluator must continue working with this Field Crew until
certain of the Crew's ability to conduct the sampling properly so that data quality is not
adversely affected. If the Evaluator finds major deficiencies in the Field Crew operations (e.g.,
major misinterpretation of protocols, equipment or performance problems) the Evaluator must
contact the following QA official:
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.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
15 REFERENCES
American Red Cross. 1979. Standard First Aid and Personal Safety. American National Red Cross.
269 pp.
Bain, M.B., and N.J. Stevenson (editors). 1999. Aquatic habitat assessment: common methods.
American Fisheries Society, Bethesda, Maryland.
Baker, J.R., D.V. Peck, and D.W. Sulton (editors). 1997. Environmental Monitoring and
Assessment Program Surface Waters Field Operations Manual for Lakes. EPA/620/R-
97/001. U.S. Environmental Protection Agency, Washington DC.
Dunne, T., and L.B. Leopold. 1978. Water in environmental planning. W.H. Freeman, New York.
818 p.
Flotemersch, J.E.I, B.C. Autrey2, and S.M. Cormierl (editors). 2000. Logistics of Ecological
Sampling on Large Rivers. U.S. Environmental Protection Agency, Cincinnati OH.
Flotemersch, J.E., J.B.Stribling, and M.J. Paul. 2006. Concepts and Approaches for the
Bioassessment of Non-wadeable Streams and Rivers. EPA 600-R-06-127. U.S.
Environmental Protection Agency, Cincinnati, OH.
Harrelson, C.C., C.L. Rawlins, and J.P. Potyondy. 1994. Stream channel reference sites: an
illustrated guide to field technique. USDA Forest Service, General Technical Report RM-
245, Rocky Mountain Forest and Range Experiment Station, Fort Collins, Colorado. 61p.
Helm, W.T. 1985. Aquatic habitat inventory: standard methods and glossary. American Fisheries
Society, Western Division, Bethesda, Maryland.
Hosea, R. C., and B. Finlayson. 2005. Controlling the spread of New Zealand mudsnails on wading
gear. California Department of Fish and Game, Office of Spill Prevention and Response,
Administrative Report 2005-02, Sacramento.
Kaufmann, P.R., P. Levine, E.G. Robison, C. Seeliger, and D.V. Peck. 1999. Quantifying Physical
Habitat in Wadeable Streams. EPA 620/R-99/003. Environmental Monitoring and
Assessment Program, U.S. Environmental Protection Agency, Corvallis, OR.
Lemmon, P.E. 1957. A new instrument for measuring forest overstory density. Journal of
Forestry 55:667-669.
Leopold, L.B. 1994. A view of the river. Harvard University Press, Cambridge, Massachusetts. 298
P-
Linsley, R.K., M.A. Kohler, and J.L.H. Paulhus. 1982. Hydrology for engineers. McGraw-Hill Book
Co. New York, NY. 508 p.
Mulvey, M., L. Cato, and R. Hafele. 1992. Oregon Nonpoint Source Monitoring Protocols Stream
Bioassessment Field Manual: For Macroinvertebrates and Habitat Assessment. Oregon
Department of Environmental Quality Laboratory Biomonitoring Section. Portland,
Oregon. 40pp.
National Institute for Occupational Safety and Health. 1981. Occupational Health Guidelines for
Chemical Hazards (Two Volumes). NIOSH/OSHA Publication No. 81-123.
-------
National Rivers and Streams Assessment 2018/19
Version 1.1 June 2018
Field Operations Manual
Wadeable
Nelson, J.S., E.J. Crossman, H. Espinosa-Perez, L.T. Findley, C.R. Gilbert, R.N. Lea, and J.D.
Williams. 2004. Common and Scientific Names of Fishes from the United States, Canada,
and Mexico. American Fisheries Society, Special Publication 29, Bethesda, Maryland.
Ohio EPA. 1990. Ohio EPA Fish Evaluation Group Safety Manual. Ohio Environmental Protection
Agency, Ecological Assessment Section, Division of Water Quality Planning and
Assessment, Columbus, Ohio.
Pont, D., R.M. Hughes, T.R. Whittier, and S. Schmutz. 2009. A predictive index of biotic integrity
model for aquatic-vertebrate assemblages of western U.S. streams. Transactions of the
American Fisheries Society 138:292-305.
Rantz, S. E., et al., 1982. Measurement and Computation of Streamflow, Vols. 1 and 2. U.S.
Geological Survey Water-Supply Paper 2175.
Robison, E.G. and R.L. Beschta. 1990. Characteristics of coarse woody debris for several coastal
streams of southeast Alaska, USA. 47(9):1684-1693.
Stauffer, Dr. Jay R., J. Karish and T.D. Stecko. 2001. Guidelines for Using Digital Photos as Fish
Vouchers for Pennsylvania Fishes. The Pennsylvania State University and National Park
Service.
Stevens, D. L., Jr., and A. R. Olsen. 2004. Spatially-balanced sampling of natural resources in the
presence of frame imperfections. Journal of American Statistical Association:99:262-
278.
U.S. Coast Guard. 1987. Federal Requirements for Recreational Boats. U.S. Department of
Transportation, United States Coast Guard, Washington, D.C.
USDA Forest Service, 1995. A guide to field identification of bankfull stage in the western United
States. Rocky Mountain Forest and Range Experiment Station, Stream Systems
Technology Center, Fort Collins, Colorado (31 minute video, closed captioned).
USDA Forest Service, 2002. Identifying bankfull stage in forested streams in the eastern United
States. Rocky Mountain Forest and Range Experiment Station, Stream Systems
Technology Center, Fort Collins, Colorado (46 minute video, closed captioned).
USEPA. 1986. Occupational Health and Safety Manual. Office of Planning and Management U.S.
Environmental Protection Agency, Washington, D.C. USEPA. 2009. National Rivers and
Streams Assessment: Field Operations Manual. EPA 841-B-07-009. U.S. Environmental
Protection Agency, Office of Water and Office of Research and Development,
Washington, D.C.
USEPA. 2000a. EPA Quality Manual for Environmental Programs 5360A1. May 2000.
http://www.epa.gov/quality/qs-docs/5360.pdf
USEPA. 2000b. EPA Order 5360.1 A2 CHG2, Policy and Program Requirements for Mandatory
Agency-wide Quality System, May 5, 2000. http://www.epa.gov/quality/qs-docs/5360-
l.pdf
Web Pages:
CO
LU
u US EPA Aquatic Monitoring Research: http://www.epa.gov/nheerl/arm
LU
NHD Plus: http://www.horizon-svstems.com/nhdplus
U_
LU
cc
166
------- |