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United States Office of Research and
Environmental Protection Development
Agency , Washington DC 20460
EPA/620/R-94/004F
September 1998
xvEPA i Surface Waters
Field Operations and
Methods for
Measuring the Ecological
Condition of Wadeable
Streams
Environmental Monitoring and
Assessment Program
-------
-------
EPA/6207R-94/004F
September 1998
ENVIRONMENTAL MONITORING AND ASSESSMENT PROGRAM-
SURFACE WATERS:
FIELD OPERATIONS AND METHODS FOR MEASURING THE
ECOLOGICAL CONDITION OF WADEABLE STREAMS
Edited by
James M. Lazorchak1, Donald J. Klemm1, and David V. Peck2
1 U.S. Environmental Protection Agency
Ecosystems Research Branch
Ecological Exposure Research Division
National Exposure Research Laboratory
Cincinnati, OH 45268
2 U.S. Environmental Protection Agency
Regional Ecology Branch
Western Ecology Division
National Health and Environmental Effects Research Laboratory
Corvallis, OR 97333
NATIONAL EXPOSURE RESEARCH LABORATORY
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
RESEARCH TRIANGLE PARK, NC 27711
NATIONAL HEALTH AND ENVIRONMENTAL EFFECTS RESEARCH LABORATORY
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
RESEARCH TRIANGLE PARK, NC 27711
Printed on Recycled Paper
-------
NOTICE
This research described in this report has been funded wholly or in part by the U.S.
Environmental Protection Agency. This document has been prepared at the EPA National
Exposure Research Laboratory (Ecological Exposure Research Division, Cincinnati, Ohio)
and the National Health and Environmental Effects Research Laboratory (Western Ecology
Division, Corvallis, Oregon), under the following contracts and cooperative agreements:
Contract 68-C6-0006 to Dynamac International, Inc.
Contract 68-C1-0022 to Technology Applications, Inc.
Contract 68-C6-0019 to SoBran, Inc.
Contract 68-W5-0065 to OAO, Inc.
Cooperative Agreement CR824682 to Oregon State University
This work is in support of the Environmental Monitoring and Assessment Program
(EMAP). It has been subjected to the Agency's peer and administrative review, and
approved for publication as an EPA document. Mention of trade names or commercial
products does not constitute endorsement or recommendation for use.
This publication represents the final revision of the EMAP field operations and
methods manual for wadeable streams. Previously, annual revisions have been produced
under the same title and EPA document number (EPA/620/R-94/004). The document
number for the final revision is modified to distinguish it from earlier revisions while
maintaining traceability.
The correct citation for this document is:
Lazorchak, J.M., Klemm, D.J., and D.V. Peck (editors). 1998. Environmental
Monitoring and Assessment Program -Surface Waters: Field Operations and
Methods for Measuring the Ecological Condition of Wadeable Streams. EPA/620/R-
94/004F. U.S. Environmental Protection Agency, Washington, D.C.
Section authors are listed on the following page. Complete addresses for authors
are also provided in each section.
-------
Section 1:
Section 2:
Section 3:
Section 4:
Section 5:
Section 6:
Section 7:
Section 8:
Section 9:
Section 10:
Section 11:
Section 12:
Section 13:
Section 14:
Section 15:
J.M. Lazorchak1, AT. Heriihy2, H.R. Preston3'4, and D.J. Klemm1
B.H. Hill1, F.H. McCormick1, J.M. Lazorchak1, D.J. Klemm1, P.A.
Lewis1-5, V.C. Rogers6-7, and M.K. McDowell5
D.J. Klemm1, B.H. Hill1, F.H. McCormick1, and M.K. McDowell5
AT. Heriihy2
A T. Heriihy2
P R. Kaufmann2
P R. Kaufmann2 and E.G. Robison2-8
B.H. Hill1
B.H. Hill1
J.M. Lazorchak1, and M. E. Smith9
D.J. Klemm1, J.M. Lazorchak1, and P.A. Lewis1'4
F.H. McCormick1 and R. M. Hughes10
R.B. Yeardley, Jr.8, J.M. Lazorchak1, and F.H. McCormick1
J.M. Lazorchak1, A. T. Heriihy2, and J. Green3
J.M. Lazorchak1
1 U.S. EPA, National Exposure Research Laboratory, Cincinnati, OH 45628.
2 Department of Fisheries and Wildlife, Oregon State University, Corvallis, OR 97333.
3 U.S. EPA, Region 3, Wheeling, WV 26003).
4 Current address: Canaan Valley Institute, Davis, WV 26260.
5 Current Address: 1037 Wylie Road, RR #2, Seaman, OH 45679'
6 OAO Corp., Corvallis, OR 97333
7 Current address: Linn-Benton Community College, Albany, OR.
8 Current Address: Oregon Department of Forestry, Salem, OR 97310
So Bran Environmental, Inc., Cincinnati, OH 45628.
Dynamac International, Inc., Corvallis, OR 97333.
iii
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FOREWORD
The National Exposure Research Laboratory (NERL) and the National Health and
Environmental Effects Research Laboratory (NHEERL) provide scientific understanding,
information and assessment tools that will reduce and quantify the uncertainty in the
Agency's exposure and risk assessments for all environmental stressors. Stressors include
chemicals, biologicals, radiation, climate, and land and water use changes.
Research at NERL focuses on: (1) characterizing the sources of environmental
stressors and the compartments of the environment in which they reside or move; (2)
studying the pathways through environmental compartments that lead to exposure of
receptors to stressors; (3) investigating intra- and inter compartmental stressor transfers
and their transformations; and (4) studying and characterizing receptors and their activities
as required to predict or measure stressor exposure. Research products from NERL
provide effects researchers and risk assessors with information on stressor sources,
pollutant transport and transformations and exposure, and state-of-the-science source-to-
receptor predictive exposure models applicable at the appropriate temporal scales and site,
watershed/regional and global scales. It also provides risk managers with receptor-
back-to-source and stressor-back-to-cause analyses and evaluations of alternative
mitigation, management or restoration strategies from an exposure perspective.
Ecological research at NHEERL contribute to improving hazard identification, dose-
response assessments, and risk characterization at multiple spatial and temporal scales.
Research products from NHEERL include improved assessment methods and improved
approaches to interpreting the data acquired by these methods. Major uncertainties in
assessing the effects on ecosystems resulting from exposure to environmental stressors
are addressed through the development of the tools necessary for effective monitoring of
ecosystems and their components, by mechanistic studies, and through modeling.
To accomplish its mission, NERL conducts fundamental and applied research
designed to:
IV
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1. Characterize air, soil, surface water, sediment, and subsurface systems to
evaluate spatial and temporal patterns, exposure to environmental stressors/
pollutants;
2. Identify, quantify, and predict the physical, chemical, biological and biochemical
behavior of stressors, including characterization of their sources, transformations
pathways and other factors that determine stressor exposure to humans and
ecosystems across multiple media
3. Characterize the ecological and human receptors potentially impacted by stress-
ors and pollutants;
4. Measure, predict, and apply data on environmental stressors to characterize
exposure to humans and ecosystems;
5. Incorporate scientific understanding of environmental processes and ecosystem
behavior, along with environmental exposure data, into predictive multimedia
models to estimate exposure and to evaluate mitigation, restoration, prevention
and management options;
6. Develop and implement receptor level exposure and dose models to provide risk
assessors with better and more refined estimates of exposure and dose.
7. Develop chemical, physical, and biological measurement methods to identify and
quantify environmental stressors and to characterize the environment;
8. Develop quality assurance methodologies for chemical, physical, radiological, and
biological analyses;
9. Develop and apply geographical informational systems, remote sensing, photo-
graphic interpretation, information management technologies, software engineer-
ing technologies, computational chemistry, expert systems, and high performance
computing to support the application of exposure and risk assessment tools;
10. Demonstrate, field test/evaluate, and transfer scientific information, measurement
and quality assurance protocols, data bases, predictive exposure and risk
assessment tools, and other innovative exposure assessment technologies, and
provide environmental education materials to support Program Offices, Regions,
State/Municipal/Tribal governments, and other Federal Agencies;
11. Provide technical support to Program Offices, Regions, State/Municipal/Tribal
governments and other Federal Agencies to help in performing state-of-the-
science exposure assessments of known certainty.
Research activities at NHEERL related to improving ecosystem risk assessment are
designed to:
1. Develop and evaluate appropriate and meaningful indicators of ecological
condition and develop associated criteria to characterize condition.
-------
2. Develop and test approaches for monitoring frameworks that are integrated
over multiple spatial and temporal scales to provide representative informa-
tion about spatial extent of ecosystem resources, their current status (i.e.,
baseline condition) and how condition is changing through time.
3. Develop approaches to demonstrate relationships between effects on
ecological condition and the relative magnitude of current stressors at
multiple scales.
This field operations and methods manual represents a collaborative effort among
principal investigators at NERL and NHEERL. The manual describes guidelines and
standardized procedures for evaluating the biological integrity of surface waters of streams.
It was developed to provide the Environmental Monitoring and Assessment Program
(EMAP) with bioassessment methods for determining the status and monitoring trends of
the environmental condition of freshwater streams. These bioassessment studies are
carried out to assess biological criteria for the recognized beneficial uses of water, to
monitor surface water quality, and to evaluate the health of the aquatic environment.
VI
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PREFACE
The Ecosystems Research Branch (ERB), Ecological Exposure Research Division,
National Exposure Research Laboratory, U.S. Environmental Protection Agency - Cincinnati
is responsible for field and laboratory exposure methods and ecological indicators that are
used in assessing aquatic ecosystems. Research areas include the development, evalua-
tion, validation, and standardization of Agency methods for the collection of biological field
and laboratory data. These methods can be used by USEPA regional, enforcement, and
research programs engaged in inland, estuarine, and marine water quality and permit
compliance monitoring, and status and/or trends monitoring for the effects of impacts on
aquatic organisms, including phytoplankton, zooplankton, periphyton, macrophyton,
macroinvertebrates, and fish. The program addresses methods and techniques for sample
collection; sample preparation; processing of structural and functional measures by using
organism identification and enumeration; the measurement of biomass and benthic
metabolism; the bioaccumulation and pathology of toxic substances; acute, chronic, and
sediment toxicity; the computerization, analysis, and interpretation of biological data; and
ecological assessments. ERB also includes field and laboratory support of the ecological
biomarker research program and transfer of monitoring technology to the regions and state
programs.
This document contains the EMAP-Surface Water field operations and bioassess-
ment methods for evaluating the health and biological integrity of wadeable freshwater
streams.
VII
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ABSTRACT
The methods and instructions for field operations presented in this manual for
surveys of wadeable streams were developed and tested during 5 years of pilot and
demonstration projects (1993 through 1997). These projects were conducted under the
sponsorship of the U.S. Environmental Protection Agency and its collaborators through the
Environmental Monitoring and Assessment Program (EMAP). This program focuses on
evaluating ecological conditions on regional and national scales. This document describes
procedures for collecting data, samples, and information about biotic assemblages,
environmental measures, or attributes of indicators of stream ecosystem condition. The
procedures presented in this manual were developed based on standard or accepted
methods, modified as necessary to adapt them to EMAP sampling requirements. They are
intended for use in field studies sponsored by EMAP, and related projects such as the
USEPA Regional Environmental Monitoring and Assessment Program (R-EMAP), and the
Temporally Integrated Monitoring of Ecosystems study (TIME). In addition to methodology,
additional information on data management, safety and health, and other logistical aspects
is integrated into the procedures and overall operational scenario. Procedures are de-
scribed for collecting field measurement data and/or acceptable index samples for several
response and stressor indicators, including water chemistry, physical habitat, benthic
macroinvertebrate assemblages, aquatic vertebrate assemblages, fish tissue contaminants,
periphyton assemblages, sediment community metabolism, and sediment toxicity. The
manual describes field implementation of these methods and the logistical foundation
constructed during field projects. Flowcharts and other graphic aids provide overall
summaries of specific field activities required to visit a stream site and collect data for these
indicators. Tables give step-by-step protocol instructions. These figures and tables can be
extracted and bound separately to make a convenient quick field reference for field teams.
The manual also includes example field data forms for recording measurements and
observations made in the field and sample tracking information. Checklists of all supplies
and equipment needed for each field task are included to help ensure that these materials
are available when required.
vni
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TABLE OF CONTENTS
Section Page
NOTICE H
FOREWORD jv
PREFACE vii
ABSTRACT viii
FIGURES xiv
TABLES . . xvii
ACKNOWLEDGMENTS xx
ACRONYMS, ABBREVIATIONS, AND MEASUREMENT UNITS xxi
1 INTRODUCTION .1
1.1 OVERVIEW OF EMAP-SURFACE WATERS 2
1.2 STREAM SAMPLING COMPONENTS OF EMAP-SURFACE WATERS 3
1.2.1 Mid-Atlantic Highlands Assessment Project 3
1.2.2 Mid-Atlantic Integrated Assessment Program 4
1.2.3 Temporal Integrated Monitoring of Ecosystems Project 4
1.2.4 Other Projects , 5
1.3 SUMMARY OF ECOLOGICAL INDICATORS . , 5
1.3.1 Water Chemistry 5
1.3.2 Physical Habitat 6
1.3.3 Periphyton Assemblage 6
1.3.4 Sediment Community Metabolism 7
1.3.5 Benthic Macroinvertebrate Assemblage 7
IX
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TABLE OF CONTENTS (CONTINUED)
Section
Page
1.3.6 Aquatic Vertebrate Assemblages 8
1.3.7 Fish Tissue Contaminants 9
1.3.8 Sediment Toxicity '• • • 10
1.4 OBJECTIVES AND SCOPE OF THE FIELD OPERATIONS AND METHODS
MANUAL 10
1.5 QUALITY ASSURANCE 11
1.6 LITERATURE CITED 12
2 OVERVIEW OF FIELD OPERATIONS • 17
2.1 DAILY OPERATIONAL SCENARIO 17
2.2 GUIDELINES FOR RECORDING DATA AND INFORMATION 18
2.3 SAFETY AND HEALTH •, • • • 20
2.3.1 General Considerations 20
2.3.2 Safety Equipment and Facilities 24
2.3.3 Safety Guidelines for Field Operations 24
2.4 LITERATURE CITED 26
3 BASE LOCATION ACTIVITIES 27
3.1 ACTIVITIES BEFORE EACH STREAM VISIT 27
3.1.1 Confirming Site Access 27
3.1.2 Daily Sampling Itinerary 29
3.1.3 Instrument Inspections and Performance Tests 29
3.1.3.1 Global Positioning System Receiver 29
3.1.3.2 Dissolved Oxygen Meter 30
3.1.3.3 Conductivity Pens or Conductivity Meters 30
3.1.3.4 Current Velocity Meters 32
3.1.4 Preparation of Equipment and Supplies 32
3.2 ACTIVITIES AFTER EACH STREAM VISIT 36
3.2.1 Equipment Care 36
3.2.2 Sample Tracking, Packing, and Shipment 37
3.3 EQUIPMENT AND SUPPLIES 42
3.4 LITERATURE CITED 44
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TABLE OF CONTENTS (CONTINUED)
Section
Page
4 INITIAL SITE PROCEDURES . , 45
4.1 SITE VERIFICATION ACTIVITIES . . . . . 45
4.1.1 Locating the Index Site ., 45
4.1.2 Determining the Sampling Status of a Stream 48
4.1.3 Sampling During or After Rain Events 48
4.1.4 Site Photographs 43
4.2 LAYING OUT THE SAMPLING REACH .49
4.3 MODIFIED PROCEDURES FOR DRY AND INTERMITTENT STREAMS ..... 53
4.4 EQUIPMENT AND SUPPLIES 53
5 WATER CHEMISTRY 57
5.1 SAMPLE COLLECTION . - .... 58
5.2 FIELD MEASUREMENTS . 53
5.3 EQUIPMENT AND SUPPLIES 59
5.4 LITERATURE CITED 59
6 STREAM DISCHARGE . . . 67
6.1 VELOCITY-AREA PROCEDURE . . . .....' 67
6.2 TIMED FILLING PROCEDURE . . 70
6.3 NEUTRALLY-BUOYANT OBJECT PROCEDURE . 72
6.4 EQUIPMENT AND SUPPLIES 74
6.5 LITERATURE CITED . 74
7 PHYSICAL HABITAT CHARACTERIZATION 77
7.1 COMPONENTS OF THE HABITAT CHARACTERIZATION 79
7.2 HABITAT SAMPLING LOCATIONS WITHIN THE SAMPLING REACH ... 81
7.3 LOGISTICS AND WORK FLOW 81
7.4 THALWEG PROFILE AND LARGE WOODY DEBRIS MEASUREMENTS 84
7.4.1 Thalweg Profile 84
7.4.2 Large Woody Debris Tally 92
XI
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TABLE OF CONTENTS (CONTINUED)
Section
Page
7.5 CHANNEL AND RIPARIAN CROSS-SECTION MEASUREMENTS 94
7.5.1 Slope and Bearing • 94
7.5.2 Substrate and Channel Dimensions ,... 99
7.5.3 Bank Characteristics 1°1
7.5.4 Canopy Cover Measurements . 105
7.5.5 Riparian Vegetation Structure ; • 109
7.5.6 Instream Fish Cover, Algae, Aquatic Macrophytes .. „• 112
7.5.7 Human Influence 114
7.6 EQUIPMENT AND SUPPLIES • 114
7.7 LITERATURE CITED • 117
8 PERIPHYTON • • • 119
8.1 SAMPLE COLLECTION • - - • • 119
8.2 PREPARATION OF LABORATORY SAMPLES 122
8.2.1 ID/Enumeration Sample 122
8.2.2 Chlorophyll Sample 124
8.2.3 Biomass Sample 128
8.2.4 Acid/Alkaline Phosphatase Activity Sample • 128
8.3 EQUIPMENT AND SUPPLIES 131
8.4 LITERATURE CITED 131
9 SEDIMENT COMMUNITY METABOLISM 133
9.1 SAMPLE COLLECTION 133
9.2 DETERMINING SEDIMENT RESPIRATION 135
9.3 EQUIPMENT AND SUPPLIES 135
10 SEDIMENT TOXICITY 141
10.1 SAMPLE COLLECTION AND PREPARATION 141
10.2 EQUIPMENT AND SUPPLIES 141
11 BENTHIC MACROINVERTEBRATES 147
11.1 SAMPLE COLLECTION 149
11.2 SAMPLE PROCESSING , , 155
11.3 EQUIPMENT AND SUPPLY CHECKLIST 155
XII
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TABLE OF CONTENTS (CONTINUED)
Section Page
11.4 LITERATURE CITED 158
12 AQUATIC VERTEBRATES 161
12.1 SAMPLE COLLECTION 161
12.1.1 Electrofishing 163
12.1.2 Seining 167
12.2 SAMPLE PROCESSING . . . 169
12.2.1 Taxonomic Identification and Tally 169
12.2.2 External Examination and Length Measurements 172
12.2.3 Preparing Voucher Specimens 175
12.3 EQUIPMENT AND SUPPLIES . . 180
12.4 LITERATURE CITED 180
13 FISH TISSUE CONTAMINANTS 183
13.1 PREPARING COMPOSITE SAMPLES FOR PRIMARY AND SECONDARY
TARGET SPECIES ,183
13.2 EQUIPMENT AND SUPPLIES . . . 189
14 RAPID HABITAT AND VISUAL STREAM ASSESSMENTS 193
14.1 RAPID HABITAT ASSESSMENT .......... 193
14.2 VISUAL STREAM ASSESSMENT 194
14.3 EQUIPMENT AND SUPPLIES 208
14.4 LITERATURE CITED 208
15 FINAL SITE ACTIVITIES 211
Appendix Page
A EQUIPMENT AND SUPPLY CHECKLISTS A-1
B QUICK REFERENCE GUIDES B-1
C FIELD DATA FORMS C-1
D SPECIES CODES FOR AQUATIC VERTEBRATES: MID-ATLANTIC REGION . . D-1
E MODIFIED PROTOCOL FOR COLLECTING BENTHIC
MACROINVERTEBRATES E-1
xiii
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FIGURES
Figure
Page
2-1. General sequence of stream sampling activities 19
3-1. Activities conducted at base locations 28
3-2. Performance test procedure for a dissolved oxygen meter. 31
3-3. Sample container labels 37
3-4. Equipment and supply checklist for base location activities 43
4-1. Verification Form (page 1) 47
4-2. Verification Form (page 2) 51
4-3. Sampling reach features 52
4-4. Equipment and supplies checklist for initial site activities 55
5-1. Completed sample labels for water chemistry 59
5-2. Sample Collection Form (page 2), showing data recorded for water
chemistry samples 61
5-3. Field Measurement Form (page 1), showing data recorded for water chemistry. ... 63
5-4. Checklist of equipment and supplies for water chemistry 64
6-1. Layout of channel cross-section for obtaining discharge data by the velocity-area
procedure 68
6-2. Field Measurement Form (page 2), showing data recorded for all three discharge
measurement procedures 71
6-3. Use of a portable weir in conjunction with a calibrated bucket to obtain an estimate
of stream discharge 72
6-4. Equipment and supply checklist for stream discharge .. 76
7-1. Sampling reach layout for physical habitat measurements (plan view). 82
7-2. Thaiweg Profile and Woody Debris Form, 87
7-3. Large woody debris influence zones. 94
XIV
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FIGURES (CONTINUED)
Figure
Page
7-4. Channel slope and bearing measurements. .96
7-5. Slope and Bearing Form 93
7-6. Substrate sampling cross-section. 101
7-7. Channel/Riparian Cross-section and Thalweg Profile Form 103
7-8. Schematic showing bankfull channel and incision for channels 106
7-9. Schematic of modified convex spherical canopy densiometer 107
7-10. Boundaries for visual estimation of riparian vegetation, fish cover, and
human influences 110
7-11. Checklist of equipment and supplies for physical habitat 116
8-1. Index sampling design for periphyton 120
8-2. Sample Collection Form (pagel) showing data recorded for periphyton samples. . 123
8-3. Completed set of periphyton sample labels 124
8-4. Filtration apparatus for preparing chlorophyll and biomass subsamples
for periphyton 127
8-5. Checklist of equipment and supplies for periphyton 132
9-1. Field Measurement Form (page 1), showing data for sediment metabolism
samples ... 133
9-2. Completed sample labels for sediment metabolism. 139
9-3. Checklist of equipment and supplies for sediment metabolism. 140
10-1. Completed sample label for sediment toxicity 143
10-2. Sample Collection Form (page 2), showing information recorded for a sediment
toxicity sample 144
10-3. Checklist of equipment and supplies for sediment toxicity 145
11-1. Modified kick net 148
11-2. Index sampling design for benthic macroinvertebrates 150
11-3. Sample Collection Form (page 1), showing information for benthic
macroinvertebrate samples 154
11-4. Checklist for benthic macroinvertebrate sampling activities 155
11-5. Completed labels for benthic macroinvertebrate samples 158
11-6. Blank labels for benthic invertebrate samples 159
xv
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FIGURES (CONTINUED)
Figure
Page
11-7. Equipment and supply checklist for benthic macroinvertebrates, 160
12-1. Index sample design for aquatic vertebrate sampling. , 162
12-2. Vertebrate Collection Form (pagel) 166
12-3. Vertebrate Collection Form (page 2) , 171
12-4. Fish length measurements 174
12-5. Vertebrate Length Recording Form (page 1) 176
12-6. Completed voucher sample label and specimen bag tag for aquatic
vertebrates 178
12-7. Equipment and supplies checklist for aquatic vertebrates 181
13-1. Completed sample labels for fish tissue contaminants 189
13-2. Sample Collection Form showing information recorded for fish tissue samples. .. 190
13-3. Equipment and supplies checklist for fish tissue contaminants 191
14-1. Rapid Habitat Assessment Form for riffle/run prevalent streams (page 1) 199
14-2. Rapid Habitat Assessment Form for riffle/run prevalent streams (page 2) 200
14-3. Rapid Habitat Assessment Form for pool/glide prevalent streams (page 1) 201
14-4. Rapid Habitat Assessment Form for glide/pool prevalent streams (page 2) 202
14-5. Assessment Form (page 1) 206
14-6. Assessment Form (page 2) 207
14-7. Checklist of equipment and supplies required for rapid habitat and visual
stream assessments 209
XVI
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TABLES
Table
Page
2-1. ESTIMATED TIMES AND DIVISION OF LABOR FOR FIELD ACTIVITIES 18
2-2. GUIDELINES FOR RECORDING FIELD DATA AND OTHER INFORMATION 21
2-3. GENERAL HEALTH AND SAFETY CONSIDERATIONS 23
2-4. GENERAL SAFETY GUIDELINES FOR FIELD OPERATIONS , 25
3-1. STOCK SOLUTIONS, USES, AND INSTRUCTIONS FOR PREPARATION 33
3-2. PERFORMANCE CHECK OF CONDUCTIVITY PENS OR CONDUCTIVITY
METERS . . 34
3-3. GENERAL PERFORMANCE CHECKS FOR CURRENT VELOCITY METERS .... 35
3-4. EQUIPMENT CARE AFTER EACH STREAM VISIT 38
3-5. GENERAL GUIDELINES FOR PACKING AND SHIPPING SAMPLES 40
4-1. SITE VERIFICATION PROCEDURES 46
4-2. GUIDELINES TO DETERMINE THE INFLUENCE OF RAIN EVENTS 49
4-3. LAYING OUT THE SAMPLING REACH 50
4-4. MODIFICATIONS FOR DRY CHANNELS AND INTERMITTENT STREAMS 54
5-1. SAMPLE COLLECTION PROCEDURES FOR WATER CHEMISTRY 60
5-2. PROCEDURES FOR STREAMSIDE AND IN SITU CHEMISTRY
MEASUREMENTS 62
6-1. VELOCITY-AREA PROCEDURE FOR DETERMINING STREAM DISCHARGE ... 69
6-2. TIMED FILLING PROCEDURE FOR DETERMINING STREAM DISCHARGE 73
6-3. NEUTRALLY BUOYANT OBJECT PROCEDURE FOR DETERMINING
STREAM DISCHARGE 75
7-1. COMPONENTS OF PHYSICAL HABITAT CHARACTERIZATION 80
7-2. THALWEG PROFILE PROCEDURE 85
7-3. CHANNEL UNIT AND POOL FORMING CATEGORIES 89
7-4. PROCEDURE FOR TALLYING LARGE WOODY DEBRIS 93
xvii
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TABLES (CONTINUED)
Table
Page
7-5. PROCEDURE FOR OBTAINING SLOPE AND BEARING DATA 97
7-6. SUBSTRATE MEASUREMENT PROCEDURE 102
7-7. PROCEDURE FOR MEASURING BANK CHARACTERISTICS 104
7-8. PROCEDURE FOR CANOPY COVER MEASUREMENTS 108
7-9. PROCEDURE FOR CHARACTERIZING RIPARIAN VEGETATION
STRUCTURE 111
7-10. PROCEDURE FOR ESTIMATING INSTREAM FISH COVER 113
7-11. PROCEDURE FOR ESTIMATING HUMAN INFLUENCE 115
8-1. PROCEDURE FOR COLLECTING COMPOSITE INDEX SAMPLES
OF PERIPHYTON 121
8-2. PREPARATION OF ID/ENUMERATION SAMPLES FOR PERIPHYTON ........ 125
8-3. PROCEDURE FOR PREPARING CHLOROPHYLL SAMPLES FOR
PERIPHYTON 126
8-4. PROCEDURE FOR PREPARING BIOMASS SAMPLES FOR PERIPHYTON 129
8-5. PROCEDURE FOR PREPARING ACID/ALKALINE PHOSPHATASE ACTIVITY
SAMPLES FOR PERIPHYTON 130
9-1. SEDIMENT COLLECTION PROCEDURE 134
9-2. PROCEDURE TO MEASURE SEDIMENT RESPIRATION 136
10-1. PROCEDURE FOR PREPARING SEDIMENT TOXICITY SAMPLES 142
11-1. PROCEDURE TO COLLECT KICK NET SAMPLES FROM RIFFLE AND
RUN HABITATS 151
11-2. PROCEDURE TO COLLECT KICK NET SAMPLES FROM POOL AND
GLIDE HABITATS 153
11-3. PROCEDURE FOR PREPARING COMPOSITE SAMPLES FOR BENTHIC
MACROINVERTEBRATES 157
12-1. PROCEDURE TO COLLECT AQUATIC VERTEBRATES BY
ELECTROFISHING 165
12-2. PROCEDURES TO COLLECT AQUATIC VERTEBRATES BY SEINING ...... 168
XVIII
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TABLES (CONTINUED)
Table
Page
12-3. PROCEDURE TO IDENTIFY, TALLY, AND EXAMINE AQUATIC
VERTEBRATES 170
12-4. EXTERNAL ANOMALY CATEGORIES AND CODES .173
12-5. GUIDELINES AND PROCEDURES FOR PREPARING AQUATIC
VERTEBRATE VOUCHER SPECIMENS . . 177
13-1. PROCEDURE TO PREPARE THE PRIMARY COMPOSITE SAMPLE FOR
FISH TISSUE CONTAMINANTS 185
13-2. PROCEDURE TO PREPARE THE SECONDARY COMPOSITE SAMPLE
FOR FISH TISSUE CONTAMINANTS 187
14-1. DESCRIPTIONS OF HABITAT PARAMETERS USED IN THE RAPID
ASSESSMENT OF STREAMS . . 195
14-2. PROCEDURE FOR CONDUCTING THE RAPID HABITAT ASSESSMENT ..... 198
14-3. PROCEDURE FOR CONDUCTING THE FINAL VISUAL ASSESSMENT
OF A STREAM ; 204
XIX
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ACKNOWLEDGMENTS
Review comments from the following persons are gratefully acknowledged: D.J.
Chaloud, (National Exposure Research Laboratory, Las Vegas, NV), P.A. Lewis (U.S. EPA,
retired), W. Thoeny (SoBran, Inc., Cincinnati, OH), P.M. Nolan (U.S. EPA Region 1, Lexing-
ton, MA), H. R. Preston, (U.S. EPA Region 3, Wheeling, WV), R.D. Spear, (U.S. EPA
Region 2, Edison, NJ), A. Euresti (EPA Region 6, Houston, TX), M.D. Bilger (U.S. Geologi-
cal Survey, Lemoyne, PA), C. Yoder and M. Smith (Ohio EPA, Columbus, OH), and C.
McFarlane (U.S. EPA, Corvallis, OR). The efforts and dedication of numerous field
personnel in implementing these protocols and providing feedback for clarification and
improvement are also recognized. M. Hails-Avery and H. Gronemyer (National Asian
Pacific Center on Aging, Senior Environmental Employment Program, Corvallis, OR)
assisted with preparing many of the figures. G. Mosher (OAO Inc., Corvallis, OR) prepared
the field data forms.
xx
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ACRONYMS, ABBREVIATIONS, AND MEASUREMENT UNITS
Acronyms and Abbreviations
AFDM Ash-free dry mass
APA Acid/Alkaline Phosphatase Activity
BPJ Best Professional Judgment
BOD Biological Oxygen Demand
CENR (White House) Committee on the Environment and Natural Resources
CFR Code of Federal Regulations
DC Direct Current
DIG Dissolved Inorganic Carbon
DLGs Digital Line Graphs
DO Dissolved oxygen
EERD Ecological Exposure Research Division
EMAP Environmental Monitoring and Assessment Program
EMAP-SW Environmental Monitoring and Assessment Program-Surface Waters
Resource Group
EPA U.S. Environmental Protection Agency
ERB Ecosystems Research Branch
GPS Global Positioning System
ID identification
LWD Large Woody Debris
MAHA Mid-Atlantic Highlands Assessment
MAIA Mid-Atlantic Integrated Assessment
NAWQA National Water-Quality Assessment Program
NERL National Exposure Research Laboratory
NHEERL National Health and Environmental Effects Research Laboratory
ORD Office of Research and Development
OSHA Occupational Safety and Health Administration
P-Hab physical habitat
PVC polyvinyl chloride
QA quality assurance
QC quality control
XXI
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ACRONYMS, ABBREVIATIONS, AND MEASUREMENT UNITS
(CONTINUED)
Acronyms and Abbreviations (continured)
RBP (EPA) Rapid Bioassessment Pf&tocol
R-EMAP Regional Environmental Monitoring and Assessment Program
SL Standard length
SOP Standard Operating Procedure
TIME Temporally Integrated Monitoring of Ecosystems
TL Total length
USGS United States Geological Survey
WED Western Ecology Division
YOY young of year
YSI Yellow Springs Instrument system
Measurement Units
amps amperes
cm centimeter
gal gallon
ha hectare
Hz Hertz
in inches
L liter
m meter
m2 square meters
mg/L milligram per liter
mm millimeter
um micrometer
uS/cm microsiemens per centimeter
msec millisecond
ppm parts per million
psi pounds per square inch
V volts
VA volt-ampere
xxii
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SECTION 1
INTRODUCTION
by
James M. Lazorchak1, Alan T. Herlihy2, H. Ronald Preston3-4 and Donald J. Klemm1
This manual contains procedures for collecting samples and measurement data
from various biotic and abiotic components of streams. These procedures were developed
and used between 1993 and 1998 in research studies of the U.S. Environmental Protection
Agency's (EPA) Environmental Monitoring and Assessment Program (EMAP). The pur-
poses of this manual are to: (1) Document the procedures used in the collection of field
data and various types of samples for the various research studies; and (2) provide these
procedures for use by other groups implementing stream monitoring programs.
These procedures are designed for use during a one-day visit by a crew of four
persons to sampling sites located on smaller, wadeable streams (stream order 1 through 3).
They were initially developed based on information gained from a workshop of academic,
State, and Federal experts (Hughes, 1993), and subsequent discussions between aquatic
biologists and ecologists within EMAP, with scientists of the U.S. Geological Survey Na-
tional Water Quality Assessment Program (NAWQA), with biologists from the U.S. Fish &
Wildlife Service, and with State and Regional biologists within EPA Region 3.
EMAP initiated additional research activities in 1997 to develop field procedures for
use in nonwadeable riverine systems. These procedures are currently still under develop-
ment and will be published separately.
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin L. King Dr.,
Cincinnati, OH 45268.
2 Department of Fisheries and Wildlife, Oregon State University, c/o U.S. EPA. 200 SW 35th St., Corvallis, OR 97333
3 U.S. EPA Region 3, Wheeling Office, 303 Methodist Bldg, 11th and Chaplin Streets, Wheeling, WV 26003.
4 Current address: Canaan Valley Institute, P.O. Box 673, Davis, WV 26260
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EMAP-SW-Streams Held Operations Manual. Section 1 (Introduction), Rev. 4, September 1998 Page 2 of 16
1.1 OVERVIEW OF EMAP-SURFACE WATERS
The U.S. EPA has designated EMAP to develop the necessary monitoring tools to
determine the current status, extent, changes and trends in the condition of our nation's
ecological resources on regional and national scales (U.S. EPA, 1998). The nation's ecolog-
ical resources are a national heritage, as essential to the country now and in the future as
they have been in the past. Data indicate that regional and international environmental
problems may be endangering these essential resources. The potential threats include acid
rain, ozone depletion, point and nonpoint sources of pollution, and climate change.
The tools being developed by EMAP include appropriate indicators of ecological
condition, and statistical sampling designs to determine the status and extent of condition,
and to detect regional-scale trends in condition. When fully implemented in a national
monitoring framework, such as that being developed by the White House Committee on
Environment and Natural Resources (CENR; Committee on Environment and Natural
Resources, 1997), these tools will provide environmental decision makers with statistically
valid interpretive reports describing the health of our nation's ecosystems (Whittier and
Paulsen, 1992). Knowledge of the health of our ecosystems will give decision makers and
resource managers the ability to make informed decisions, set rational priorities, and make
known to the public costs, benefits, and risks of proceeding or refraining from implementing
specific environmental regulatory actions. Ecological status and trend data will allow deci-
sion makers to objectively assess whether or not the nation's ecological resources are
responding positively, negatively, or not at all, to existing or future regulatory programs.
The following three objectives guide EMAP research activities (U.S. EPA, 1998):
Estimate the current status, extent, changes and trends in indicators of
the condition of the nation's ecological resources on a regional basis
with known confidence.
Monitor indicators of pollutant exposure and habitat condition and
seek associations between human-induced stresses and ecological
condition.
Provide periodic statistical summaries and interpretive reports on
ecological status and trends to resource managers and the public.
The EMAP Surface Waters Resource Group (EMAP-SW) is charged with developing
the appropriate tools to assess the health of lakes, streams, and wetlands in the United
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 3 of 16
States. The first phase of the program started with a study of northeastern lakes between
1991 and 1996 (Larsen and Christie, 1993; Baker et ai., 1997). In 1992 and 1993, a pilot
study of wetland ecosystems was conducted in the Prairie Pothole region of the northern
plains region of the U.S. (Peterson et al., 1997). The specific research studies dealing with
streams are described in more detail in the following section.
1.2 STREAM SAMPLING COMPONENTS OF EMAP-SURFACE WATERS
The procedures presented in this manual were developed and refined during several
different research projects conducted between 1993 and 1997. These projects represent
two types of field activities to be performed prior to full-scale implementation of a monitoring
program that addresses EMAP objectives. Pilot projects are intended to answer questions
about proposed ecological indicators, such as plot design (how to obtain representative
samples and data from each stream site), responsiveness to various stressors, evaluation
of alternative methods, and logistical constraints. Pilot studies are not primarily intended to
provide regional estimates of condition, but may provide these estimates for a few indica-
tors.
Demonstration projects are conducted at larger geographic scales, and may be
designed to answer many of the same questions as pilot studies. Additional objectives of
these larger studies are related to characterizing spatial and temporal variability of ecologi-
cal indicators, and to demonstrating the ability of a suite of ecological indicators to estimate
the condition of regional populations of aquatic resources.
1.2.1 Mid-Atlantic Highlands Assessment Project
The stream sampling component of EMAP-SW was initiated in 1993 in the mid-
Appalachian region of the eastern United States, in conjunction with a Regional-EMAP (R-
EMAP) project being conducted by EPA Region 3. This R-EMAP study was known as the
Mid-Atlantic Highlands Assessment study (MAHA), and was carried out over a 4-year pe-
riod. The MAHA project was designed to test the EMAP approach in a few of the most
heavily impacted ecoregions of Region 3, the mid-Appalachians, the Ridge and Valley, the
Central Appalachians, the Piedmont and some of the Coastal Plain.
The Region 3 R-EMAP project was designed to answer the following questions:
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 4 of 16
• What are biological reference conditions for the Central Appalachian Ridge
and Valley Ecoregion?
• Do biological communities differ between subregions?
• What is the status of mid-Atlantic Highlands stream biota?
Can linkages be established between impairment and possible causes of
impairment?
How can an EMAP-like approach be used to design programs to restore and
manage stream resources on a regional scale?
During the MAHA study, 577 wadeable stream sites throughout EPA Region 3 (DE,
MD, VA, WV, PA) and the Catskill Mts. of New York were visited and sampled using the
field protocols being developed by EMAP. Streams were sampled each year during a 10-
week index period from April to July by field crews from EPA, the U.S. Fish and Wildlife
Service, State, and contract personnel.
1.2.2 Mid-Atlantic Integrated Assessment Program
In 1997 and 1998 the EMAP Surface Waters Program became a collaborator in the
Mid-Atlantic Integrated Assessment (MAIA) project, which is attempting to produce an
assessment of the condition of surface water and estuarine resources. The MAIA project
represented a follow-up to the MAHA study, with an expanded geographic scope (southern
New York to northern North Carolina, with more sites located in the Piedmont and Coastal
Plain ecoregions) and a different index period (July-September). The first year of the MAIA
study, approximately 200 sites (150 wadeable sites, 13 repeated wadeable sites, and ap-
proximately 30 riverine sites) were visited for sampling.
1.2.3 Temporal Integrated Monitoring of Ecosystems Project
A special interest component of EMAP-SW is the Temporal Integrated Monitoring of
Ecosystems Project (TIME). The purpose of the TIME project is to assess the changes and
trends in chemical condition in acid-sensitive surface waters (lakes and streams) of the
northeastern and eastern U.S. resulting from changes in acidic deposition caused by the
1990 Clean Air Act Amendments. The TIME project has three goals (Stoddard, 1990):
1. Monitor current status and trends in chemical indicators of acidification
in acid-sensitive regions of the U.S.
2. Relate changes in deposition to changes in surface water conditions.
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 5 of 16
3. Assess the effectiveness of the Clean Air Act emissions reductions in
improving the acid/base status of surface waters.
1.2.4 Other Projects
The basic procedures and methods presented in this manual have also been used in
other areas of the U.S. as part of R-EMAP projects being conducted by other EPA Regions.
These include Regions 7 (central U.S.), 8 (Colorado), 9 (California), and 10 (Oregon and
Washington). Each of these projects have modified the basic procedures to be compatible
with the geographic region or other project-specific requirements.
1.3 SUMMARY OF ECOLOGICAL INDICATORS
The following sections describe the rationale for each of the ecological indicators
currently included in the stream sampling procedures presented in this manual. Evaluation
activities to determine the suitability of individual indicators to robustly determine ecological
condition are ongoing at this time. This information is presented to help users understand
the various field procedures and the significance of certain aspects of the methodologies.
Currently, EMAP considers two principal types of indicators, condition and stressor
(U.S. EPA, 1998). Condition indicators are biotic or abiotic characteristics of an ecosystem
that can provide an estimate of the condition of an ecological resource with respect to some
environmental value, such as biotic integrity. Stressor indicators are characteristics that are
expected to change the condition of a resource if the intensity or magnitude is altered.
1.3.1 Water Chemistry
Data are collected from each stream for a variety of physical and chemical constitu-
ents. Information from these analyses is used to evaluate stream condition with respect to
stressors such as acidic deposition (of importance to the TIME project), nutrient enrichment,
and other inorganic contaminants. In addition, streams can be classified with respect to
water chemistry type, water clarity, mass balance budgets of constituents, temperature
regime, and presence of anoxic conditions.
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4. September 1998 Page 6 of 16
1.3.2 Physical Habitat
Naturally occurring differences among surface waters in physical habitat structure
and associated hydraulic characteristics contributes to much of the observed variation in
species composition and abundance within a zoogeographic province. The structural
complexity of aquatic habitats provides the variety of physical and chemical conditions to
support diverse biotic assemblages .and maintain long-term stability. Anthropogenic alter-
ations of riparian areas and stream channels, wetland drainage, grazing and agricultural
practices, and stream bank modifications such as revetments or development, generally
act to reduce the complexity of aquatic habitat and result in a loss of species and ecosys-
tem degradation.
Stressor indicators derived from data collected about physical habitat quality will be
used to help explain or diagnose stream condition relative to various condition indicators.
Important attributes of physical habitat in streams are channel dimensions, gradient, sub-
strate characteristics; habitat complexity and cover; riparian vegetation cover and structure;
disturbance due to human activity, and channel-riparian interaction (Kaufmann, 1993).
Overall objectives for this indicator are to develop quantitative and reproducible indices,
using both multivariate and multimetric approaches, to classify streams and to monitor
biologically relevant changes in habitat quality and intensity of disturbance. Kaufmann et al.
(in preparation) discuss procedures for reducing EMAP field habitat measurements and
observations to metrics that describe channel and riparian habitat at the reach scale.
1.3.3 Periphyton Assemblage
Periphyton are the algae, fungi, bacteria, and protozoa associated with substrates in
aquatic habitats. These organisms exhibit high diversity and are a major component in
energy flow and nutrient cycling in aquatic ecosystems. Many characteristics of periphyton
community structure and function can be used to develop indicators of ecological conditions
in streams. Periphyton are sensitive to many environmental conditions, which can be
detected by changes in species composition, cell density, ash free dry mass (AFDM),
chlorophyll, and enzyme activity (e.g., alkaline and acid phosphatase). Each of these
characteristics may be used, singly or in concert, to assess condition with respect to soci-
etal values such as biological integrity and trophic condition.
A hierarchical framework is being used in the development of the periphyton indices
of stream condition. The framework involves the calculation of composite indices for biotic
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 7 of 16
integrity, ecological sustainability, and trophic condition. The composite indices will be
calculated from measured or derived first-order and second-order indices. The first-order
indices include species composition (richness, diversity), cell density, AFDM, chlorophyll,
and enzyme activity (e.g., Saylor et al., 1979), which individually are indicators of ecological
condition in streams. Second-order indices will be calculated from periphyton characteris-
tics, such as the autotrophic index (Weber, 1973), community similarity compared to refer-
ence sites, and autecological indices (e.g., Lowe, 1974; Lange-Bertalot, 1979; Charles,
1985; Dixitetal, 1992).
1.3.4 Sediment Community Metabolism
Ecosystems are complex, self-regulating, functional units defined by rates and
processes, such as energy flow or material cycling. These processes are mediated by the
trophic structure of the ecosystem, and integrate the functioning of the entire community.
Energy flow and material cycling are important components of two major concepts in stream
ecology: The river continuum concept and resource spiraling. Heterotrophic microorgan-
isms (bacteria and fungi) are responsible for oxygen sags in streams and for much of the
decomposition of organic matter deposited in them. Measuring the rate of oxygen con-
sumption within the soft sediments of a stream provides a functional indicator of energy flow
and material transformation within the ecosystem
1.3.5 Benthic Macroinvertebrate Assemblage
Benthic macroinvertebrates inhabit the sediment or live on the bottom substrates of
streams. The macroinvertebrate assemblages in streams reflect overall biological integrity
of the benthic community , and monitoring these assemblages is useful in assessing the
status of the water body and discerning trends. Benthic communities respond differently to
a wide array of stressors. As a result of this, it is often possible to determine the type of
stress that has affected a benthic macroinvertebrate community (Plafkin et al., 1989; Klemm
et al., 1990). Because many macroinvertebrates have relatively long life cycles of a year or
more and are relatively immobile, macroinvertebrate community structure is a function of
past conditions.
Two different approaches are currently being evaluated to developing ecological
indicators based on benthic invertebrate assemblages. The first is a multimetric approach,
where different structural and functional attributes of the assemblage are characterized as
"metrics". Individual metrics that respond to different types of stressors are scored against
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 8 of 16
expectations under conditions of minimal human disturbance. The individual metric scores
are then summed into an overall index value that is used to judge the overall level of impair-
ment of an individual stream reach. Examples of multimetric indices based on benthic
invertebrate assemblages include Kerans and Karr (1993), Fore et al. (1996) and Barbour
etal. (1995; 1996).
The second approach being investigated is to develop indicators of condition based
on multivariate analysis of benthic assemblages and associated abiotic variables. Exam-
ples of this type of approach as applied to benthic invertebrate assemblages include
RIVPACS (Wright, 1995), and BEAST (Reynoldson et al., 1995). Rosenberg and Resh
(1993) present various approaches to biological monitoring using benthic invertebrates, and
Norris (1995) briefly summarizes and discusses approaches to analyzing benthic macro-
invertebrate community data.
1.3.6 Aquatic Vertebrate Assemblages
Aquatic vertebrate assemblages of interest to EMAP include fish and amphibians.
The fish assemblage represents a critical component of biological integrity from both an
ecosystem function and a public interest perspective. Historically, fish assemblages have
been used for biological monitoring in streams more often than in lakes (e.g., Plafkin et al.,
1989; Karr, 1991). Fish assemblages can serve as good indicators of ecological conditions
because fish are long-lived and mobile, forage at different trophic levels, integrate effects of
lower trophic levels, and are reasonably easy to identify in the field (Plafkin et al., 1989).
Amphibians comprise a substantial portion of vertebrate biomass in streams of many areas
of the U.S. (Hairston, 1987; Bury et al., 1991). Reports of dramatic declines in amphibian
biodiversity (e.g., Blaustein and Wake, 1990; Phillips, 1990) has increased the level of
interest in monitoring these assemblages. Amphibians may also provide more information
about ecosystem condition in headwater or intermittent streams in certain areas of the
country than other biological response indicators (Hughes, 1993). The objective of field
sampling is to collect a representative sample of the aquatic vertebrate assemblage by
methods designed to 1) collect all except very rare species in the assemblage and 2) pro-
vide a measure of the abundance of species in the assemblages (McCormick, 1993).
Information collected for EMAP that is related to vertebrate assemblages in streams in-
cludes assemblage attributes (e.g., species composition and relative abundance) and the
incidence of external pathological conditions.
8
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 9 of 16
Indicators based on vertebrate assemblages are being developed primarily using the
multimetric approach described in Section 1.3.5 for benthic macroinvertebrates, and origi-
nally conceived by Karr and others (Karr et al., 1986). Simon and Lyons (1995) provide a
recent review of multimetric indicators as applied to stream fish assemblages.
1.3.7 Fish Tissue Contaminants
Indicators of fish tissue contaminants attempt to provide measures of bioaccumula-
tion of toxic chemicals in fish. When coupled with study designs such as those being
developed by EMAP, these indicators can be used to estimate regional risks Of consumption
to predators of fish (either wildlife or human), and to track how this risk changes with time
in a region. It is also meant to be used in conjunction with the other stressor indicators
(physical habitat, water chemistry, land use, population density, other records of relevant
anthropogenic stresses) and condition indicators (fish, macroinvertebrates, periphyton) to
help diagnose whether the probable cause of stream degradation, when it is shown by the
condition indicators to occur, is water quality, physical habitat, or both.
The various studies that have been done on fish tissue contaminants have focused
on different parts of the fish: whole fish, fillets, livers. For EMAP-SW, the focus is on
whole fish because of the emphasis on the ecological health of the whole stream (as op-
posed to a focus on human health concerns). Whole fish are a better indicator of risk to
piscivorous wildlife than fillets. It is hoped to also be able to say something about risks to
human health by analyzing whole fish. Whole fish also present fewer logistical problems for
field crews (no gutting required in the field) and the analytical lab (no filleting necessary).
Samples are prepared for two major categories of fish species. One sample is
prepared using a species whose adults are small (e.g., small minnows, sculpins, or darters).
The second sample is prepared using a species whose adults are of larger size (e.g.,
suckers, bass, trout, sunfish, carp). In addition to being more ubiquitous than the larger fish
(and therefore more likely to be present in sufficient numbers to composite), small fish have
other advantages over large fish. Most importantly, it may be possible to get a more repre-
sentative sample of the contaminant load in that stream segment (although it could be at a
lower level of bioaccumulation) by creating a composite sample from a larger number of
small individuals than by compositing a few individuals of larger species. Small fish may be
a more appropriate indicator for assessing ecological risk, as they might be expected to be
prey for a larger number of fish-eating animals (the majority of which will be piscivorous
birds and small mammals). The major advantage that larger fish could potentially offer,
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EMAP-SW-Streams Field Operations Manual. Section 1 (Introduction), Rev. 4. September 1998 Page 10 of 16
whether predators (piscivores) or bottom feeders, is a higher level of bioaccumulation and
thus greater sensitivity to detect contaminants. The relative bioaccumulation of contami-
nants by large and small stream fish is not known, thus the reason for preparing two sam-
ples in this study.
1.3.8 Sediment Toxicity
Sediment toxicity testing has been used to evaluate the contaminant levels of fresh-
water harbors and rivers, as well as estuaries, marine bays, and marsh lands. Most of its
use in the past has been in evaluating sites that were known or suspected to be highly
contaminated. EMAP-SW is the first program to use sediment toxicity on such a large scale
in freshwater lakes and streams. Sediment toxicity tests, using the freshwater amphipod
Hyalella azteca, will be used to determine the status of sediment contamination in streams.
Sediment toxicity can also be used to indicate the affects of non-contaminant stressors,
such as physical habitat degradation. The measurements for sediment toxicity are simple
and easy to determine. The survival in each sample is determined at the end of the test
and compared to survival in a test using a "reference" sediment.
1.4 OBJECTIVES AND SCOPE OF THE FIELD OPERATIONS AND METHODS
MANUAL
Only field-related sampling and data collection activities are presented in this man-
ual. Laboratory procedures and methods (including sample processing and analytical
methods) associated with each ecological indicator are summarized in Chaloud and Peck
(1994); detailed procedures will be published as a separate document.
**,
This manual is organized to follow the sequence of field activities during the 1-day
site visit. Section 2 presents a general overview of all field activities. Section 3 presents
those procedures that are conducted at a "base" location before and after a stream site
visit. Section 4 presents the procedures for verifying the site location and defining a reach
of the stream where subsequent sampling and data collection activities are conducted.
Sections 5 through 14 describes the procedures for collecting samples and field measure-
ment data for various condition and stressor indicators. Specific procedures associated with
each indicator are presented in standalone tables that can be copied, laminated, and taken
into the field for quick reference. Section 15 describes the final activities that are conducted
before leaving a stream site. Appendix A contains a list of all equipment and supplies
required by a crew to complete all field activities at a stream. Appendix B presents a set of
10
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 11 of 16
brief summaries of field procedures and activities that can be laminated, collated into a 3-
ring binder, and taken into the field along with the procedure tables. This waterproof hand-
book can serve as the primary field reference for field teams after they complete an inten-
sive training program. Appendix C provides a complete set of blank field data forms as
used in 1997. Appendix D contains a list of vertebrate species names and corresponding
species codes developed for use in the Mid-Atlantic region. This information documents the
common and scientific names used for the various Mid-Atlantic studies, and also provides
an example that can be adapted for use in other areas of the country. Appendix E presents
a modified protocol for collecting benthic macroinvertebrates that has been used in EMAP
studies in some parts of the U.S.
Depending on the specific project and approach to information management, field
teams may also be provided with an information management handbook that contains
instructions for tracking samples and generating sampling status reports as well as using
the computers and associated hardware and software. Field teams are also required to
keep the field operations and methods manual available in the field for reference and to
address questions pertaining to protocols that might arise.
1.5 QUALITY ASSURANCE
Large-scale and/or long-term monitoring programs such as those envisioned for
EMAP require a rigorous quality assurance (QA) program that can be implemented consis-
tently by all participants throughout the duration of the monitoring period. Quality assurance
is a required element of all EPA-sponsored studies that involve the collection of environ-
mental data (Stanley and Verner, 1986). Field teams should be provided a copy of the QA
project plan (e.g., Chaloud and Peck, 1994 for EMAP-SW activities). The QA plan contains
more detailed information regarding QA/QC activities and procedures associated with
general field operations, sample collection, measurement data collection for specific indica-
tors, and data reporting activities.
Quality control (QC) activities associated with field operations are integrated into the
field procedures. Important QA activities associated with field operations include a compre-
hensive training program that includes practice sampling visits, and the use of a qualified
museum facility or laboratory to confirm any field identifications of biological specimens.
The overall sampling design for EMAP-SW related studies usually includes a subset of sites
(10 to 15 percent) that are revisited within a single sampling period and/or across years
(e.g., Larsen, 1997; Urquhart et al., 1998). Information from these repeat visits is used in
11
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EMAP-SW-Streatns Field Operations Manual. Section 1 (Introduction), Rev. 4, September 1998 Page 12 of 16
part to describe overall sampling and measurement precision for the various ecological
indicators.
1.6 LITERATURE CITED
Barbour, M.T., J.B. Stribling, and J.R. Karr. 1995. The multimetric approach for establish-
ing biocriteria and measuring biological condition, pp. 69-80 IN: W.S. Davis and T.P.
Simon (eds.) Biological Assessment and Criteria: Tools for Water Resource Planning
and Decision-making. Lewis Publishers, Chelsea, Michigan.
Barbour, M.T., J. Gerritsen, G.E. Griffith, R. Frydenborg, E. McCarron, J.S. White, and M.L.
Bastian. 1996. A framework for biological criteria for Florida streams using benthic
macroinvertebrates. Journal of the North American Benthological Society 15(2): 185-
211.
Baker, J.R., D.V. Peck, and D.W. Sutton (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, D.C.
Blaustein, A.R. and D.B. Wake. 1990. Declining amphibian populations: a global phenom-
enon? Trends in Ecology and Evolution 5:203-204.
Bury, R.B., P.C. Corn, K.B. Autry, F.F. Gilbert, and L.L.C. Jones. 1991. Aquatic amphibian
communities in Oregon and Washington, pp. 353-362 ]N: L.F. Ruggiero, K.B. Aubry,
A.B. Carey, and M.H. Huff (coordinators). Wildlife and Vegetation ofUnmanaged
Douglas-Fir Forests. General Technical Report PNW-GRT-285. USDA Forest Ser-
vice, Portland, Oregon.
Chaloud, D. J., and D. V. Peck (eds.). 1994. Environmental Monitoring and Assessment
Program: Integrated Quality Assurance Project Plan for the Surface Waters Resource
Group. EPA 600/X-917080. Revision 2.00. U.S. Environmental Protection Agency,
Las Vegas, Nevada.
Charles, D.F. 1985. Relationships between surface sediment diatom assemblages and
lakewater characteristics in Adirondack lakes. Ecology 66:994-1011.
12
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 13 of 16
Committee on Environment and Natural Resources. 1997. Integrating the Nation's
Environmental Monitoring and research Networks and Programs: A Proposed Frame-
work. March 1997 revision. Office of Science and Technology Policy, Washington,
DC.
Dixit, S.S., J.P. Smol, J.C. Kingston, and D.F. Charles. 1992. Diatoms: Powerful indicators
of environmental change. Environmental Science and Technology 26:22-33.
Fore, L.S., J.R. Karr, and R.W. Wissernan. 1996. Assessing invertebrate responses to
human activities, evaluating alternative approaches. Journal of the North American
Benthological Society 15:212-231.
Hairston, N.G. 1987. Community Ecology and Salamander Guilds. Cambridge University
Press.
Hughes, R.M. (ed.). 1993. Stream Indicator and Design Workshop. EPA/600/R-93/138.
U.S. Environmental Protection Agency, Corvallis, Oregon.
Karr, J.R. 1991. Biological integrity: a long neglected aspect of water resource manage-
ment. Ecological Applications 1:66-84.
Karr, J.R., K.D. Fausch, P.L. Angermeier, P.R. Yant, and I.J. Schlosser. 1986. Assessing
Biological Integrity in Running Waters: A Method and its Rationale. Illinois Natural
History Survey Special Publication 5. Champaign, IL
Kaufmann, P.R. (ed.). 1993. Physical Habitat, pp. 59-69 ]N: R.M. Hughes (ed.). Stream
Indicator and Design Workshop. EPA/600/R-93/138. U.S. Environmental Protection
Agency, Corvallis, Oregon.
Kaufmann, P.R., P. Levine, E.G. Robison, C. Seeliger, and D.V. Peck. In preparation.
Quantifying Physical Habitat in Wadeable Streams. Environmental Monitoring and
Assessment Program, U.S. Environmental Protection Agency, Corvallis, Oregon.
Kerans, B.L., and J.R. Karr. 1994. A benthic index of biotic integrity (B-IBI) for rivers of the
Tennessee Valley. Ecological Applications 4:768-785.
13
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 14 of 16
Klemm, D.J., P.A. Lewis, F. Fulk, J.M. Lazorchak. 1990. Macroinvertebrate Field and
Laboratory Methods for Evaluating the Biological Integrity of Surface Waters.
EPA/600/4-90/030. U.S. Environmental Protection Agency, Cincinnati, Ohio.
Larsen, D.P. and S.J. Christie (eds.) 1993. EMAP-Surface Waters 1991 Pilot Report.
EPA/620/R-93/003. U.S. Environmental Protection Agency, Washington, D.C.
Lange-Bertalot, H. 1979. Pollution tolerance of diatoms as criterion for water quality estima-
tion. NovaHedwigia 64:285-304.
Larsen, D.P. 1997. Sample survey design issues for bioassessment of inland aquatic
ecosystems. Human and Ecological Risk Assessment 3:979-991.
Lowe, R.L. 1974. Environmental Requirements and Pollution Tolerance of Freshwater
Diatoms. U.S. Environmental Protection Agency, Environmental Monitoring Series,
National Environmental Research Center, Cincinnati, Ohio.
McCormick, F.H. 1993. Fish. pp. 29-36 IN: R.M. Hughes (ed.). Stream Indicator Work-
shop. EPA/600/R-93/138. U.S. Environmental Protection Agency. Corvallis, Oregon.
Norris, R.H. 1995. Biological monitoring: the dilemma of data analysis. Journal of the
North American Benthological Society 14:440-450.
Peterson, S.A., L. Carpenter, G. Gutenspergen, and L.M. Cowardin (editors). 1997. Pilot
Test of Wetland Condition Indicators in the Prairie Pothole Region of the United States.
EPA/620/R-97/002. U.S. Environmental Protection Agency, Washington, D.C.
Phillips, K. 1990. Where have all the frogs and toads gone? Bioscience 40:422-424.
Plafkin, J.L., M.T. Barbour, K.D. Porter, S.K. Gross, and R.M. Hughes. 1989. Rapid Bio-
assessment Protocols for Use in Streams and Rivers: Benthic Macroinvertebrates and
Fish. EPA/440/4-89/001. U.S. Environmental Protection Agency, Washington, D.C.
Reynoldson, T.B., R.C. Bailey, K.E. Day, and R.H. Norris, 1995. Biological guidelines for
freshwater sediment based on Benthic Assessment Sediment (the BEAST) using a
multivariate approach for predicting biological state. Australian Journal of Ecology
20:198-219.
14
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EMAP-SW-Streams Field Operations Manual. Section 1 (Introduction). Rev. 4, September 1998 Page 15 of 16
Rosenberg, D.M. and V.H. Resh. 1993. Freshwater Biomonitoring and Benthic Macro-
invertebrtates. Chapman and Hall, New York.
Sayler; G.S., M. Puziss, and M. Silver. 1979. Alkaline phosphatase assay for freshwater-
sediments: application to perturbed sediment systems. Applied and Environmental
Microbiology 38:922-927.
Simon, T.P. and J. Lyons. 1995. Application of the index of biotic integrity to evaluate
water resources integrity in freshwater ecosystems, pp. 245-262 JIN: W.S. Davis and
T.P. Simon (eds.), Biological Assessment and Criteria: Tools for Water Resource
Planning and Decision Making. Lewis Publishers, Boca Raton, Florida.
Stanley, T.W., and S.S. Verner. 1986. The U.S. Environmental Protections Agency's
quality assurance program, pp. 12-19 IN: J.K. Taylor and T.W. Stanley (eds.). Quality
Assurance for Environmental Measurements. ASTM STP 867, American Society for
Testing and Materials, Philadelphia, Pennsylvania.
Stoddard, J.L. 1990. Plan for Converting NAPAP Aquatic Effects Long-Term Monitoring
(LTM) Project to the Temporally Integrated Monitoring of Ecosystems (TIME) Project.
U.S. Environmental Protection Agency, Environmental Research Laboratory, Corvallis,
Oregon.
Urquhart, N.S., S.G. Paulsen, and D.P. Larsen. 1998. Monitoring for policy-relevant re-
gional trends over time. Ecological Applications 8:246-257'.
U.S. EPA. 1998. Environmental Monitoring and Assessment Program (EMAP): Research
Plan 1997. EPA/620/R-98/002. U.S. Environmental Protection Agency, Washington,
D.C.
Weber, C.I. 1973. Recent developments in the measurement of the response of plankton
and periphyton to changes in their environment, pp. 119-138 ]N: G. Glass (ed.),
Bioassay Techniques and Environmental Chemistry. Ann Arbor Science Publishers,
Ann Arbor, Michigan.
Whittier, T.R. and S.G. Paulsen. 1992. The surface waters component of the Environmen-
tal Monitoring and Assessment Program (EMAP): an overview. Journal of Aquatic
Ecosystem Health 1:119-126.
15
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EMAP-SW-Streams Field Operations Manual, Section 1 (Introduction), Rev. 4, September 1998 Page 16 of 16
Wright, J.F. 1995. Development and use of a system for predicting the macroinvertebrate
fauna in flowing waters. Australian Journal of Ecology 20:181-197.
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SECTION 2
OVERVIEW OF FIELD OPERATIONS
Brian H. Hill1, Frank H. McCormick1, James M. Lazorchak1, Donald J. Klemm1,
Philip A. Lewis1'2, Victoria C. Rogers3-4, and Michael K. McDowell3
This section presents a general overview of the activities a 4-person field team
conducts during a typical one-day sampling visit to a stream site. General guidelines for
recording data and using standardized field data forms and sample labels are also pre-
sented. Finally, safety and health considerations and guidelines related to field operations
are provided.
2.1 DAILY OPERATIONAL SCENARIO
The field team is divided into two groups, termed the "Geomorphs" and the "Bio-
morphs," that reflect their initial responsibilities more than their expertise. The geomorphs
are primarily responsible for conducting the intensive physical habitat characterization. The
biomorphs are primarily responsible for collecting biological samples. Table 2-1 provides
the estimated time required to conduct various field activities. Figure 2-1 presents the
general sequence of activities conducted at each stream reach.
Upon arrival at a stream site, the geomorphs are responsible for verifying and docu-
menting the site location, determining the length of stream reach to be sampled, and estab-
lishing the required transects (Section 4). The biomorphs collect samples and field mea-
surements for water chemistry (Section 5) and determine stream discharge (Section 6).
The biomorphs also collect sediment for the sediment metabolism determination (Section 9)
1 U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 Martin L. King Dr.,
Cincinnati, OH 45268.
2 Current address: 1037 Wylie Road, RR #2, Seaman, OH 45679.
3 OAO, Inc., 200 SW 35th St., Corvallis, OR 97333.
4 Current address: Linn-Benton Community College.6500 SW Pacific Blvd., Albany, OR 97321.
17
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EMAP-SW-Streams Field Operations Manual. Section 2 (Overview of Field Operations), Rev. 3, September 1998 Page 2 of 10
TABLE 2-1. ESTIMATED TIMES AND DIVISION OF LABOR FOR FIELD ACTIVITIES
Activity
Site verification and establishing sampling reach
and transects
Water chemistry sampling and stream discharge
determination
Collecting and processing benthos, periphyton
and sediment metabolism samples
Intensive physical habitat characterization
Aquatic vertebrate sampling and processing
Rapid habitat assessment
Visual stream assessment
Sample tracking and packing
SUMMARY
Group
Geomorphs (2 persons)
Biomorphs (2 persons)
Biomorphs (2 persons)
Geomorphs (2 pesons)
Geomorphs and
Biomorphs (4 persons)
Biomorphs (2 persons)
Geomorphs (2 persons)
28 to 32 person-hours
Est. Time
Required
2 hours
1 hour
3.5 hours
2 to 3 hours
2 to 5 hours
0.5 hours
1 hour
7 to 8 hours
per team
and sediment toxicity testing (Section 10), and collect periphyton and benthos samples
(Sections 8 and 11, respectively). The geomorphs conduct the intensive physical habitat
characterization (Section 7). Both groups are involved with collecting aquatic vertebrates
(Section 12) and preparing samples for fish tissue contaminants (Section 13). Finally, the
biomorphs conduct a habitat characterization based on the Rapid Bioassessment Protocols
(RBP; Plafkin et al., 1989) and a visual stream assessment (Section 14), while the geo-
morphs prepare samples for transport and shipment (Section 3).
2.2 GUIDELINES FOR RECORDING DATA AND INFORMATION
During the one-day visit to a stream, a field team is required to obtain and record a
substantial amount of data and other information for all of the various ecological indicators
described in Section 1.3. In addition, all the associated information for each sample col-
lected must be recorded on labels and field data forms to ensure accurate tracking and
subsequent linkage of other data with the results of sample analyses.
It is imperative that field and sample information be recorded accurately, consis-
tently, and legibly. Measurement data that cannot be accurately interpreted by others
18
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EMAP-SW-Streams Field Operations Manual, Section 2 (Overview of Field Operations), Rev. 3, September 1998 Page 3 of 10
CGEOMORPHS"N
(2 persons) J
CBIOMORPHS"^\
(2 persons) /
SITE LOCATION AND VERIFICATION
•Verify Stream and reach locations
• Mark Index site and habitat transects
WATER CHEMISTRY
Collect samples
Conduct field measurements
PHYSICAL HABITAT CHARACTERIZATION
(Intensive)
Thalweg profile measurements
Substrate size and channel dimensions
Large woody debris tally
Riparian vegetation types and structure
Canopy density
Bank characteristics
Instream fish cover
Human disturbance
STREAM DISCHARGE
STREAM PERIPHYTON
• Collect and prepare
composite samles
BENTHIC
MACROtNVERTEBRATES
• Collect kick net samples
and prepare composite
samples
SEDIMENT METABOLISM AND
SEDIMENT TOXICITY
Collect composite sediment sample
]
AQUATIC VERTEBRATE ASSEMBLAGE
FISH TISSUE CONTAMINANTS
Conduct electrofishing and seining
ID and tally vertebrates collected; length measurements and external anomaly
examination
Prepare voucher and "unknown" specimens
Select specimens and prepare tissue samples
PHYSICAL HABITAT QUALITY
(Rapid)
Conduct RBP habitat characterization
Complete visual stream assessment
SEDIMENT METABOLISM AND SEDIMENT TOXIC1TY |
• Prepare incubation chamber .
- Prepare replicate metabolism samples
• Prepare sediment toxicity sample
* Measure O2 of replicates after incubation
• Prepare biomass samples
STREAM PERIPHYTON
Prepare ID/Enumeration sample
Prepare chlorophyll sample
Prepare biomass sample
Prepare activity sample
FINAL ACTIVITIES
Review field data forms
Inspect and package samples
Clean up stream site
BENTHIC
MACROINVERTEBRATES
Process composite samples
I
NEXT DAY ACTIVITIES
• Ship samples and data forms
• Travel to next stream
Figure 2-1. General sequence of stream sampling activities (modified from Chaloud and
Peck, 1994).
19
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EMAP-SW-Streams Field Operations Manual, Section 2 (Overview of Field Operations), Rev. 3, September 1998 Page 4 of 10
besides the field teams, and/or samples with incorrect or illegible information associated
with them, are lost to the program. The cost of a sampling visit coupled with the short index
period severely limits the ability to re-sample a stream because the initial information re-
corded was inaccurate or illegible. Some guidelines to assist field personnel with recording
information are presented in Table 2-2. Examples of completed data forms and labels are
presented in the sections describing field sampling and measurement procedures for differ-
ent indicators, and a complete set of blank field data forms are included as Appendix C.
2.3 SAFETY AND HEALTH
Collection and analysis of samples (e.g., benthic invertebrates, fish, periphyton,
sediment) can involve significant risks to personal safety and health (drowning, electrical
shock, pathogens, etc.). While safety is often not considered an integral part of field sam-
pling routines, personnel must be aware of unsafe working conditions, hazards connected
with the operation of sampling gear, boats, and other risks (Berry et al., 1983). Personnel
safety and health are of the highest priority for ail investigative activities and must be em-
phasized in safety and health plans for field, laboratory, and materials handling operations.
Preventive safety measures and emergency actions must be emphasized. Management
should assign health and safety responsibilities and establish a program for training in
safety, accident reporting, and medical and first aid treatment. Safety documents and
standard operating procedures (SOPs) containing necessary and specific safety precau-
tions should be available to all field personnel. Additional sources of information regarding
field and laboratory safety related to biomonitoring studies include Berry et al. (1983), U.S.
EPA (1986) and Ohio EPA (1990).
2.3.1 General Considerations
Important considerations related to field safety are presented in Table 2-3. It is the
responsibility of the group safety officer or project leader to ensure that the necessary
safety courses are taken by all field personnel and that all safety policies and procedures
are followed. Sources of information regarding safety-related training include the American
Red Cross (1989), the National Institute for Occupational Safety and Health (1981), U.S.
Coast Guard (1987) and Ohio EPA (1990).
Persons using sampling devices should become familiar with the hazards involved
and establish appropriate safety practices prior to using them. Individuals involved in
electrofishing must be trained by a person experienced in this method or by attending a
20
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EMAP-SW-Streams Field Operations Manual, Section 2 (Overview of Field Operations), Rev. 3. September 1998 Page 5 of 10
TABLE 2-2. GUIDELINES FOR RECORDING FIELD DATA AND OTHER INFORMATION
Activity
Guidelines
Field Measurements:
Data Recording
Record measurement values and/or observations on data forms preprinted
on water-resistant paper.
Record information on forms using No. 2 pencil only. Erase mistakes com-
pletely and write the correct value whenever you can. If you must
line out an incorrect value, place the correct value nearby so the data
entry operator can easily find it.
Headers on the second pages of all forms link the data. Fill in all headers
of all pages or data will be lost (this is a good one to review at the
end of the day).
Record data and information so that all entries are obvious. Enter data
completely in every field that you use. Follow the "comb" guidelines-
print each number or letter in the individual space provided. Keep
letters and numerals from overlapping. Record data to the number of
decimal places provided on the forms. Illegible information is equiva-
lent to no information.
Print neatly, using block capital letters in alphabetical fields. Clearly distin-
guish letters from numbers (e.g., 0 versus O, 2 versus Z, 7 versus T
or F, etc.). Do not put lines through 7's,p's, or Z's. Do not use
slashes.
Record information on each line, even if it has to be recorded repeatedly
on a series of lines (e.g., fish species codes or physical habitat char-
acteristics). Do not use "ditto marks" (") or a straight vertical line.
When recording comments, print or write legibly. Make notations in com-
ments field only. Avoid marginal notes, etc. Be concise, but avoid
using abbreviations and/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 and/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 team dur-
ing a particular sampling visit (also used for qualifying sam-
ples).
Explain all flags in comments section on data form.
Review of Data
Forms
Field team reviews data forms for accuracy, completeness, and legibility
before leaving a stream.
Data forms from all teams are reviewed for completeness, accuracy, and
legibility before transfer to the information management staff.
(continued)
21
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EMAP-SW-Streams Field Operations Manual. Section 2 (Overview of Field Operations), Rev. 3, September 1998 Page 6 of 10
TABLE 2-2 (Continued)
Activity
Guidelines
Sample Collection and Tracking
Sample Labels
Use adhesive labels with preprinted ID numbers and a standard recording
format for each type of sample.
Record information on labels using a fine-point indelible marker. Cover com-
pleted labels with clear tape.
Sample Collection
Information
Record sample ID number from the label and associated collection informa-
tion on sample collection form preprinted on water-resistant paper.
Record information on field data forms using No. 2 pencil only (fine-point
indelible fine-tipped markers can be used if necessary).
Record collection information using correct format as provided on the collec-
tion form.
Sample Qualifiers
(Flags)
Use only defined flag codes and record on sample collection form in appropri-
ate field.
K Sample not collected or lost before shipment; re-sampling not
possible.
U Suspect sample (e.g., possible contamination, does not meet mini-
mum acceptability requirements, or collected using a nonstandard
procedure)
Fn Miscellaneous flags (n=1, 2, etc.) assigned by a field team during
a particular sampling visit (also used for field measurements).
Explain all flags in comments section on sample collection form.
Review of Labels
and Collection
Forms
The field team compares information recorded on labels and sample collec-
tion form for accuracy before leaving a stream.
The field team reviews labels and collection form for accuracy, completeness,
and legibility before leaving a stream.
Sample collection forms are reviewed for completeness, accuracy, and legibil-
ity before transfer to the information management staff.
If boats are used to access sampling sites, personnel must consider and prepare for
hazards associated with the operation of motor vehicles, boats, winches, tools, and other
incidental equipment. Boat operators should be familiar with U.S. Coast Guard rules and
regulations for safe boating contained in a pamphlet, "Federal Requirements for Recre-
ational Boats," available from a local U.S. Coast Guard Director or Auxiliary or State Boat-
ing Official (U.S. Coast Guard, 1987). All boats with motors must have fire extinguishers,
boat horns, life jackets or flotation cushions, and flares or communication devices.
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
22
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EMAP-SW-Streams Field Operations Manual, Section 2 (Overview of Field Operations), Rev. 3. September 1998 Page 7 of 10
TABLE 2-3. GENERAL HEALTH AND SAFETY CONSIDERATIONS
Training:
First aid
Cardiopulmonary resuscitation (CPR)
Vehicle safety (e.g., operation of 4-wheel drive vehicles)
Boating and water safety (if boats are required to access sites)
Field safety (e.g., weather conditions, personal safety, orienteering, reconnaissance of sites
prior to sampling
Equipment design, operation, and maintenance
Electrofishing safety
Handling of chemicals and other hazardous materials
Communications
Check-in schedule
Sampling itinerary (vehicle used and its description, time of departure, travel route, esti-
mated time of return)
Contacts for police, ambulance, fire departments, search and rescue personnel
Emergency services available near each sampling site and base location
Personal Safety .
Field clothing and other protective gear
Medical and personal information (allergies, personal health conditions)
Personal contacts (family, telephone numbers, etc.)
Physical exams and immunizations
certified electrofishing training course. Reynolds (1983) and Ohio EPA (1990) provide
additional information regarding electrofishing safety procedures and practices, have a daily
check-in procedure for safety. An emergency communications plan should include contacts
for police, ambulance, fire departments, and search and rescue personnel.
Proper field clothing should be worn to prevent hypothermia, heat exhaustion, sun-
stroke, drowning, or other dangers. Field personnel should be able to swim. Chest waders
made of rubberized or neoprene material and suitable footwear must always be worn with a
belt to prevent them from filling with water in case of a fall. The use of a life jacket is advis-
able at dangerous wading stations if one is not a strong swimmer because of the possibility
of sliding into deep water.
23
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EMAP-SW-Streams Field Operations Manual, Section 2 (Overview of Field Operations), Rev. 3. September 1998 Page 8 of 10
Many hazards lie out of sight in the bottoms of lakes, 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 sus-
pected 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.
Prior to a sampling trip, personnel should determine that all necessary equipment is
in safe working condition. Good housekeeping practice should be followed in the field.
These practices protect staff from injury, prevent or reduce exposure to hazardous or toxic
substances, and prevent damage to equipment and subsequent down time and/or loss of
valid data.
2.3.2 Safety Equipment and Facilities
Appropriate safety apparel such as waders, lab coats, gloves, safety glasses, etc.
must be available and used when necessary. Bright colored caps (e.g., orange) must be
available and worn during field activities. First aid kits, fire extinguishers, and blankets must
be readily available in the field. A properly installed and operating fume hood must be
provided in the laboratory for use when working with carcinogenic"chemicals (e.g., formalde-
hyde, formalin) that may produce dangerous fumes. Cellular telephones or portable radios
should be provided to field teams working in remote areas for use in case of an emergency.
Facilities and supplies must be available for cleaning of exposed body parts that may have
been contaminated by pollutants in the water. Soap and an adequate supply of clean water
or ethyl alcohol, or equivalent, should be suitable for this purpose.
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 on a regular or infrequent basis should be in sound physical
condition and have a physical exam annually or in accordance with Regional, State, or
organizational requirements. All surface waters and sediments should be considered
potential health hazards due to toxic substances or pathogens. Persons must become
familiar with the health hazards associated with using chemical fixing and/or preserving
agents. Formaldehyde (or formalin) is highly allergenic, toxic, and dangerous to human
24
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EMAP-SW-Streams Field Operations Manual, Section 2 (Overview of Field Operations), Rev. 3, September 1998 Page 9 of 10
TABLE 2-4. GENERAL SAFETY GUIDELINES FOR FIELD OPERATIONS
• Two persons (three to four persons for electrofishing) must be present during all sample
collection activities, and no one should be left alone while in the field.
• Exposure to stream 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 and when turning over rocks during hand picking.
• Use appropriate protective equipment (e.g., gloves, safety glasses) when handling and
using hazardous chemicals
• Persons working in areas where poisonous snakes may be encountered 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.
If local advice is not available and medical assistance is more than an hour away,
carry a snake bite kit and be familiar with its use.
• Any person allergic to bee stings, other insect bites, or plants must take proper precautions
and have any needed medications handy.
• Field personnel should also protect themselves against the bite of deer or wood ticks be-
cause of the potential risk of acquiring pathogens that cause Rocky Mountain spotted fever
and Lyme disease.
• All 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 freez-
ing (up to 10°C or 50°F) if he or she is exposed to wind or becomes wet.
• Handle and dispose of chemical wastes properly. Do not dispose any chemicals in the field.
health (carcinogenic) if utilized improperly. Chemical wastes can cause various hazards
due to flammability, explosiveness, toxicity, causticity, or chemical reactivity. All chemical
wastes must be discarded according to standardized health and hazards procedures (e.g.,
National Institute for Occupational Safety and Health [1981]; U.S. EPA [1986]). .
25
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EMAP-SW-Streams Field Operations Manual. Section 2 (Overview of Field Operations). Rev. 3, September 1998 Page 10 of 10
2.4 LITERATURE CITED
American Red Cross. 1979. Standard First Aid and Personal Safety. American National
Red Cross. 269 pp.
Berry, C.R. Jr., W.T. Helm, and J. M. Neuhold. 1983. Safety in fishery field work. pp. 43-
60 1M: Nielsen, L.A., and D. L. Johnson (eds.). Fisheries Techniques. American
Fisheries Society, Bethesda, MD.
Chaloud, D. J., and D. V. Peck (eds.). 1994 Environmental Monitoring and Assessment
Program: Integrated Quality Assurance Project Plan for the Surface Waters Resource
Group. 'EPA 600/X-917080. Revision 2.00. U.S. Environmental Protection Agency,
Las Vegas, Nevada.
National Institute for Occupational Safety and Health. 1981. Occupational Health Guide-
lines for Chemical Hazards (Two Volumes). NIOSH/OSHA Publication No. 81-123.
U.S. Government Printing Office, Washington, D.C.
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.
Plafkin, J.L., M.T. Barbour, K.D. Porter, S.K. Gross, and R.M. Hughes. 1989. Rapid
Bioassessment Protocols for Use in Streams and Rivers: Benthic Macroinvertebrates
and Fish. EPA/440/4-89/001. U.S. Environmental Protection Agency, Washington,
D.C.
Reynolds, J. B. 1983. Electrofishing. pp. 147-163. ]N: L. A. Nielsen and D. L Johnson
(eds.). Fisheries Techniques. American Fisheries Society, Bethesda, MD.
U.S. Coast Guard. 1987. Federal Requirements for Recreational Boats. U.S. Department
of Transportation, United States Coast Guard, Washington, D.C.
U.S. EPA. 1986. Occupational Health and Safety Manual. Office of Planning and Manage-
ment, U.S. Environmental Protection Agency, Washington, D.C.
26
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SECTIONS
BASE LOCATION ACTIVITIES
Donald J. Klemm1, Brian H. Hill1, Frank H. McCormick1, and Michael K. McDowell2
Field teams conduct a number of activities at a "base" location before and after
visiting each stream site. These activities are generally conducted on the same day as the
sampling visit. Close attention to these activities is required to ensure that the field teams
know where they are going, that access to the stream site is possible and permissible, that
all the necessary equipment and supplies are in good order to complete the sampling effort,
and that samples are packaged and shipped correctly and promptly.
Figure 3-1 illustrates operations and activities that are conducted before and after
each visit to a stream site. Activities that are conducted after a stream visit include equip-
ment cleanup and maintenance, packing and shipping samples, and communications with
project management to report the status of the visit.
3.1 ACTIVITIES BEFORE EACH STREAM VISIT
Before each stream visit, each field team should confirm access to the stream site,
develop a sampling itinerary, inspect and repair equipment, check to make sure all supplies
required for the visit are available, and prepare sample containers. Procedures to accom-
plish these activities are described in the following sections.
3.1.1 Confirming Site Access
Field crews should be provided with dossiers containing important locational and
access information for each stream they are scheduled to visit. Before visiting a stream, the
crew should review the contents of the specific stream dossier. The landowner(s) listed in
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin L. King Dr.,
Cincinnati, OH.
2 OAO, Inc., c/o U.S. EPA, 200 SW 35th St., Corvallis, OR 97333
27
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities). Rev. 0, September 1998 Page 2 of 18
BASE LOCATION ACTIVITIES
BEFORE EACH STREAM VISIT
Team Leader
Review stream dossier
information
Make access contacts
Prepare itinerary
Team Members
Test and calibrate oxygen meter
and conductivity pen or meter
Initialize GPS (if necessary)
Prepare sample containers and
labels
Pack equipment and supplies using
checklist
AFTER EACH STREAM VISIT
Team Leader
Review forms and labels
Enter sample tracking information into computer
Package and ship samples and data forms
File status report with field coordinator or other
central contact person
Team Members
Clean and check equipment
Charge or replace batteries
Check and refuel vehicles ~
Obtain ice and other consumable supplies as
needed
Figure 3-1. Activities conducted at base locations.
28
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0, September 1998 Page 3 of 18
the dossier should be contacted to confirm permission to sample and identify any revisions
to the information contained in the dossier.
3.1.2 Daily Sampling Itinerary
Based upon the sampling schedule provided to each team, team leaders are respon-
sible for developing daily itineraries. The team leader reviews each stream dossier to
ensure that it contains the appropriate maps, contact information, copies of permission
letters, and access instructions. Additional activities include determining the best access
routes, calling the landowners or local contacts to confirm permission, confirming lodging
plans for the upcoming evening, and coordinating rendezvous locations with individuals who
must meet with field teams prior to accessing a site. This information is used to develop an
itinerary for the stream. The itinerary should include anticipated departure time, routes of
travel, location of any intermediate stops (e.g., to drop off samples, pick up supplies, etc.)
and estimated time of arrival at the final destination after completing the stream visit. This
information (and any changes that occur due to unforeseen circumstances), should be
provided to the field coordinator or other central contact person identified for the specific
field study. Failure to adhere to the reported itinerary can result in the initiation of expensive
search and rescue procedures and disruption of carefully planned schedules. In addition,
each team should carry individual emergency medical and personal information with them,
possibly in the form of a "safety log" that remains in the vehicle (see Section 2).
3.1.3 Instrument Inspections and Performance Tests
Each field team is required to test and calibrate instruments prior to departure for the
stream site. Field instruments include a global positioning system (GPS) receiver, a current
velocity meter, a conductivity pen (or a conductivity meter), and a dissolved oxygen meter.
Backup instruments should be available if instruments fail the performance tests or calibra-
tions described in the following subsections.
3.1.3.1 Global Positioning System Receiver—
Specific performance checks will vary among different brands of GPS receivers.
Follow the instructions in the receiver's operating manual to make sure the unit is function-
ing properly. Turn on the receiver and check the batteries. Replace batteries immediately if
a battery warning is displayed. Make sure extra batteries are stored with the receiver and
will be available in the field.if necessary.
29
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0. September 1998 Page 4 of 18
Before the initial use, or, in some cases, if batteries are replaced, the receiver may
require inputting the coordinates of a positional reference point that is nearby (e.g., a U.S.
Geological Survey benchmark identified on a topographic map). Follow the manufacturer's
instructions for initializing the receiver.
3.1.3.2 Dissolved Oxygen Meter—
As an initial performance test before use each year, dissolved oxygen (DO) meters
should be tested for accuracy against the Winkler titration method, In addition, inspect and
test the dissolved oxygen meters at the base location before each stream site visit. The
inspection and testing procedure, based on the use of Yellow Springs Instruments (YSI)
Model 53 oxygen meters, is summarized in Figure 3-2. Some modification to the procedure
may be necessary for other models or types of dissolved oxygen meters.
Inspect the meter by checking the status of the batteries, and the functioning of the
electronics. Confirm the meter is adjusted correctly for measurements in fresh water.
Inspect the membrane of the probe. If bubbles are present, if the membrane is discolored,
or if the membrane is torn, use a backup probe and/or replace the membrane on the origi-
nal probe. (NOTE: For older models of meters, new membranes may require conditioning
for 24 hours before use).
After inspecting the meter and probe, attempt to calibrate it, following the instruc-
tions in the instrument operating manual. Do not record the calibration information obtained
during the performance test. The meter is calibrated again at each stream site, at which
time the calibration information is recorded on the field data form. If the meter cannot be
successfully calibrated, replace the meter and/or probe. After the test, turn the meter off,
and store the probe according to the manufacturer's instructions.
3.1.3.3 Conductivity Pens or Conductivity Meters--
If conductivity "pens" are being used, check the pen for outward signs of fouling
daily. Refer to the instrument manual for probe cleaning instructions. Do not touch the
electrodes inside the probe with any object. Always keep the pen's electrode moist by
keeping deionized water in the pen cap. If deionized water is not available, use stream-
water or tap water rather than let the electrode dry out. Before using a pen which has been
stored dry, soak the electrodes in deionized water (by filling the caps) for 24 hours. If
conductivity meters are used, follow the operating manual provided with the instrument to
check the batteries, the electronics, and to inspect the probe. New probes or probes that
have been stored dry may require conditioning before use.
30
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities). Rev. 0, September 1998 Page 5 of 18
DISSOLVED OXYGEN METER PERFORMANCE CHECK
Replace
batteries
or meter
CHECK METER
Turn meter on
Adjust electronic zero
Adjust salinity knob (0-FRESH)
EQUILIBRATE PROBE
Empty water from calibration
chamber
Insert probe into calibration chamber
Equilibrate in calibration chamber
water bath for 15 minutes
FAIL
Temperature
Check
(within ±1 °C)?
PASS
CALIBRATE METER
Adjust meter to theoretical O2 value for water-
saturated air at chamber temperature and pressure
I
Successful \ NO
vCalibration?^
YES
TAKE METER TO THE STREAM
Refill calibration chamber with water
to store probe
Replace probe
and/or meter
Figure 3-2. Performance test procedure for a dissolved oxygen meter.
31
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EMAP-SW-Streams Field Operations Manual. Section 3 (Base Location Activities), Rev. 0, September 1998 Page 6 of 18
The operation of the conductivity pen or conductivity meter is checked at the base
location using a standard solution of known conductivity. A daily quality control check
sample (QCCS) is prepared as described in Table 3-1. The daily QCCS can be prepared
as either of two dilutions of the stock standard, depending on the theoretical conductivity
desired. A 1:100 dilution of the stock provides a QCCS with a conductivity of 75.3 uS/cm at
25 °C (Metcalf and Peck, 1993). A 1:200 dilution results in a QCCS with a conductivity of
37.8 uS/cm at 25 °C (Peck and Metcalf, 1991). A fresh lot of the daily QCCS should be
prepared every two weeks from the stock standard solution. Check the performance of the
conductivity pen or conductivity meter by following the procedure presented in Table 3-2.
3.1.3.4 Current Velocity Meters-
Field teams may be using one of three types of current velocity meters, a vertical
axis meter (e.g., Price type AA), an electromagnetic type meter (e.g., Marsh McBirney
Model 201D), or a photo-optical impeller type meter (e.g., Swoffer Model 2100). General
guidelines regarding performance checks and inspection of current meters are presented in
Table 3-3. Consult the operating manual for the specific meter and modify this information
as necessary.
3.1.4 Preparation of Equipment and Supplies
To ensure that all activities at a stream can be conducted completely and efficiently,
field teams should check all equipment and supplies before traveling to a stream site. In
addition, they should prepare sample containers and labels for use to the extent possible.
Check the inventory of equipment and supplies prior to departure using the stream-
visit checklists presented in Appendix A. Pack meters, probes, and sampling gear in such a
way as to minimize physical shock and vibration during transport. If necessary, prepare
stock preservative solutions as described in Table 3-1. Follow the regulations of the Occu-
pational Safety and Health Administration (OSHA) for handling and transporting hazardous
materials such as formalin and ethanol. Regulations pertaining to formalin are in the Code
of Federal Regulations (CFR; specifically 29 CFR 1910.1048). These requirements should
be summarized for all hazardous materials being used for the project and provided to field
personnel. Transport formalin and ethanol in appropriate containers with absorbent mate-
rial.
32
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities). Rev. 0, September 1998 Page 7 of 18
TABLE 3-1. STOCK SOLUTIONS, USES, AND INSTRUCTIONS FOR PREPARATION
SOLUTION
Bleach
(10%)
Conductivity
Standard
Stock Solution3
Quality Control
Check Sample
Formalin, borax
buffered0
(pH 7-8)
Ethanol
USE
Clean seines, dip nets,
kick nets, or other equip-
ment that is immersed in
the stream
To prepare conductivity
quality control check sam-
ple solution
To check operation of con-
ductivity pen or conductiv-
ity meter
Preservative for fish speci-
mens and periphyton sam-
ples
Preservative for benthic
macroinvertebrate sam-
ples.
PREPARATION
Dilute 400 ml_ chlorine bleach solution to 4 L
with tap water.
Dissolve 3.4022 g KH2PO4 and 3.5490 g
Na2HPO4 (analytical grade; dried at 120 °C for 3
h and stored desiccated) in 1000.0 g (1.0018 L
at 20 °C, 1 .0029 L at 25 °C) reagent water.
1:100 dilution of standard stock solution with
reagent water (theoretical conductivity = 75.3
MS/cm at 25 °C)a
1 :200 dilution of standard stock solution with
reagent water (theoretical conductivity = 37.6
uS/cm at 25 °C)6
Add 400 g borax detergent (e.g., Twenty Mule
Team®) to each 20-L container of 100% forma-
lin. Test with pH paper.
None.
Metcalf and Peck (1993)
b Peck and Metcalf (1991)
c Handle formalin according to 29 CFR 1910.1048.
33
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EMAP-SW-Streams Held Operations Manual, Section 3 (Base Location Activities), Rev. 0. September 1998 Page 8 of 18
TABLE 3-2. PERFORMANCE CHECK OF CONDUCTIVITY PENS OR CONDUCTIVITY METERS
1. Check the functioning of the pen or meter according to the manufacturer's operating manual
(e.g., zero and "red line" of the meter).
2. Swirl the electrodes (pen) or probe (meter) for 3-5 seconds in a 250-mL bottle containing the
daily QCCS solution labeled "RINSE".
3. Transfer the probe from the "RINSE" bottle to a second 250-mL bottle of QCCS labeled
"TEST1. Let stabilize for 20 seconds.
4. If the measured value of the QCCS is within ±10% or ±10 uS/cm of the theoretical value, rinse
the pen or probe in deronized water. Store as described in the operating manual and package
the pen or meter for transport to the stream site.
If the measured value of the QCCS is not within ±10% or ±10 uS/cm of theoretical value,
repeat Steps 1 through 3.
If the value is still unacceptable, replace the QCCS in both the "rinse" and "test"
bottles and repeat the measurement process.
If the measured value is still not acceptable, clean the pen or conductivity probe as
described in the manual, check the batteries, soak in deionized water for 24 hours,
and repeat Steps 1 through 3.
If the measured value is still unacceptable, replace the pen or meter.
34
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0, September 1998 Page 9 of 18
TABLE 3-3. GENERAL PERFORMANCE CHECKS FOR CURRENT VELOCITY METERS
Vertical-axis Meters (from Smoot and Novak, 1968)
• Inspect the bucket and wheel hub assembly, yoke, cups, tailpiece, and the pivot point each
day before use.
• Inspect the bearings and check the contact chamber for proper adjustment.
• Periodically conduct a spin test of the meter. The minimum spin time is 1.5 minutes, while
the recommended time is between 3 and 4 minutes.
Electromagnetic Meters
• Check the meter calibration daily as part of morning routine. Calibration value should be
2.00 + 0.05.
• Once per week, check the zero value using a bucket of quiescent water. Place the probe in
the bucket and allow to sit for 30 minutes with no disturbance. The velocity value obtained
should be 0.0 + 0.1. Adjust the meter zero if the value is outside this range.
Photoelectric Impeller Meters
• Check that the calibration adjustment cover screws are tightly fitted on the display case.
• Periodically check the condition of the connector fitting between the display unit and the
sensor.
• Connect the sensor to the display unit and check the calibration value stored in memory. If
this value is less than the correct value for the display unit-sensor rotor combination, replace
the batteries.
• Periodically perform a spin test of the rotor assembly, following the instructions in the me-
ter's operating manual. A displayed count value of 300 or greater is indicative of satisfac-
tory performance at low current velocities.
• If a buzzing sound occurs when the rotor assembly is spun by hand, or if the shaft shows
visible wear, replace the rotor assembly.
• Periodically examine the thrust-bearing nut on the rotor assembly. If a "cup" begins to form
on the bottom surface of the nut, it should be replaced.
35
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EMAP-SW-Streams Held Operations Manual. Section 3 (Base Location Activities), Rev. 0. September 1998 Page 10 of 18
Inspect the vehicles every morning before departure. Refuel vehicles and conduct
maintenance activities the night before a sampling trip. Check vehicle lights, turn signals,
brake lights, and air pressure in the tires.
Some sample containers can be labeled before departing from the base site. Figure
3-3 illustrates the preprinted labels. A set of three water chemistry sample containers all
having the same ID number (one for the 4-L cubitainer and two for the 60-mL syringes) can
be pre-labeled with the appropriate information (described in Section 5). After labeling,
place the syringes in their plastic container, and place the cubitainer and beakers in a clean
self-sealing plastic bag to prevent contamination. Sample containers for biological and
sediment samples should NOT be pre-labeled before reaching the stream site. Problems in
sample tracking can result if jars are labeled and then are not used at a stream.
3.2 ACTIVITIES AFTER EACH STREAM VISIT
Upon reaching a lodging location after sampling a stream, the team reviews all
completed data forms and sample labels for accuracy, completeness, and legibility, and
makes a final inspection of samples. If information is missing from the forms or labels, the
team leader should fill in the missing information as accurately as possible. The team
leader initials all data forms after review. The other team member should inspect and clean
sampling equipment, check the inventory of supplies, and prepare samples for shipment.
Other activities include shipping samples and communicating with the field coordinator or
other central contact person.
3.2.1 Equipment Care
Equipment cleaning procedures are given in Table 3-4. Inspect all equipment,
including nets, and clean off any plant and animal material. This effort ensures that intro-
ductions of nuisance species do not occur between streams, and prevents possible cross-
contamination of samples. If nets cannot be cleaned thoroughly using water and detergent,
clean and disinfect them with a 10 percent chlorine bleach solution (Table 3-1). Use bleach
only as a last resort, as repeated use will destroy the net material. Take care to avoid
damage to lawns or other property.
36
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0, September 1998 Page 11 of 18
WATER CHEMISTRY
CU SI S2
SITE ID: MAIA -
DATE / /98
22S015
PERIPHYTON
APA MOMASS CMLA ID
SITF O; MWA
DATE: .' .'88
HASH A I': TOCL RIFFLt/RUN
S JBSAhP. P VC1I UHF . . .ml
COMPOSITE VOLUME:
225004
SEDIMENT METABOLISM
SITtlDMAW •
DATE: __/__/98
SAMPLb TWO R1 R2 R5 R4 RS
229003
SEDIMENT TOXICITY
SITE ID: MWA -
DATE: I /98
STATION:
229011
PERIPHYTON
APA WOMASS CMLA ID
SITF ID: MAIA
DATE: i
HASriAtt TOCL
SJBSAKIJ.F VtllUMF
COMPOSITE VOLUME:
,'98
RIFFLC/RUN
.ml
mL
229004
COMPOSITE BENTHOS
SITE ID: MAIA ______
DATE:
_/ I 98
HABITAT: Riffle Pool
FISH TISSUE
SITE ID: MAIA -
DATE: __/__/98
229013
229003
FISH-JAR
SITE ID: MAIA -
DATE: . . /
229003
Figure 3-3. Sample container labels.
3.2.2 Sample Tracking, Packing, and Shipment
Each field team packs and ships samples from each stream visit as soon as possible
after collection, normally the day following a stream visit. Field teams must be provided with
specific information for the shipping destinations, contact persons, and the required ship-
ping schedule for each type of sample.
Sample tracking information (including sample types, sample ID numbers, and other
field-related information that is required by the laboratory to conduct analyses and associate
results to a specific sample and stream site) is recorded during the packing process.
37
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0, September 1998 Page 12 of 18
TABLE 3-4. EQUIPMENT CARE AFTER EACH STREAM VISIT
1. Clean for biological contaminants (e.g., plant and animal material).
• Prior to departing a stream, drain all water from live wells and buckets used to hold and
process fish.
Inspect sampling gear for evidence of plant fragments and remove any fragments ob-
served.
• At the stream or base site, dry out seines, dip nets, and kick nets, and inspect and re-
move any remnant vegetation or animal life. If the weather is rainy and gear cannot be
dried out, then use a different (backup) set of gear, if available. If an additional set "of
gear is not available, disinfect gear with 10 percent bleach solution.
2. Clean and dry other equipment prior to storage.
• Rinse chlorophyll filtration chamber three times with distilled water after each use.
Rinse periphyton sampling equipment with tap water at the base site.
• Rinse coolers with water to clean off any dirt or debris on the outside and inside.
Make sure conductivity pens or conductivity meter probes are rinsed with deionized water
and are stored moist.
• Rinse all beakers used to collect water chemistry samples three times with deionized
water to prevent contamination of the next stream sample. Place the beakers in a 1-
gallon self-sealing plastic bag with a cubitainer for use at the next stream.
3. Check fish nets for holes and repair, if possible; otherwise, set damaged gear aside and locate
replacements.
4. Inventory equipment and supply needs and relay orders to the Field Coordinator through the
Communications Center.
5. Remove DO meters and GPS receivers from carrying cases and set up for pre-visit inspections
and performance tests. Examine the DO membrane for cracks, wrinkles, or bubbles; replace if
necessary.
6. Recharge all batteries overnight if possible (12-V wet cells, current meter, computer battery).
Replace others (GPS, DO meter) as necessary.
7. Recheck field forms from the day's sampling activities. Make corrections and completions
where possible, and initial each form after review.
8. Replenish fuel in vehicles and/or electrofishing generator (if necessary).
38
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0, September 1998 Page 13 of 18
Depending upon the project, this information may be recorded manually onto paper forms,
or otherwise recorded electronically into a portable computer, using such tools as barcode
scanners and customized entry and reporting software. Procedures for conducting sample
tracking activities should be provided to each field team by the information management
staff, possibly as a separate operations manual or handbook.
The sample tracking system should also identify the intermediate and final destina-
tions for each sample. In some cases, intermediate storage "depots" may be used to
accumulate samples prior to shipment to the support laboratory. The tracking system
should provide an informal "chain-of custody" to prevent the loss of samples and associated
information.
General guidelines for packing and shipping the various types of samples described
in this manual are presented in Table 3-5. When shipping samples using ice, use fresh ice.
Use block ice when available; it should be sealed in a large plastic bags. If block ice is not
available, contain the ice in several self-sealing plastic bags. Label each bag of ice as "ICE"
with an indelible marker to prevent any leakage of meltwater from being misidentified by
couriers as a possible hazardous material spill.
Water chemistry samples must be shipped as soon as possible after collection in
order to meet holding time requirements for some laboratory analyses. To ship water
chemistry samples, place a large (30-gallon) plastic bag in an insulated shipping container
(e.g., a plastic or metal cooler). The sample labels on the cubitainer and syringes should be
completely covered with clear tape to prevent damage from water or condensation during
shipment. Place the four syringes into a separate plastic container for shipment. Place the
cubitainer and syringe container into a second large plastic bag and close. Place the bag
containing the samples inside the plastic bag lining the shipping container. Place bags of
ice around the bag of samples, but inside the plastic bag lining the shipping container.
Then close the outer plastic bag. Seal the cooler with clear tape. Place the required sample
tracking forms in the shipping container and close it. Seal the container with shipping tape
and affix any required shipping-related labels to the outside of the container. Attach an
adhesive plastic sleeve to the lid of the container and insert any required shipping forms.
Sediment toxicity samples can be held for extended periods (e.g., a week), if they
can be kept refrigerated in the field. Transport or ship sediment toxicity samples in a sepa-
rate container from water chemistry samples if possible to avoid possible contamination of
39
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EMAP-SW-Streams Field Operations Manual. Section 3 (Base Location Activities). Rev. 0, September 1998 Page 14 of 18
TABLE 3-5. GENERAL GUIDELINES FOR PACKING AND SHIPPING SAMPLES
Sample Type
(container)
Guidelines
Samples requiring refrigeration (4 °C)
Water Chemistry
(4-L cubitainer and 60-mL
syringes)
Ship on day of collection or within 24 hr by overnight courier.
Use fresh ice in labeled plastic bags for shipping.
Line each shipping container with a large plastic bag.
Place syringes in a plastic container.
Place syringe container and cubitainer inside of a second plastic bag.
Cover labels completely with clear tape.
The cubitainer and syringes should have same sample ID number
assigned.
Confirm the sample ID assigned on the labels matches the ID number
recorded on the field collection form (or other sample tracking report).
Sediment Toxicity
(1-gal plastic bag)
Ship on day of collection or within 24 hr by overnight courier. Keep
chilled if extended storage time in the field is necessary.
Use a separate shipping container from water chemistry samples.
Package and ship using the same instructions as for water chemistry
samples.
If available, place the plastic bag containing the sample into a plastic
container to protect it during transport and shipment.
Cover labels completely with clear tape.
Confirm the sample ID assigned on the label matches the ID number
recorded on the field collection form (or other sample tracking report).
Samples requiring freezing (-20 °C) within 24 hours of collection
Periphyton chlorophyll (fil-
ter inaluminurnfoil)
Periphyton biomass (filter
in a numbered container)
Periphyton activity (50-mL
centrifuge tube)
Sediment metabolism
(50-mL centrifuge tubes)
If samples cannot be kept frozen in the field, ship on day of collection
or within 24 h by overnight courier.
Cover the label completely with clear tape.
Protect samples from meltwater if ice is used by double bagging ice
and placing samples in a plastic container.
Confirm the sample ID assigned on the label matches the ID number
recorded on the field collection form (or other sample tracking report).
If dry ice is used to transport or ship samples, special shipping contain-
ers, outside labeling, and shipping forms may be required.
Fish Tissue
(aluminum foil; two 30-gal
plastic bags)
If samples cannot be kept frozen in the field, ship on day of collection
or within 24 h by overnight courier.
Cover labels completely with clear tape.
Label on each bag should have identical Sample ID number assigned.
Confirm the sample ID assigned on the label matches the ID number
recorded on the field collection form (or other sample tracking report).
Protect samples from meltwater if ice is used by double bagging ice.
Special shipping containers, outside labeling, and shipping forms may
be required for shipments containing dry ice.
(continued)
40
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0, September 1998 Page 15 of 18
TABLE 3-5. (Continued)
Sample Type
(container)
Preservative
Guidelines
Samples requiring preservation in formalin
Periphyton ID (50-mL cen-
trifuge tube)
Fish Specimens
(1 -L and/or 4-L jars)
Benthic Macro-
invertebrates
(500-mL or 1-L jars)
10% buffered
formalin f"
10% buffered
formalin
Labels or tags placed inside of the jar must be of
water-resistant paper or 100% rag content paper.
The label on outside of the container should be
completely covered with clear tape.
Confirm the sample ID assigned on the label
matches the ID number recorded on the field col-
lection form (or other sample tracking report).
Special shipping containers, outside labeling, and
snipping forms may be required for shipments con-
taining formalin.
Samples requiring preservation in ethanol
70 % ethanol
Confirm the sample ID assigned on the label
matches the ID number recorded on the field col-
lection form (or other sample tracking report).
Special shipping containers, outside labeling, and
snipping forms may be required for shipments con-
taining ethanol.
41
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0, September 1998 Page 16 of 18
the water samples. Pack and ship sediment toxicity samples using the same type of insu-
lated container and plastic bag arrangement as described above for water chemistry sam-
ples. If available, sediment toxicity samples can be placed inside a plastic container (similar
to the one used for syringe samples) to protect it during shipment.
Samples requiring freezing (Table 3-5) may be stored in the field in a portable
freezer or on dry ice for a short period (e.g., one week). If only ice is available for field
storage, the samples should be shipped to the laboratory as soon as possible after collec-
tion, using fresh ice to keep them as cold as possible. When using ice, double bag the ice
and tape the last bag shut to prevent contamination of samples by melting ice. If possible,
place samples into a sealed plastic container to protect them from meltwater. Dry ice may
also be used for shipping. Note that dry ice is considered a hazardous material, and re-
quires special shipping containers, shipping labels, and shipping forms for ground or air
transport. If dry ice is used, the requirements and directions for packing and shipping
samples ice should be provided to each field team.
Samples that are preserved in buffered formalin (periphyton ID samples and fish
voucher specimens) or ethanol (benthic macroinvertebrate samples) should be transported
in appropriate containers and surrounded with some type of acceptable absorbent material
(e.g., vermiculite). The total volume of formalin in the periphyton ID samples (2 mL per 50-
mL centrifuge tube) may be small enough that they may be shipped without designating
them as a hazardous material. Specific directions for packing, labeling, transporting, and
shipping samples containing formalin or ethanol should be provided to each field team.
Each team leader should contact the field coordinator or other central contact per-
son after each stream visit to provide a brief update of each sampling visit, and to request
replenishment of supplies if necessary. For each shipment, provide the stream identifica-
tion number, date sampled, date that samples are being shipped, and the airbill number
from the courier's shipping form. If the shipment date is on a Friday, call the contact person
or leave a message that a Saturday delivery is coming. Teams should inventory their
supplies after each stream visit and submit requests for replenishment well in advance of
exhausting on-hand stocks.
3.3 EQUIPMENT AND SUPPLIES
A checklist of equipment and supplies required to conduct the activities described in
Section 3 is presented in Figure 3-4. This checklist is similar to the checklist in Appendix
42
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EMAP-SW-Streams Field Operations Manual, Section 3 (Base Location Activities), Rev. 0, September 1998 Page 17 of 18
BASE LOCATION ACTIVITIES
QTY.
ITEM
Before Departure for Stream
1
1
1
1
1
1
1
2
1
1 set
1 set
1 box
1
Dossier of access information for scheduled stream site
Sampling itinerary form or notebook
Safety log and/or personal safety information for each team member
GPS receiver with extra batteries
Dissolved oxygen/temperature meter with probe
Conductivity meter with probe, or conductivity pen
500-mL plastic bottle containing deionized water
500-mL plastic bottles containing conductivity QCCS, labeled "Rinse" and "Test"
Current velocity meter with probe and wading rod
Assorted extra batteries for.dissolved, conductivity, and current velocity meters
Completed water chemistry sample labels (3 labels with same barcode)
Water chemistry sample containers (one 4-L Cubitainer and two 60-mL syringes
with a plastic storage container
Clear tape strips to cover completed sample labels
Checklist of all equipment and supplies required for a stream visit
Packing and Shipping Samples
1 box
1-box
2
2
1
Ice (also dry ice if it is used to ship frozen samples)
1-gal heavy-duty sealable plastic bags
30-gal plastic garbage bags
Insulated shipping containers for frozen samples and sediment toxicity sample
(special containers may be needed if dry ice is used)
Containers and absorbent material suitable to transport and/or ship samples
preserved I formalin and ethanol
Plastic container to hold the sediment toxicity sample
Shipping airbills and adhesive plastic sleeves
Figure 3-4. Equipment and supply checklist for base location activities.
43
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EMAP-SW-Streams Field Operations Manual. Section 3 (Base Location Activities). Rev. 0, September 1998 Page 18 of 18
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 stream site in order to conduct the activities efficiently.
3.4 LITERATURE CITED
Metcalf, R. C., and D. V. Peck. 1993. A dilute standard for pH, conductivity, and acid
neutralizing capacity measurement. Journal of Freshwater Ecology 8:67-72.
Peck, D. V., and R. C. Metcalf. 1991. Dilute, neutral pH standard of known conductivity
and acid neutralizing capacity. Analyst 116:221 -231.
Smoot, G. F., and C. E. Novak. 1968. Calibration and Maintenance of Vertical-axis Type
Current Meters. Book 8, Chapter B2 IN; Techniques of Water-Resources Investiga-
tions of the United States Geological Survey. U.S. Government Printing Office, Wash-
ington, D.C.
44
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SECTION 4
INITIAL SITE PROCEDURES
by
Alan T. Herlihy1
When a field team first arrives at a stream site, they must first confirm they are at
the correct site. Then they determine if the stream meets certain criteria for sampling and
data collection activities to occur. They must decide whether the stream is unduly influ-
enced by rain events which could affect the representativeness of field data and samples.
Certain conditions at the time of the visit may warrant the collection of only a subset of field
measurements and samples. Finally, if it is determined that the stream is to be sampled,
the team lays out a defined reach of the stream within which all subsequent sampling and
measurement activities are conducted.
4.1 SITE VERIFICATION ACTIVITIES
4.1.1 Locating the Index Site
Stream sampling points were chosen from the "blue line" stream network repre-
sented on 1:100,000- scale USGS maps, following a systematic randomized selection
process developed for EMAP stream sampling. Sample sites were then marked with an "X"
on finer-resolution 1:24,000-scale USGS maps. This spot is referred to as the "index site"
or "X-site". The latitude/longitude of the X-site will be listed on a stream information sheet
that is part of the dossier compiled for each stream (see Section 3).
Complete a verification form for each stream visited (regardless of whether you end
up sampling it), following the procedures described in Table 4-1. While traveling from a
base location to a site, record a detailed description of the route taken on page 1 of the
Verification Form (Figure 4-1). This information will allow others to find the site again in the
future. Upon reaching the X-site for a stream, confirm its location and that the team is
1 Dept. of Fisheries and Wildlife, Oregon State University, c/o U.S. EPA, 200 SW 35th St., Corvallis, OR 97333.
45
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EMAP-SW-Streams Field Operations Manual, Section 4 (Initial Site Procedures), Rev. No. 4. September 1998 Page 2 of 12
TABLE 4-1. SITE VERIFICATION PROCEDURES
1. Find the stream location in the field corresponding to the "X" marked on a 7.5" topographic
map (X-site) that is provided with the dossier for each site. Record the routes taken and other
directions on the Verification Form so that someone can visit the same location in the future.
2. Use a GPS receiver to confirm the latitude and longitude at the X-site against the coordinates
provided in the dossier for the site. Record these on the Verification Form.
3. Use all available means to insure that you are at the correct stream as marked on the map,
including: 1:24,000 USGS map orienteering, topographic landmarks, county road maps, local
contacts, etc.
4. Scan the stream channel upstream and downstream from the X-site, and assign one of the
following sampling status categories to the stream. Record the category on the Verification
Form.
Target Categories
A. Regular Wadeable Stream: - The stream can be sampled with wadeable stream procedures.
B. Regular-Mot Wadeable Stream (river): The stream channel is too deep to be safely sampled by wadeable
stream procedures.
If over half of the reach is unwadeable, classify the reach as unwadeable.
If more than half of the reach appears to be wadeable (e.g., only a couple of deep pools), classify the
reach as "Regular-Wadeable" and sample those portions of the reach that can be" safely sampled.
C. Intermittent Stream: The flow of water is not continual, but the channel is wet. Sample using modified proce-
dures.
D. Dry Channel: A discernible stream channel is present but there is no water at the site. Sample using modified
procedures.
E. Altered Channel: There is a stream at the location marked with the X-site on the .map, but the stream channel
does not appear the way it is drawn on the map. An example would be a channel rerouting following a flood
event that cut off a loop of the stream.
Establish a new X-site at the same relative position in the altered channel. Make careful notes and
sketches of the changes on the Verification Form.
Non-target Categories
A. No Stream Channel (map error): No water body or stream channel is present at the coordinates provided for the
X-site.
B. Impounded stream: The stream is submerged under a lake or pond due to man-made or natural (e.g., beaver
dam) impoundments.
If the impounded stream, however, fs still wadeable, record the stream as Altered (Target category E)
and sample the stream.
C. Marsh/Wetland: There is standing water present, but no definable stream channel.
In cases of wetlands surrounding a stream channel, define the site as Target but restrict sampling to the
stream channel.
Inaccessible Categories
A. Physical Barriers: If you are physically unable to reach the X-site because of heavy wetlands, steep gorge or
other barrier that prohibits safe entry.
B. No Permission: You are denied access to the site by the landowners.
5. Do not sample "Non-target" or "Inaccessible" sites. Place an "X" in the appropriate box in the
"NON-SAMPLEABLE" section of the Verification Form and provide an explanation in the com-
ments section.
46
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EMAP-SW-Streams Field Operations Manual. Section 4 (Initial Site Procedures). Rev. No. 4, September 1998 Page 3 of 12
Reviewed by (initial):
VERIFICATION FORM - STREAMS/RIVERS
SITE NAME:
£W*
DATE: 7//^"/97 VISIT: 811.02 _
SITE ID: MAIA97-_2_
TEAM ID (X): H1 D2 D3 O4 D5 D6 D7 D8
STREAM/RIVER VERIFICATION INFORMATION
STREAM/RIVER VERIFIED BY (X all that apply): [g GPS [53 LOCAL CONTACT Q SIGNS j£J ROADS J^TOPO.MAP •
[] OTHER (DESCRIBE HERE): Q NOT VERIFIED (EXPLAIN IN COMMENTS)
. COORDINATES
LATITUDE (dd mm ss) North
LONGITUDE (dddmmss) West
TYPEOFGPSF.X
MAP:
GPS:
.72.
H3D
EYES
D NO
INDEX SITE STATUS - X ONE BOX FROM ONE SECTION ONLY
SAMPLEABLE
jy] REGULAR-WADEABLE
Q REGULAR-NOTWADEABLE
£3 INTEHMITTENT-DRY SPOTS ALONG REACH
[] DRY-No WATER ANYWHERE ALONG REACH
Q ALTERED-STREAM/RIVER PRESENT BUT NOT AS OH MAP
Q OTHER (EXPLAIN IN COMMENTS)
NON-SAMPLEABLE (No SAMPLE TAKEN)
Q No CHANNEL OR WATERBODY PRESENT
Q IMPOUNDED (UNDERNEATH LAKE/POND)
Q WETLAND (NO DEFINABLE CHANNEL)
NO ACCESS
Q ACCESS PERMISSION DENIED
Q INACCESSIBLE (UNABLE TO REACH srre)
DIRECTIONS TO STREAM/RIVER SITE
•••/
rf. ««>/ «.
-------
EMAP-SW-Streams Field Operations Manual, Section 4 (Initial Site Procedures), Rev. No. 4, September 1998 Page 4 of 12
at the correct stream. Use all available means to accomplish this, and record the informa-
tion on page 1 of the Verification Form (Figure 4-1).
4.1.2 Determining the Sampling Status of a Stream
Not all chosen stream sites will turn out to be streams. On the basis of previous
synoptic surveys, it was found that the maps are far from perfect representations of the
stream network. A significant part of EMAP is the estimation of the actual extent of stream
length in the area. After the stream and location of the X-site are confirmed, evaluate the
stream reach surrounding the X-site and classify the stream into one of three major sam-
pling status categories (Table 4-1). The primary distinction between "Non-target" and "Tar-
get" streams is based on the presence of a defined stream channel and its depth.
Record the site class and pertinent site verification information on the Verification
Form (Figure 4-1). If the site is non-target or inaccessible, the site visit is completed, and
no further sampling activities are conducted.
4.1.3 Sampling During or After Rain Events
Avoid sampling during high flow rainstorm events. For one, it is often unsafe to be in
the water during such times. In addition, biological and chemical conditions during episodes
are often quite different from those during baseflow. On the other hand, sampling cannot
be restricted to only strict baseflow conditions. It would be next to impossible to define
"strict baseflow" with any certainty at an unstudied site. Such a restriction would also
greatly shorten the index period when sampling activities can be conducted. Thus, some
compromise is necessary regarding whether to sample a given stream because of storm
events. To a great extent, this decision is based on the judgment of the field team. Some
guidelines to help make this decision are presented in Table 4-2. The major indicator of the
influence of storm events will be the condition of the stream itself. If a field team decides a
site is unduly influenced by a storm event, do not sample the site that day. Notify the field
coordinator or other central contact person to reschedule the stream for another visit.
4.1.4 Site Photographs
Taking site photographs is an optional activity, but should be considered if the site
has unusual natural or man-made features associated with it. If you do take any photo-
graphs at a stream, start the sequence with one photograph of an 8.5 * 11 inch piece of
48
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EMAP-SW-Streams Field Operations Manual, Section 4 (Initial Site Procedures), Rev. No. 4. September 1998 Page 5 of 12
TABLE 4-2. 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 if it is unsafe to wade in the majority of the stream reach.
• 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 un-
duly influenced by storm events, go ahead and sample it, even if it has recently rained or
is raining.
paper with the stream ID and date printed in large letters. After the photo of the stream 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. For pictures of aquatic vertebrates (see Section 12) or other small objects, place
the paper with the stream ID and date in each snapshot.
4.2 LAYING OUT THE SAMPLING REACH
Unlike chemistry, which can be measured at a point, most of the biological and
habitat structure measures require sampling a certain length of a stream to get a represen-
tative picture of the ecological community. Previous EMAP pilot studies have suggested
that a length of 40 times the channel width is necessary to collect at least 90% of the fish
species occurring in the stream reach. Thus, a support reach that is 40 channel widths long
around the X-site is required to characterize the community and habitat associated with the
sampling point. Establish the sampling reach about the X-site using the procedures de-
scribed in Table 4-3. 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 (or the midpoint of the reach) on page 2 of the Verification Form as shown in
Figure 4-2. Figure 4-3 illustrates the principal features of the established sampling reach,
including the location of 11 cross-section transects used for physical habitat characterization
(Section 7), and specific sampling points on each cross-section transect for later collection
of periphyton samples (Section 8) and benthic macroinvertebrate samples (Section 11).
49
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EMAP-SW-Streams Field Operations Manual. Section 4 (Initial Site Procedures), Rev. No. 4, September 1998 Page 6 of 12
TABLE 4-3. LAYING OUT THE SAMPLING REACH
1. Use a surveyor's rod or tape measure to determine the wetted width of the channel at five
places considered to be of "typical" width within approximately 5 channel widths upstream and
downstream from the X-site. Average the five readings together and round to the nearest 1
m. If the average width is less than 4 m, use 150 m as a minimum sample reach length.
Record this width on page 2 of the Verification Form.
For dry or intermittent channels, estimate the width based on the unvegetated width of
the channel.
2. Check the condition of the stream upstream and downstream of the X-site by having one team
member go upstream and one downstream. Each person proceeds until they can see the
stream to a distance of 20 times the average channel width (equal to one-half the sampling
reach length) determined in Step 1 from the X-site.
For example, if the reach length is determined to be 150 m, each person would proceed
75 m from the X-site to lay out the reach boundaries.
3. Determine if the reach needs to be adjusted about the X-site due to confluences with higher
order streams (downstream), lower order streams (upstream), or lakes, reservoirs, or ponds.
If such a confluence is reached, note the distance and flag the confluence as the end-
point of the reach. Move the other endpoint of the reach an equivalent distance away
from the X-site.
NOTE: Do not slide the reach to avoid man-made obstacles such as bridges,
culverts, rip-rap, or channelization.
4. Starting back at the X-site (or the new midpoint of the reach if it had to be adjusted as de-
scribed in Step 3), measure a distance of 20 channel widths down the middle of the stream
using a tape measure. Be careful not to "cut corners". Enter the channel to make measure-
ments only when necessary to avoid disturbing the stream channel prior to sampling activities.
This endpoint is the downstream end of the reach, and is flagged as transect "A".
5. Using the tape measure, measure 1/10 (4 channel widths in big streams or 15 m in small
streams) of the required stream length upstream from the start point (transect A). Flag this
spot as the next cross-section or transect (transect B). For transect B, roll the dice to deter-
mine if it is a left (L), center (C), or right (R) sampling point for collecting periphyton and
benthic macroinvertebrate samples. A roll of 1 or 2 indicates L, 3 or 4 indicates C, and 5 or 6
indicates R (or use a digital wristwatch and glance at the last digit (1-3=L, 4-6=C, 7-9=R).
Mark L, C, or R on the transect flagging.
6. Proceed upstream with the tape measure and flag the positions of 9 additional transects
(labeled "C" through "J" as you move upstream) at intervals equal to 1/10 of the reach length.
Assign sampling spots to each transect in order as L, C, R after the first random selection.
For example, if the sampling spot assigned to transect "B" was C, transect "C" is as-
signed R, transect "D" is L, transect "E" is C, etc.
50
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EMAP-SW-Streams Field Operations Manual. Section 4 (Initial Site Procedures). Rev. No. 4, September 1998 Page 7 of 12
Reviewed by (initial):
VERIFICATION FORM - STREAMS/RIVERS (continued)
SITE NAME: M fit. CtfFK.
DATE: n f/ff&7 VISIT: fcl Q2
SITEID:
TEAM ID (X): 81 D2 D3 D4 O5 D6 D7 D8
STREAM/RIVER REACH DETERMINATION
CHANNEL WIDTH USED TO DEFINE
REACH (u) (XX):
DISTANCE (M) FROM X-SITE
UPSTREAM LENGTH DOWNSTREAM LENGTH
COMMENT
Rev. 06/02/97 (strwad^T)
VERIFICATION FORM - STREAMS/RIVERS - 2
Figure 4-2. Verification Form (page 2).
51
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EMAP-SW-Streams Reid Operations Manual, Section 4 (Initial Site Procedures), Rev. No. 4, September 1998 Page 8 of 12
D
Rimmcm
SAMPLING POINTS
• L=Left C=Center R=Right
• First point (transect B)
determined at random
• Subsequent points assigned in
order L, C, R
Distance between transects=4 times
mean wetted width at X-site
Total reach length=40 times mean wetted width at X-site (minimum=150 m)
Figure 4-3. Sampling reach features.
There are some conditions that may require adjusting the reach about the X-site
(i.e., the X-site no longer is located at the midpoint of the reach) to avoid features we do not
wish to sample across. 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). If such a confluence is reached, note the distance and flag the confluence as
the endpoint of the reach. Make up for the loss of reach length by moving ("sliding") the
other end of the reach an equivalent distance away from the X-site. Similarly, if you run into
a lake, reservoir, or pond while laying out the reach, stop, flag the lake/stream confluence
as the reach end, and make up for the loss of reach length by moving the other end of the
reach an equivalent distance from the X-site. Do not "slide" the reach so that the X-site falls
outside of the reach boundaries. Also, do not "slide" a reach to avoid man-made obstacles
52
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EMAP-SW-Streams Field Operations Manual, Section 4 (Initial Site Procedures), Rev. No. 4, September 1998 Page 9 of 12
such as bridges, culverts, rip-rap, or channelization. These represent features and effects
that EMAP is attempting to study. ,
Before leaving the stream, complete a rough sketch map of the stream reach you
sampled on the page 2 of the Verification Form (Figure 4-2). In addition to any other inter-
esting features that should be marked on the map, note any landmarks/directions that can
be used to find the X-site for future visits.
4.3 MODIFIED PROCEDURES FOR DRY AND INTERMITTENT STREAMS
The full complement of field data and samples cannot be collected from streams that
are categorized as "Dry Channel" or "Intermittent" (Table 4-1). Physical habitat information
(Section 7) is collected in all streams. Intermittent streams will have some cross-sections
with biological measurements and some with none. Totally dry channels will have no bio-
logical sampling. Modified procedures for dry and intermittent streams are presented in
Table 4-4.
Samples and measurements for water chemistry (Section 5) should be collected at
the X-site (even if the reach has been adjusted by "sliding" it). If the X-site is dry, the sam-
ple and chemical measurements are taken from a location having water with a surface area
greater than 1 m2 and a depth greater than 10 cm.
All data for the physical habitat indicator (Section 7) are collected from all streams,
regardless of the amount of water present in the channel or at the transects. Depth mea-
surements along the deepest part of the channel (the "thalweg") are obtained along the
entire sampling reach for ALL target streams, whether they are dry, intermittent, or com-
pletely flowing. The thalweg profile provides 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.
4.4 EQUIPMENT AND SUPPLIES
A list of the equipment and supplies required to conduct the stream verification and
to lay out the sampling reach is presented in Figure 4-4. This checklist is similar to the
checklist presented in Appendix A, which is used at the base location (Section 3) to ensure
that all of
53
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EMAP-SW-Streams Reid Operations Manual, Section 4 (Initial Site Procedures). Rev. No. 4, September 1998 Page 10 of 12
TABLE 4-4. MODIFICATIONS FOR DRY CHANNELS AND INTERMITTENT STREAMS
Water Chemistry
If the X-site is dry but there is flowing water or a pool of water having a surface area greater
than 1 m2 and a depth greater than 10 cm somewhere along the defined sampling reach, take
the water sample and water chemistry measurements 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 chemistry sample on the sample collection form and explain why the
sample was not collected in the comments section of the form.
Physical Habitat Characterization, Periphyton, Sediment Metabolism,
and Benthic Macroinvertebrates
Obtain a complete thalweg profile for the entire reach, even if the channel is completely dry.
At points where channel is dry, record depth as 0 cm and wetted width as 0 m.
At each of the transects (cross sections), classify the stream as:
DRY CHANNEL: No surface water anywhere in cross section;
Collect all physical habitat data. Use the unvegetated area of the channel to deter-
mine 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.
DAMP CHANNEL: Wet spots in cross section but NO flowing water or pools > 10 cm
deep;
Collect all physical habitat data.
Collect periphyton samples from the wet spots. These are great environments for
algae.
Collect sediments for metabolism if there are enough fine wet sediments available.
Do not collect a benthic macroinvertebrate sample.
ENDURING POOLS: No flowing water but pools > 10 cm deep;
Collect all data and measurements for physical habitat, periphyton, sediment
metabolism, and benthic macroinvertebrate indicators, using standard procedures.
FLOWING WATER: Flowing water in cross section
Collect all data and measurements for physical habitat, periphyton, sediment
metabolism, and benthic macroinvertebrate indicators, using standard procedures.
Aquatic Vertebrates
Do not sample if the entire reach is dry.
In intermittent streams (including those having damp channels and/or enduring pools), sample
any wet areas within the sampling reach that are potential habitat for aquatic vertebrates. Do
not sample downstream of Transect "A" or upstream of Transect "K", even if there
appears to be good habitat present.
54
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EMAP-SW-Streams Field Operations Manual, Section 4 (Initial Site Procedures), Rev. No. 4, September 1998 Page 11 of 12
EQUIPMENT AND SUPPLIES FOR INITIAL SITE ACTIVITIES
QTY.
1
1
1
1
1
1
1
2 rolls
1
1 copy
1 set
Item
Dossier of site and access information
Topographic map with "X-site" marked
Site information sheet with map coordinates and elevation of X-site
GPS receiver and operating manual
Extra batteries for GPS receiver
Verification Form
Soft lead (#2) pencils
Surveyor's telescoping leveling rod
50-m fiberglass measuring tape with reel
Surveyor's flagging tape (2 colors)
Fine-tipped indelible markers to write on flagging
Waterproof camera and film
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for initial
site activities
Figure 4-4. Equipment and supplies checklist for initial site activities.
55
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EMAP-SW-Streams Field Operations Manual, Section 4 (Initial Site Procedures). Rev. No. 4, September 1998 Page 12 of 12
the required equipment is brought to the stream. Use this checklist to ensure that equip-
ment and supplies are organized and available at the stream site in order to conduct the
activities efficiently.
56
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SECTION 5
WATER CHEMISTRY
by
Alan T. Herlihy1
There are two components to collecting water chemistry information: Collecting
samples of stream water to ship to the analytical laboratory, and obtaining in situ or
streamside measurements of specific conductance, dissolved oxygen, and temperature. At
each stream, teams fill one 4-L container and two 60 mL syringes with streamwater. These
samples are stored in a cooler packed with plastic bags filled with ice and are shipped or
driven to the analytical laboratory within 24 hours of collection (see Section 3). The primary
purposes of the water samples and the field chemical measurements are to determine:
• Acid-base status
Trophic condition (nutrient enrichment)
Chemical Stressors
Classification of water chemistry type.
Water from the 4-L bulk sample is used to measure the major cations and anions,
nutrients, total iron and manganese, turbidity and color. The syringe samples are analyzed
for pH, dissolved inorganic carbon, and monomeric aluminum species. Syringes are used
to seal off the samples from the atmosphere because the pH, dissolved inorganic carbon
(DIG), and aluminum concentrations will all change if the streamwater equilibrates with
atmospheric CO2. Overnight express mail for these samples is required because the
syringe samples need to be analyzed, and the 4-L bulk sample needs to be stabilized (by
filtration and/or acidification) within a short period of time (72 hours) after collection.
In situ and streamside measurements are made using field meters and recorded on
standard data forms. Specific conductance (or conductivity) is a measure of the ability of
the water to pass an electrical current which is related to the ionic strength of a solution.
Dissolved oxygen (DO) is a measure of the amount of oxygen dissolved in solution. In
Department of Fisheries and Wildlife, Oregon State University, c/o U.S. EPA, 200 SW 35th St., Corvallis, OR 97333
57
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EMAP-SW-Streams Field Operations Manual, Section 5 (Water Chemistry), Rev. No. 4, September 1998 Page 2 of 9
natural waters, minimal concentrations of oxygen are essential for survival of most aquatic
organisms. Measures of DO and temperature are used to assess water quality and the
potential for healthy aerobic organism populations. Most of the procedures outlined in this
section are similar to the ones utilized by the EPA in streams for the National Surface Water
Survey (Kaufmann et al., 1988) and have been adapted from the Survey's field operations
handbook (U.S. EPA, 1989).
5.1 SAMPLE COLLECTION
Before leaving the base location, package the sample containers (one 4-L
cubitainer and two 60 mL syringes) and the stream sample beaker to prevent contamination
(see Section 3). Fill out a set of water chemistry sample labels as shown in Figure 5-1.
Attach a completed label to the cubitainer and each syringe and cover with clear tape strips
as described in Section 3. Make sure the syringe labels do not cover the volume gradations
on the syringe. In the field, make sure that the labels all have the same sample ID number
(barcode), and that the labels are securely attached.
The procedure to collect a water chemistry sample is described in Table 5-1. The
sample is collected from the middle of the stream channel at the X-site, unless no water is
present at that location (see Section 4). Throughout the sampling process, it is important to
take precautions to avoid contaminating the sample. Rinse all sample containers three
times with portions of stream water before filling them with the sample. Many of the
streams have a very low ionic strength and can be contaminated quite easily by perspiration
from hands, sneezing, smoking, insect repellent, or other chemicals used when collecting
other types of samples. Thus, make sure that none of the water sample contacts your
hands before going into the cubitainer. All of the chemical analyses conducted using the
syringe samples are affected by equilibration with atmospheric carbon dioxide; thus, it is
essential that no outside air contact the syringe samples during or after collection.
Record the information from the sample label on the Sample Collection Form as
shown in Figure 5-2. Note any problems related to possible contamination in the comments
section of the form.
5.2 FIELD MEASUREMENTS
Table 5-2 presents the procedures for obtaining field measurement data for the
water chemistry indicator. The conductivity and dissolved oxygen meters are checked in
58
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EMAP-SW-Streams Field Operations Manual. Section 5 (Water Chemistry), Rev. No. 4, September 1998 Page 3 of 9
WATER CHEMISTRY
CLl(si)s2
SITEID:MAIA_2.2--?.ll
DATE: JL/j£/98
229015
WATER CHEMISTRY
(CU)S1 S2
SITE ID: MAIA_2_Z-_l_i_£
DATE:
229015
WATER CHEMISTRY
cu si (32)
SITE ID: MAIA_2_2-.2.Z2.
DATE:
229015
Figure 5-1. Completed sample labels for water chemistry.
the field using the same procedures as those used at a base location (Section 3). The
quality control check sample solution (QCCS) is prepared according to directions presented
in Section 3. The results of field checks of these meters, as well as the measured values
for specific conductance, dissolved oxygen, and stream temperature, are recorded on the
Field Measurement Form as shown in Figure 5-3.
5.3 EQUIPMENT AND SUPPLIES
A list of equipment and supplies required to collect samples and field data for the
water chemistry indicator is presented in Figure 5-4. This checklist is similar to the checklist
presented in Appendix A, which is used at the base location (Section 3) to ensure that all of
the required equipment is brought to the stream. Use this checklist to ensure that
equipment and supplies are organized and available at the stream site in order to conduct
the activities efficiently.
5.4 LITERATURE CITED
Kaufmann, P., A. Herlihy, J. Elwood, M. Mitch, S. Overton, M. Sale, J. Messer, K. Reckhow,
K. Cougan, D. Peck, J, Coe, A. Kinney, S. Christie, D. Brown, C. Hagley, and Y. Jager.
1988. Chemical Characteristics of Streams in the Mid-Atlantic and Southeastern
59
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EMAP-SW-Streams Field Operations Manual, Section 5 (Water Chemistry), Rev. No. 4, September 1998 Page 4 of 9
TABLE 5-1. SAMPLE COLLECTION PROCEDURES FOR WATER CHEMISTRY
Collect the water samples from the X-site in a flowing portion near the middle of the stream.
1. Rinse the 500 ml_ sample beaker three times with streamwater, Discard the rinse
downstream.
2. Remove the cubitainer lid and expand the cubitainer by pulling out the sides. NOTE: DO
NOT BLOW into the cubitainers to expand them, this will cause contamination.
3. Fill the beaker with streamwater and slowly pour 30-50 ml_ into the cubitainer. Cap the
cubitainer and rotate it so that the. water contacts all the surfaces. Discard the water
downstream. Repeat the above rinsing procedure two more times.
4. Collect additional portions of streamwater with the beaker and pour them into the cubitainer.
Let the weight of the water expand the cubitainer. The first two portions will have to be
poured slowly as the cubitainer expands. Fill the cubitainer to its maximum volume. Rinse
the cubitainer lid with streamwater. Eliminate any air space from the cubitainer, arid cap it
tightly. Make sure the cap is tightly sealed and not on at an angle.
5. Place the cubitainer in a cooler (on ice or streamwater) and shut the lid. If a cooler is not
available, place the cubitainer in an opaque garbage bag and immerse it in the stream.
6. Submergie a 60-mL syringe halfway into the stream and withdraw a 15-20 mL aliquot. Pull
the plunger to its maximum extension and shake the syringe so the water contacts all
surfaces. Point the syringe downstream and discard the water by depressing the plunger.
Repeat the rinsing procedure two more times.
7. Submerge the syringe into the stream again and slowly fill the syringe with a fresh sample.
Try not to get any air bubbles in the syringe. If more than 1-2 tiny bubbles are present,
discard the sample and draw another one.
8. Invert the syringe (tip pointing up), and cap it with a syringe valve. Tap the syringe lightly to
detach any trapped air bubbles. With the valve open, expel the air bubbles and a small
volume of water, leaving between 50 and 60 mL of sample in the syringe. Close the syringe
valve. If any air bubbles were drawn into the syringe during this process, discard the sample
and fill the syringe again (step 8).
9. Repeat Steps 6 through 8 with a second syringe. Place the syringes together in the cooler or
in the streamwater with the cubitainer.
10. Record the barcode number (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, record an "X" 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.
11. After carrying the samples out to the vehicles, place the cubitainer and syringes in a cooler
and surround with 1 gallon self-sealing plastic bags filled with ice.
60
-------
EMAP-SW-Streams Field Operations Manual, Section 5 (Water Chemistry), Rev. No. 4, September 1998 Page 5 of 9
Reviewed by (initial):
SAMPLE COLLECTION FORM - STREAMS (continued)
SITE NAME:
DATE: "7
VISIT: H1 D2
SITE ID: MAIA97-
TEAM ID 00: H1 D2 D3 H4 HS
C17 H8
CHEMISTRY AND MICROBIAL WATER SAMPLE (Chem: 4-L Cubitainer and 2 Syringes, Micro: Glass Bottle)
SAMPLE ID (BARCODE)
TRANSECT
FLAG
COMMENTS
CHEMISTRY
MICROBIAL.
SEDIMENT TOXICITY SAMPLES
SAMPLE ID (BARCODE)
FLAG
COMMENTS
FISH TISSUE SAMPLES - PRIMARY SAMPLE (min. 50g total wgt)
SAMPLE ID (BARCODE) -
LINE
SPECIES CODE
COMMON NAME
NUMBER OF
INDIVIDUALS
FLAG
P1
B/
t»e
IS COMPOSITE SAMPLE COMPOSED OF INDIVIDUALS COLLECTED FROM THROUGHOUT REACH? (X) -
H YES D No
IF No, EXPLAIN:
FISH TISSUE SAMPLES - SECONDARY SAMPLE (where available; 5 Individuals)
SAMPLE ID (BARCODE) -
LINE
SPECIES CODE
COMMON NAME
TOTAL LENGTH (MM)
FLAG
S1
b->tit
jzr
S2
S3
sucke
S4
S5
fTe Stfckt
IS COMPOSITE SAMPLE COMPOSED OF INDIVIDUALS COLLECTED FROM THROUGHOUT REACH? (X) -
BYES Q No
IF No, EXPLAIN:
LINE
COMMENT OR FLAG EXPLANATION FOR FISH TISSUE
Fl
60 a.
nag codes: K= sampla not collected; U= Suspect sample; F1.F2, otc.= misc. flag assigned by fldd crew. Explain all flags In Comments sections.
Rev. Q&02I97 (sL_saco.9T) SAMPLE COLLECTION FORM - STREAMS - 2
Figure 5-2. Sample Collection Form (page 2), showing data recorded for water chemistry
samples.
61
-------
EMAP-SW-Streams Field Operations Manual, Section 5 (Water Chemistry), Rev. No. 4, September 1998 Page 6 of 9
TABLE 5-2. PROCEDURES FOR STREAMSIDE AND IN SITU CHEMISTRY MEASUREMENTS
Specific Conductance
1. Check the batteries and electronic functions (e.g., zero, "red line") of the conductivity meter (or
a conductivity pen) as instructed by the operating manual.
2. Insert the probe into the "RINSE" container of the quality control check sample (QCCS) and
swirl for 3 to 5 seconds. Transfer the probe to the "TEST" container of QCCS let stabilize for
20 seconds. Record the conductivity of the QCCS on the Field Measurement Form.
If the measured conductivity is not within 10% or 10 uS/cm of theoretical value, repeat
the measurement process. If the value is still unacceptable, flag the conductivity data
on the Field Measurement Form.
3. Submerge the probe in and area of flowing water near the middle of the channel at the same
location where the water chemistry sample is collected. Record the measured conductivity on
the Field Measurement Form.
Dissolved Oxygen and Temperature
1. Inspect the probe for outward signs of fouling and for an intact membrane. Do not touch the
electrodes inside the probe with any object. Always keep the probe moist by keeping it inside
its calibration chamber.
2. Check the batteries and electronic functions of the meter as described in the operating
manual. Record the results of these checks on the Field Measurement Form.
2. Calibrate the oxygen probe in water-saturated air as described in the operating manual. Allow
at least 15 minutes for the probe to equilibrate before attempting to calibrate, try to perform
the calibration as close to stream temperature as possible (not air temperature) by using
stream water to fill the calibration chamber prior to equilibration. For doing the elevation
correction, the elevation of the sample site is given on the site Information sheet in the dossier
for the site. Record the pertinent calibration information on the Field Measurement Form.
3. After the calibration, submerge the probe in midstream at mid-depth at the same location
where the water chemistry sample is collected. Face the membrane of the probe upstream,
and allow the probe to equilibrate. Record the measured DO and stream temperature on the
Field Measurement Form. If the DO meter is not functioning, measure the stream
temperature with a field thermometer and record the reading on the Field Measurement Form
along with pertinent data flags and comments.
NOTE: Older model dissolved oxygen probes require a continuous movement of
water (0.3 to 0.5 mis) across the probe to provide accurate measurements. If the
velocity of the stream is appreciably less than that, jiggle the probe in the water as
you are taking the measurement.
62
-------
EMAP-SW-Streams Field Operations Manual, Section 5 (Water Chemistry). Rev. No. 4. September 1998 Page 7 of 9
Reviewed by (Initial):
FIELD MEASUREMENT FORM - STREAMS/RIVERS
SITE NAME: AtlU CttfeK DATE: 7 / fSTl 97 VISIT: H1 D2
SITE ID: M A I A97-_5__?_-2_ TEAM ID (X): H1 D2 Q3 D4 D5 Q6 Q7 Q8
WEATHER CONDITIONS (X)
CLOUD COVER
PRECIPITATION
PREVIOUS PRECIPITATION (24 H)
AIR TEMPERATURE XX
H<5% DS-25% D 25-50% Q 50-75% D>75%
HNONE D LIGHT D MODERATE D HEAVY
DNONE H LIGHT D MODERATE D HEAVY
z r "c
IN SITU MEASUREMENTS STATION ID: £" \ Assume X-sIle unless marked
QCCSCOND«S/CM
STREAM/RIVER COND ^S/CM
STREAM/RIVER DO MQ/L
STREAM/RIVER TEMP °c
FLAG COMMENTS
y_2^
S" o o
H.&
Z 0 .jQ_
STREAM/RIVER METABOLISM DETERMINATION
INITIAL 02 . ''
(MO/L) !£
—M...2. A
SAMPLE ID
(BARCODE)
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MITIAL
*P.(°C)
0..3L
FINAL O2
(Moa.)
— 3L.S-
L.JL
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Ttec«IIx«tloitvi]u>l>cbti]n>dl>rmult!plyli>gtlia
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motar rMdng to tto olfcratlon vahw.
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COMMENTS:
Flag Codes: K - no measurement or observation made; U= suspect measurement or observation; Q = unacceptable QC check associated with
measurement; F1, F2, etc. = miscellaneous flags assigned by each field crew. Explain all flags In comments section.
Rev. 06/02/97 (strvfldm.97)
FIELD MEASUREMENT FORM - STREAMS/RIVERS - 1
Figure 5-3. Field Measurement Form (page 1), showing data recorded for water chemistry.
63
-------
EMAP-SW-Streams Field Operations Manual, Section 5 (Water Chemistry), Rev. No. 4. September 1998 Page 8 of 9
EQUIPMENT AND SUPPLIES FOR WATER CHEMISTRY
QTY.
1
1
1
1
1
1
1
1
1
2
1
2
1
1
1
1 copy
1 set
Item
Dissolved oxygen/Temperature meter with probe
DO repair kit containing additional membranes and probe filling solution
Conductivity meter with probe
500-mL plastic bottle of conductivity QCCS labeled "Rinse" (in plastic bag)
500-mL plastic bottle of conductivity QCCS labeled "Test" (in plastic bag)
500-mL plastic bottle of deionized water to store conductivity probe
Field thermometer
500 mL plastic beaker with handle (in clean plastic bag)
4-L cubitainer with completed sample label attached (in clean plastic bag)
60 mL plastic syringes (with Luer type tip) with completed sample labels
attached
Plastic container with snap-on lid to hold filled syringes
Syringe valves (Mininert® with Luer type adapter, or equivalent, available from a
chromatography supply company)
Cooler with 4 to 6 plastic bags (1-gal) of ice OR
a medium or large opaque garbage bag to store the water sample at streamside
Sample Collection From
Field Measurement Form
Soft-lead pencils for filling out field data forms
Fine-tipped indelible markers for filling out labels
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for water
chemistry
Figure 5-4. Checklist of equipment and supplies for water chemistry.
64
-------
EMAP-SW-Streams Field Operations Manual, Section 5 (Water Chemistry), Rev. No. 4. September 1998 Page 9 of 9
United States. Volume I: Population Descriptions and Physico-Chemical
Relationships. EPA 600/3-88/021 a. U.S. Environmental Protection Agency,
Washington, D.C.
U.S. EPA. 1989. Handbook of Methods for Acid Deposition Studies: Field Operations for
Surface Water Chemistry. EPA 600/4-89/020. U.S. Environmental Protection Agency,
Office of Research and Development, Washington, D.C.
65
-------
-------
SECTION 6
STREAM DISCHARGE
by
Philip R. Kaufmann1
Stream discharge is equal to the product of the mean current velocity and vertical
cross sectional area of flowing water. Discharge measurements are critical for assessing
trends in streamwater acidity and other characteristics that are very sensitive to streamflow
differences. Discharge should be measured at a suitable location within the sample reach
that is as close as possible to the location where chemical samples are collected (typically
the X-site; see Section 5), so that these data correspond.
Discharge is usually determined after collecting water chemistry samples. Although
discharge is part of the physical habitat indicator (Section 7), it is presented as a separate
section because the "biomorphs" measure while the "geomorphs" conduct the other habitat
characterization procedures (see Section 2).
No single method for measuring discharge is applicable to all types of stream chan-
nels. The preferred procedure for obtaining discharge data is based on "velocity-area"
methods (e.g., Rantz and others, 1982; Linsley et al., 1982). For streams that are too small
or too shallow to use the equipment required for the velocity-area procedure, two alternative
procedures are presented. One procedure is based on timing the filling of a volume of
water in a calibrated bucket. The second procedure is based on timing the movement of a
neutrally buoyant object (e.g., an orange) through a measured length of the channel, after
measuring one or more cross-sectional depth profiles within that length.
6.1 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 flow cross-sectional area of
many increments across a channel (Figure 6-1). Each increment gives a subtotal of the
1 Department of Fisheries and Wildlife, Oregon State University, c/o U.S. EPA, 200 SW 35th St., Corvallis, OR 97333.
67
-------
EMAP-SW-Streams Field Operations Manual. Section 6 (Stream Discharge), Rev. 5. September 1998 Page 2 of 10
15 to 20 equally spaced
intervals across stream
Midpoints of intervals: measure
stream depth and obtain velocity
measurements at 0.6 depth
WATER SURFACE
Jl
Extended surveyor's
rod or tape measure
Figure 6-1. Layout of channel cross-section for obtaining discharge data by the velocity-area
procedure.
stream discharge, and the whole is calculated as the sum of these parts. Discharge mea-
surements are made at only one carefully chosen channel cross section within the
sampling reach. It is important to choose a channel cross section that is as much like a
canal as possible. A glide area with a "U" shaped channel cross section that is free of
obstructions provides the best conditions for measuring discharge by the velocity-area
method. You may remove rocks and other obstructions to improve the cross-section before
any measurements are made. However, because removing obstacles from one part of a
cross-section affects adjacent water velocities, you must not change the cross-section once
you commence collecting the set of velocity and depth measurements.
The procedure for obtaining depth and velocity measurements is outlined in Table
6-1. Record the data from each measurement on page 2 of the Field Measurement Form
68
-------
EMAP-SW-Streams Field Operations Manual, Section 6 (Stream Discharge), Rev. 5, September 1998 Page 3 of 10
TABLE 6-1. VELOCITY-AREA PROCEDURE FOR DETERMINING STREAM DISCHARGE
1. Locate a cross-section of the stream channel for discharge determination that has most of the
following qualities (based on Rantz and others, 1982):
Segment of stream above and below cross-section is straight
• Depths mostly greater than 15 centimeters, and velocities mostly greater than 0.15
meters/second. Do not measure discharge in a pool.
"U" shaped, with a uniform streambed free of large boulders, woody debris or brush, and
dense aquatic vegetation.
Flow is relatively uniform, with no eddies, backwaters, or excessive turbulence.
2. Lay the surveyor's rod (or stretch a meter 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. Place an
"X" in the "VELOCITY AREA" box in the "STREAM DISCHARGE" section of the Field Measure-
ment 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.
5. Stand downstream of the rod or tape and to the side of the midpoint of the first interval (closest
to the left bank if looking downstream).
6. Place the wading rod in the stream at the midpoint of the interval and adjust the probe or
propeller so that it is at the water surface. Record the distance from the left bank (in centime-
ters) and the depth indicated on the wading rod (in centimeters) on the Field Measurement
Form.
7. Stand downstream of the probe or propeller to avoid disrupting the stream flow. Adjust
the position of the probe on the wading rod so it is at 0.6 of the measured depth below the
surface of the water. Face the probe upstream at a right angle to the cross-section, even if
local flow eddies hit at oblique angles to the cross-section.
8. Wait 20 seconds to allow the meter to equilibrate, then measure the velocity. Record the value
on the Field Measurement Form.
For the electromagnetic current meter Ce.g..-Marsh-McBirnev). use the lowest time con-
stant 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 mea-
surements.
9. Move to the midpoint of the next interval and repeat Steps 6 through 8. Continue until depth
and velocity measurements have been recorded for all intervals.
69
-------
EMAP-SW-Streams Field Operations Manual, Section 6 (Stream Discharge), Rev. 5. September 1998 Page 4 of 10
as shown in Figure 6-2. To reduce redundancy and to conserve space, Figure 6-2 shows
measurement data recorded for all three procedures. In the field, data will be recorded
using only one of the three available procedures.
6.2 TIMED FILLING PROCEDURE
In channels too "small" for the velocity-area method, discharge can sometimes be
determined directly by measuring the time it takes to fill a container of known volume.
"Small" is defined as a channel so shallow that the current velocity probe cannot be placed
in the water, or where the channel is broken up and irregular due to rocks and debris, and a
suitable cross-section for using the velocity area procedure is not available. This can be an
extremely precise and accurate method, but requires a natural or constructed spillway of
free-falling water. If obtaining data by this procedure will result in a lot of channel distur-
bance or stir up a lot of sediment, wait until after all biological and chemical measurements
and sampling activities have been completed.
Choose a cross-section of the stream that contains one or more natural spillways or
plunges that collectively include the entire stream flow. A temporary spillway can also be
constructed using a portable V-notch weir, plastic sheeting, or other materials that are
available onsite. Choose a location within the sampling reach that is narrow and easy to
block when using a portable weir. Position the weir in the channel so that the entire flow of
the stream is completely rerouted through its notch (Figure 6-3). Impound the flow with the
weir, making sure that water is not flowing beneath or around the side of the weir. Use mud
or stones and plastic sheeting to get a good waterproof seal. The notch must be high
enough to create a small spillway as water flows over its sharp crest.
The timed filling procedure is presented in Table 6-2. 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 Field Measurement Form as shown in Figure 6-2. Repeat the procedure 5 times. If
the cross-section contains multiple spillways, you will need to do separate determinations
for each spillway. If so, clearly indicate which time and volume data replicates should be
averaged together for each spillway; use additional field measurement forms if necessary.
70
-------
EMAP-SW-Streams Field Operations Manual, Section 6 (Stream Discharge), Rev. 5, September 1998 Page 5 of 10
Reviewed by (initial):
FIELD MEASUREMENT FORM - STREAMS (continued)
SITE NAME: A)/ti CxtSLg. DATE: 7 / /i"/ 97 VISIT: H1 D2
SITE ID: MAIA97- 9 ^ 9 TEAM ID (X): Hi D2 D3 D4 OS D6 D7 D8
STREAM DISCHARGE
E2 VELOCITY AREA
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
DlST. FROM
BANK (CM)
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MEASUREMENT
WIDTH (m)
DEPTH 1 (cm)
DEPTH 2 (cm)
DEPTH 3 (cm)
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Flag Codes: K = no measurement or observation made; U = suspect measurement or observation; Q = unacceptable QC check associated
with measurement; F1f F2, etc. =* miscellaneous flags assigned by each field crew. Explain all flags In comments section.
Rev. 06/02/97 (strvfldm.97)
FIELD MEASUREMENT FORM - STREAMS/RIVERS - 3
Figure 6-2. Field Measurement Form (page 2), showing data recorded for all three discharge
measurement procedures.
7-1
-------
EMAP-SW-Streams Reid Operations Manual. Section 6 (Stream Discharge), Rev. 5, September 1998 Page 6 of 10
Water Level
Bucket
Figure 6-3. Use of a portable weir in conjunction with a calibrated bucket to obtain an esti-
mate of stream discharge.
6.3 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.
72
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EMAP-SW-Streams Field Operations Manual, Section 6 (Stream Discharge), Rev. 5, September 1998 Page 7 of 10
TABLE 6-2. TIMED FILLING PROCEDURE FOR DETERMINING STREAM DISCHARGE
NOTE: If measuring discharge by this procedure will result in significant channel disturbance or will
stir up sediment, delay determining discharge until all biological and chemical measurement and
sampling activities have been completed.
1. Choose a cross-section that contains one or more natural spillways or plunges, or construct a
temporary one using on-site materials, or install a portable weir using a plastic sheet and on-
site materials.
2. Place an "X" in the "TIMED FILLING" box in the stream discharge section of the Field Measure-
ment 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 Field Measurement
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
73
-------
EMAP-SW-Streams Field Operations Manual. Section 6 (Stream Discharge), Rev. 5, September 1998 Page 8 of 10
The neutrally buoyant object procedure is described in Table 6-3. Examples of
suitable objects include oranges, 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-sec-
tion, and that is long enough to require 10 to 30 seconds for an object to float through it.
Select one to three cross-sections to represent the channel dimensions within the segment,
depending on the variability of width and/or depth. Determine the stream depth at 5 equally
spaced points at each cross-section. Three separate times, measure the time required for
the object to pass through the segment that includes all of the selected cross-sections.
Record data on the Field Measurement Form as shown in Figure 6-2.
6.4 EQUIPMENT AND SUPPLIES
Figure 6-4 shows the list of equipment and supplies necessary to measure stream
discharge. This checklist is similar to the checklist presented in Appendix A, which is used
at the base location (Section 3) to ensure that all of the required equipment is brought to the
stream. Use this checklist to ensure that equipment and supplies are organized and avail-
able at the stream site in order to conduct the activities efficiently.
6.5 LITERATURE CITED
Linsley, R.K., M.A. Kohler, and J.L.H. Paulhus. 1982. Hydrology for Engineers.
McGraw-Hill Book Co. New York.
Rantz, S.E. and others. 1982. Measurement and Computation of Streamflow: Volume 1.
Measurement of Stage and Discharge. U.S. Geological Survey Water-Supply Paper
2175.
74
-------
EMAP-SW-Streams Field Operations Manual, Section 6 (Stream Discharge). Rev. 5. September 1998 Page 9 of 10
TABLE 6-3. NEUTRALLY BUOYANT OBJECT PROCEDURE FOR DETERMINING
STREAM DISCHARGE
1. Place an "X" in the "NEUTRALLY BUOYANT OBJECT" box on the Field Measurement 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. Record the length
of the segment in the "FLOAT DISTANCE" field of the Field Measurement 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 (m) using a surveyor's rod or tape measure,
and record on the Field Measurement 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 depths (not the distances) in centimeters on the Field Mea-
surement Form.
5. Repeat Step 4 for the remaining cross-sections.
6. Use a stopwatch to determine the time required for the object to travel through the segment.
Record the time in the "FLOAT TIME" field of the Field Measurement Form.
7. Repeat Step 6 two more times. The float distance may differ somewhat for the three trials
75
-------
EMAP-SW-Streams Field Operations Manual, Section 6 (Stream Discharge), Rev. 5, September 1998 Page 10 of 10
EQUIPMENT AND SUPPLIES FOR STREAM DISCHARGE
QTY.
1
1
1
1
1
1
1
1
1
1
1 copy
1 set
ITEM
Surveyor's telescoping leveling rod
50-m fiberglass measuring tape and reel
Current velocity meter, probe, and operating manual
Top-set wading rod (metric scale) for use with current velocity meter
Portable Weir with 60° "V" notch (optional)
Plastic sheeting to use with weir
Plastic bucket (or similar container) with volume graduations
Stopwatch
Neutrally buoyant object (e.g., orange, small rubber ball, stick)
Covered clipboard
Soft (#2) lead pencils
Field Measurement Forms (1 per stream plus extras if needed for timed filling
procedure)
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for stream
discharge
Figure 6-4. Equipment and supply checklist for stream discharge.
76
-------
SECTION 7
PHYSICAL HABITAT CHARACTERIZATION
by
Philip R. Kaufmann1 and E. George Robison2,
,23
In the broad sense, physical habitat in streams includes all those physical attributes
that influence or provide sustenance to organisms within the stream. Stream physical
habitat varies naturally, as do biological characteristics; thus, expectations differ even in the
absence of anthropogenic disturbance. Within a given physiographic-climatic region,
stream drainage area and overall stream gradient are likely to be strong natural determi-
nants of many aspects of stream habitat, because of their influence on discharge, flood
stage, and stream power (the product of discharge times gradient). Summarizing the
habitat results of a workshop conducted by EMAP on stream monitoring design, Kaufmann
(1993) identified seven general physical habitat attributes important in influencing stream
ecology:
• Channel Dimensions
• Channel Gradient
• Channel Substrate Size and Type
• Habitat Complexity and Cover
• Riparian Vegetation Cover and Structure
• Anthropogenic Alterations
• Channel-Riparian Interaction
All of these attributes may be directly or indirectly altered by anthropogenic activities.
Nevertheless, their expected values tend to vary systematically with stream size (drainage
area) and overall gradient (as measured from topographic maps). The relationships of
specific physical habitat measurements described in this section to these seven attributes
1 Department of Fisheries and Wildlife, Oregon State University, c/o U.S. EPA, 200 SW 35th St., Corvallis, OR 97333.
2 Department of Forest Engineering, Oregon State University, Corvallis, OR 97331.
3 Current address: Oregon Department of Forestry, 2600 State St., Salem, OR 97310
77
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 2 of 42
are discussed by Kaufmann (1993). Aquatic macrophytes, riparian vegetation, and large
woody debris are included in this and other physical habitat assessments because of their
role in modifying habitat structure and light inputs, even though they are actually biological
measures. The field physical habitat measurements from this field habitat characterization
are used in the context of water chemistry, temperature, and other data sources (e.g.,
remote sensing of basin land use and land cover). The combined data analyses will more
comprehensively describe additional habitat attributes and larger scales of physical habitat
or human disturbance than are evaluated by the field assessment alone. A comprehensive
data analysis guide (Kaufmann et al., in preparation) discusses the detailed procedures
used to calculate metrics related to stream reach and riparian habitat quality from filed data
collected using the EMAP field protocols. This guide also discusses the precision associ-
ated with these measurements and metrics.
These procedures are intended for evaluating physical habitat in wadeable streams.
The EMAP field procedures are most efficiently applied during low flow conditions and
during times when terrestrial vegetation is active, but may be applied during other seasons
and higher flows except as limited by safety considerations. This collection of procedures is
designed for monitoring applications where robust, quantitative descriptions of reach-scale
habitat are desired, but time is limited. The qualitative nature of the habitat quality rank
scores produced by many currently available rapid habitat assessment methods (e.g., those
described in Section 14) have not been demonstrated, as yet, to meet the objectives of
EMAP, where more quantitative assessment is needed for site classification, trend interpre-
tation, and analysis of possible causes of biotic impairment.
The habitat characterization protocol developed for EMAP differs from other rapid
habitat assessment approaches (e.g., Plafkin etal., 1989, Rankin, 1995) by employing a
randomized, systematic spatial sampling design that minimizes bias in the placement and
positioning of measurements. Measures are taken over defined channel areas and these
sampling areas or points are placed systematically at spacings that are proportional to
baseflow channel width. This systematic sampling design scales the sampling reach length
and resolution in proportion to stream size. It also allows statistical and series analyses of
the data that are not possible under other designs. We strive to make the protocol objective
and repeatable by using easily learned, repeatable measures of physical habitat in place of
estimation techniques wherever possible. Where estimation is employed, we direct the
sampling team to estimate attributes that are otherwise measurable, rather than estimating
the quality or importance of the attribute to the biota or its importance as an indicator of
disturbance. We have included the more traditional visual classification of channel unit
78
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 3 of 42
scale habitat types because they have been useful in past studies and enhance comparabil-
ity with other work.
The time commitment to gain repeatability and precision is greater than that required
for more qualitative methods. In our field trials, two people typically complete the specified
channel, riparian, and discharge measurements in about three hours of field time (see
Section 2, Table 2-1). However, the time required can vary considerably with channel
characteristics. On streams up to about 4 meters wide with sparse woody debris, measure-
ments can be completed in less than two hours, whereas crews may require up to five hours
in large (>10 m wide), complex streams with abundant woody debris and deep water, if 100
width measurements are required. However, reducing the number of width measurements
from 100 to 21 locations on sample reaches limits time to < 4 hours even on large, complex
wadeable streams.
The procedures are employed on a sampling reach length 40 times its low flow
wetted width, as described in Section 4. Measurement points are systematically placed to
statistically represent the entire reach. Stream depth and wetted width are measured at
very tightly spaced intervals, whereas channel cross-section profiles, substrate, bank char-
acteristics and riparian vegetation structure are measured at larger spacings. Woody debris
is tallied along the full length of the sampling reach, and discharge is measured at one
location (see Section 6). 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.
7.1 COMPONENTS OF THE HABITAT CHARACTERIZATION
There are four different components of the EMAP physical habitat characterization
(Table 7-1), including stream discharge, which is described in Section 6. Measurements for
the remaining three components are recorded on 11 copies of a two-sided field form, plus
an a separate form for recording slope and bearing measurements. The thalweg profile is
a longitudinal survey of depth, habitat class, and presence of soft/small sediment at 100
equally spaced intervals (150 in streams less than 2.5 m wide) along the centerline between
the two ends of the sampling reach. "Thalweg" refers to the flow path of the deepest water
in a stream channel. Wetted width is measured at 21 equally spaced intervals. Data for the
second component, the woody debris tally, are recorded for each of 10 segments of
stream located between the 11 transects. The third component, the channel and riparian
characterization, includes measures and/or visual estimates of channel dimensions,
79
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EMAP-SW-Streams Field Operations Manual. Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 4 of 42
TABLE 7-1. COMPONENTS OF PHYSICAL HABITAT CHARACTERIZATION
Component
Description
Thalweg Profile:
(Section 7.4.1)
Woody Debris Tally:
(Section 7.4.2)
Channel and Riparian
Characterization:
(Section 7.5)
Discharge:
(see Section 6)
Measure maximum depth, classify habitat and pool-forming
features, and determine presence of soft sediment at 10-15
equally spaced intervals between each of 11 channel cross-sec-
tion transects (100 or 150 individual measurements along entire
reach).
Measure wetted width at 11 channel cross-section transects and
midway between them (21 measurements).
Between each of the channel cross sections, tally large woody
debris numbers within and above the bankfull channel according
to length and diameter classes (10 separate tallies).
At 11 cross-section transects placed at equal intervals along
reach length:
Measure: channel cross section dimensions, bank height,
bank undercut distance, bank angle, slope and compass
bearing (backsite), and riparian canopy density (densio-
meter).
Visually Estimate3: substrate size class and embeddedness;
areal cover class and type (e.g., woody trees) of riparian
vegetation in Canopy, Mid-Layer and Ground Cover; areal
cover class offish concealment features, aquatic macro-
phytes and filamentous algae.
Observe & Record3: human disturbances and their proximity
to the channel.
In medium and large streams (defined in Section 6) measure wa-
ter depth and velocity at 0.6 depth at 15 to 20 equally spaced
intervals across one carefully chosen channel cross-section.
In very small streams, measure discharge by timing the filling of a
bucket or timing the passage of a neutral buoyant object through
a segment whose cross-sectional area has been estimated.
* Substrate size class and embeddedness are estimated, and depth is measured for a total of 55 particles taken at 5 equally-
spaced points along each of 11 cross-section transects. 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.
substrate, fish cover, bank characteristics, riparian vegetation structure, and evidence of
human disturbance. These data are obtained at each of the 11 equally-spaced transects
established within the sampling reach. In addition, measurements of the stream slope and
compass bearing between stations are obtained, providing information necessary for calcu-
lating reach gradient, residual pool volume, and channel sinuosity.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization). Rev. 4, Sept. 1998 Page 5 of 42
7.2 HABITAT SAMPLING LOCATIONS WITHIN THE SAMPLING REACH
Measurements are made at two scales of resolution along the length of the reach; the
results are later aggregated and expressed for the entire reach, a third level of resolution.
Figure 7-1 illustrates the locations within the sampling reach where data for the different
components of the physical habitat characterization are obtained. We assess habitat over
stream reach lengths that are approximately 40 times their average wetted width at base-
flow, but not less than 150 m long. This allows us to adjust the sample reach length to
accommodate varying sizes of streams (see Section 2). Many of the channel and riparian
features are characterized on 11 cross-sections and pairs of riparian plots spaced at 4
channel-width intervals. The thalweg profile measurements must be spaced evenly over the
entire sampling reach. In addition, they must be sufficiently close together that they do not
"miss" deep areas and habitat units that are in a size range of about Va to Vz of the average
channel width. Follow these specifications for choosing the interval between thalweg profile
measurements:
• Channel Width < 2.5 m
• Channel Width 2.5-3.5 m
• Channel Width > 3.5 m
— interval = 1.0 m
— interval = 1.5 m
— interval = 0.01 * (reach length)
Following these guidelines, you will be making 150 evenly spaced thalweg profile measure-
ments in the smallest category of streams, 15 between each detailed channel cross section.
In all of the larger stream sizes, you will make 100 measurements, 10 between each cross
section. For practical reasons, we specify width measurements only at the 11 cross-section
transects and at the thalweg measurement points midway between each pair of transects (a
total of 21 wetted widths). If more resolution is desired, width measurements may be made
at all 100 or 150 thalweg profile locations.
7.3 LOGISTICS AND WORK FLOW
The four components (Table 7-1) of the habitat characterization are organized into
three grouped activities:
1. Thalweg Profile and Large Woody Debris Tally (Section 7.4).
Two people (the "geomorphs") proceed upstream from the
downstream end of the sampling reach (see Figure 7-1) mak-
ing observations and measurements at the chosen
81
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EMAP-SW-Streams Field Operations Manual. Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 6 of 42
Instream fish cover
Riparian Vegetation &
Human Disturbance
Upstream end of
sampling reach
Channel/Riparian
Cross section
Transect
Substrate and Channel
Measurements
Thalweg profile
intervals
Woody Debris Tally
(between transects)
Downstream end of
sampling reach
Figure 7-1. Sampling reach layout for physical habitat measurements (plan view).
82
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 7 of 42
increment spacing. One person is in the channel making width and depth
measurements, and determining whether soft/small sediment is present
under his/her staff. The other person records these measurements, classi-
fies the channel habitat, and tallies large woody debris. Each time this team
reaches a flag marking a new cross-section transect, they start filling out a
new copy of the Thalweg Profile and Woody Debris Form. They interrupt the
thalweg profile and woody debris tallying activities to complete data collection
at each cross-section transect as it comes.
3.
Channel/Riparian Cross-Sections (Section 7.5). One person proceeds
with the channel cross-section dimension, substrate, bank, and can-
opy cover measurements. The second person records those mea-
surements on the Channel/Riparian Cross-section and Thalweg Pro-
file Form while making visual estimates of riparian vegetation struc-
ture, instream fish cover,-and human disturbance specified on that
form. Slope and bearing are determined together by backsiting to the
previous transect. Intermediate flagging (of a different color) may
have to be used if the stream is extremely brushy, sinuous, or steep
to the point that you cannot site for slope and bearing measures be-
tween two adjacent transects. (Note that the crews could tally woody
debris while doing the backsite, rather than during the thalweg profile
measurements.)
Discharge (Section 6). Discharge measurements are made
after collecting the chemistry sample. They are done at a cho-
sen optimal cross section (but not necessarily at a trapsect)
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 chem-
ical and biological sampling has been completed.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1993 Page 8 of 42
7.4 THALWEG PROFILE AND LARGE WOODY DEBRIS MEASUREMENTS
7.4.1 Thalweg Profile
"Thalweg" refers to the flow path of the deepest water in a stream channel. The
thalweg profile is a longitudinal survey of maximum depth and several other selected char-
acteristics at 100 or 150 equally spaced points along the centerline of the stream between
the two ends of the stream reach. Data from the thalweg profile allows calculation of indi-
ces of residual pool volume, stream size, channel complexity, and the relative proportions of
habitat types such as riffles and pools. The EMAP-SW habitat assessment modifies tradi-
tional methods by proceeding upstream in the middle of the channel, rather than along the
thalweg itself (though each thalweg depth measurement is taken at the deepest point at
each incremental position). One field person walks upstream (wearing felt-soled waders)
carrying a fiberglass telescoping (1.5 to 7.5 m) surveyor's rod and a 1-m metric ruler (or a
calibrated rod or pole, such as a ski pole). A second person on the bank or in the stream
carries a clipboard with 11 copies of the field data form.
The procedure for obtaining thalweg profile measurements is presented in Table
7-2. Record data on the Thalweg Profile and Woody Debris Data Form as shown in Figure
7-2. Use the surveyor's rod and a metric ruler or calibrated rod or pole to make the required
depth and width measurements, and to measure off the distance between measurement
points as you proceed upstream. Ideally, every tenth thalweg measurement will bring you
within one increment spacing from the flag marking a new cross-section profile. The flag
will have been set previously by carefully taping along the channel, making the same bends
that you do while measuring the thalweg profile (refer to Figure 7-1). However, you may still
need to make minor adjustments to align each 10th measurement to be one thalweg incre-
ment short of the cross section. In streams with average widths smaller than 2.5m, you will
be making thalweg measurements at 1-meter increments. Because the minimum reach
length is set at 150 meters, there will be 15 measurements between each cross section.
Use the 5 extra lines on the thalweg profile portion of the data form (Figure 7-2) to record
these measurements.
It is very important that thalweg depths are obtained from all measurement points.
Missing depths at the ends of the sampling reach (e.g., due to the stream flowing into or out
of a culvert or under a-large pile of debris) can be tolerated, but those occurring in the
middle of the sampling reach are more difficult to deal with. Flag these missing measure-
ments using a "K" code and explain the reason for the missing measurements in the
84
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4. Sept. 1998 Page 9 of 42
TABLE 7-2. THALWEG PROFILE PROCEDURE
1. Determine the interval between measurement stations based on the wetted width used to
determine the length of the sampling reach.
For widths < 2.5 m, establish stations every 1 m.
For widths between 2.5 and 3.5 m, establish stations every 1.5 m
For widths > 3.5 m, establish stations at increments equal to 0.01 times the sampling
reach length.
2. Complete the header information on the thalweg profile and woody debris section of a Chan-
nel/Riparian Cross-section and Thalweg Profile Form, noting the transect pair (downstream to
upstream). Record the interval distance determined in Step 1 in the "INCREMENT" field on the
field data form.
• NOTE: If a side channel is present, and contains between 16 and 49% of the total flow,
establish secondary cross-section transects and thalweg measurement stations as nec-
essary. Use separate field data forms to record data for the side channel, and designate
each secondary transect as "X" followed by the primary transect letter (e.g., XA, XB, etc.).
Collect all channel and riparian cross-section measurements from the side channel.
3. Begin at the downstream end (station "0") of the first transect QTransect "A").
4. Measure the wetted width if you are at station "0", 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 widths up to about 3 meters, and to the nearest
5% for widths > 3 m. This is 0.2 m for widths of 4 to 6 m, 0.3 m for widths of 7 to 8 m, and 0.5
m for widths of 9 or 10 m, and so on. For dry and intermittent streams, where no water is in the
channel, record zeros for wetted width.
NOTE: If a mid-channel bar is present at a station where wetted width is measured,
measure the bar width and record it on the field data form.
5. At each thalweg profile station, use a meter ruler or a calibrated pole or rod to locate the
deepest point (the "thalweg"), which may not always be located at mid-channel. Measure the
thalweg depth to the nearest cm, and record it on the thalweg profile form. Read the depth on
the side of the ruler, rod, or pole to avoid inaccuracies due to the wave formed by the rod in
moving water.
NOTE: For dry and intermittent streams, where no water is in the channel, record zeros
for depth.
• NOTE: At stations where the thalweg is too deep to measure directly, stand in shallower
water and extend the surveyor's rod or calibrated rod or pole at an angle to reach the
thalweg. Determine the rod angle using the external scale of the clinometer. Leave the
depth reading for the station blank, and record a "U" flag. Record the water level on the
. rod and the rod angle in the comments section of the field data form;
(continued)
85
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 10 of 42
TABLE 7-2 (Continued)
6. At the point where the thalweg depth is determined, observe whether small, loose, soft sedi-
ments 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
staff. Record presence or absence in the "SOFT/SMALL SEDIMENT" field on the field data form.
7. Determine the channel unit code and pool forming element codes for the station. Record
these on the field data form using the standard codes provided. For dry and intermittent
streams, where no water is in the channel, record habitat type as dry channel (DR).
8. If the station cross-section intersects a mid-channel bar, Indicate the presence of the bar in the
"BAR WIDTH" field on the field data form.
9. 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.
10. Proceed upstream to the next station, and repeat Steps 4 through 9.
11. Repeat Steps 4 through 10 until you reach the next transect. Prepare a new Channel/Riparian
Cross-section and Thalweg Profile Form, then repeat Steps 2 through 10 for each of the reach
segments, until you reach the upstream end of the sampling reach (Transect "K").
86
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EMAP-SW-Streams Field Operations Manual. Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 11 of 42
Reviewed by (initial):
PHab: THALWEG PROFILE & WOODY DEBRIS FORM - STREAMS
SITE NAME: MlLL CtefK
SITE ID: MAlA97-_i_3_3
DATE: 1 1 lf\yi VISIT: E91 D2
TEAM ID (X): Hi D2 O3 D4 US D6 D7 D8
TRANSECT(X): LTA-B HB-C DC-D DD-E DE-F DF-G DG-H DH-I Dl-J Dj-K
THALWEG PROFILE
STA-
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COMMENTS
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Flag Codes: K=nomeasurementmade; U^suspect measurement F1.F2, etc.=mlsc. flags assigned by each field crew. Explain all nags In comments. 1= Measure
Bar Width at Station 0 and Mid-Station (5 or 7), X small column if bar present at the rest of the stations.
Rev. 06/02/97 (st_phct.97)
PHab: CHANNEL/RIPARIAN CROSS-SECTION & THALWEG PROFILE FORM -STREAMS -2
Figure 7-2. Thalweg Profile and Woody Debris Form.
87
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 12 of 42
comments section of the field data form. At points where a direct depth measurement
cannot be obtained, make your best estimate of the depth, record it on the field form, and
flag the value using a "U" code (for suspect measurement), explaining that it is an estimated
value in the comments section of the field data form. Where the thalweg points are too
deep for wading, measure the depth by extending the surveyor's rod at an angle to reach
the thalweg point. Record the water level on the rod, and the rod angle, as determined
using the external scale on the clinometer (vertical = 90°).
At every thalweg measurement increment, determine by sight or feel whether soft/
small sediment is 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
NOT the same as "Fines" described when determining the substrate particle sizes at the
cross-section transects (Section 7.5.2). For the thalweg profile, determine if soft/small
sediment deposits are readily obvious by feeling the bottom with your boot, the surveyor's
rod, or the calibrated rod or pole.
Wetted width is measured at each transect (station 0), and midway between tran-
sects (station 5 for larger streams having 100 measurement points, or station 7 for smaller
streams having 150 measurement points). The wetted width boundary is the point at which
substrate particles are no longer surrounded by free water.
While recording the width and depth measurements and the presence of soft/small
sediments, the second person chooses and records the habitat class and the pool forming
element codes (Table 7-3) applicable to each of the 100 (or 150) measurement points along
the length of the reach. These channel unit habitat classifications and pool-forming ele-
ments are modified from those of Bisson et al. (1982) and Frissell et al. (1986). The result-
ing database of traditional visual habitat classifications will provide a bridge of common
understanding with other studies. With the exception of backwater pools, channel unit scale
habitat classifications are to be made 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
7-3. Before being considered large enough to be identified as a channel-unit scale habitat
feature, the unit should be at least as long as the channel is wide. For instance, if there is a
'small deep (pool-like) area at the thalweg within a large riffle area, don't record it as a pool
unless it occupies an area about as wide or long as the channel is wide.
Mid-channel bars, islands, and side channels pose some problems for the sampler
conducting a thalweg profile and necessitate some guidance. Bars are defined here as
88
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EMAP-SW-Strearns Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 13 of 42
TABLE 7-3. CHANNEL UNIT AND POOL FORMING ELEMENT CATEGORIES
Channel Unit Habitat Classes3
Class (Code)
Description
Pools: Still water, low velocity, smooth, glassy surface, usually deep compared to other parts
of the channel:
Plunge Pool (PP) Pool at base of plunging cascade or falls.
Trench Pool (PT) Pool-like trench in the center of the stream
Lateral Scour Pool (PL) Pool scoured along a bank.
Backwater Pool (PB) Pool separated from main flow off the side of the channel.
Impoundment Pool (PD) Pool formed by impoundment above dam or constriction.
Pool (P)
Glide(GL)
Riffle (Ri)
Rapid (RA)
Cascade (CA)
Falls (FA)
Dry Channel (DR)
Pool (unspecified type).
Water moving slowly, with a smooth, unbroken surface. Low turbu-
lence.
Water moving, with small ripples, waves and eddies — waves not
breaking, surface tension not broken. Sound: "babbling", "gurgling".
Water movement rapid and turbulent, surface with intermittent white-
water with breaking waves. Sound: continuous rushing, but not as loud
as cascade.
Water movement rapid and very turbulent over steep channel bottom.
Most of the water surface is broken in short, irregular plunges, mostly
Whitewater. Sound: roaring.
Free falling water over a vertical or near vertical drop into plunge,
water turbulent and white over high falls. Sound: from splash to roar.
No water in the channel
(continued)
Note that in order for a channel habitat unit (other than a backwater pool) to be distinguished, it must be at least as wide
or long as the channel is wide.
89
-------
TABLE 7-3 (Continued)
Categories of Pool-forming Elements"
Code
N
W
R
B
F
WR, RW, RBW
Category
Not Applicable, Habitat Unit is not a pool
Large Woody Debris.
Rootwad
Boulder or Bedrock
Unknown cause (unseen fluvial processes)
Combinations
Other (describe in the comments section of field form)
6 Remember that most pools are formed at high flows, so you may need to look at features, such as large woody debris, that
are dry at baseflow, but still within the bankfull channel.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 15 of 42
mid-channel features below the bankfull flow mark that are dry during baseflow conditions
(see Section 7.5.3 for the definition of bankfull channel). Islands are mid-channel features
that are dry even when the stream is experiencing a bankfull flow. Both bars and islands
cause the stream to split into side channels. When a mid-channel bar is encountered along
the thalweg profile, it is noted on the field form and the active channel is considered to
include the bar. Therefore, the wetted width is measured as the distance between wetted
left and right banks. It is measured across and over mid-channel bars and boulders. If mid-
channel bars are present, record the bar width in the space provided.
If a mid-channel feature is as high as the surrounding flood plain, it is considered an
island. Treat side channels resulting from islands different from mid-channel bars. Handle
the ensuing side channel based on visual estimates of the percent of total flow within the
side channel as follows:
Less than 15%
16 to 49%
Indicate the presence of a side channel on the field data form.
Indicate the presence of a side channel on the field data form.
Establish a secondary transect across the side channel and
designate it as "X" plus the primary transect letter; e.g., XA).
Complete the detailed channel and riparian cross-section
measurements for the side channel, using a separate copy of
the field data form.
When a side channel occurs due to an island, reflect its presence with continuous entries in
the "Side Channel" field on the thalweg profile form (Figure 7-2). In addition, note the points
of divergence and confluence of the side channel in the comments section of the thalweg
profile form. Begin entries at the point where the side channel converges with the main
channel; note the side channel presence continuously until the upstream point where it
diverges. When doing width measures with a side channel separated by an island, include
only the width of the main channel in the measures at the time and then measure the side
channel width separately.
For dry and intermittent streams, where no water is in the channel at a thalweg
station, record zeros for depth and wetted width. Record the habitat type as dry channel
(DR).
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4. Sept. 1998 Page 16 of 42
7.4.2 Large Woody Debris Tally
Methods for large woody debris (LWD) measurement are a simplified adaptation of
those described by Robison and Beschta (1990). This component of the EMAP physical
habitat characterization allows quantitative estimates of the number, size, total volume and
distribution of wood within the stream reach. 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.).
The procedure for tallying LWD is presented in Table 7-4. The tally includes all
pieces of LWD that are at least partially in the baseflow channel, the "active channel" (flood
channel up to bankfull stage), or spanning above the active channel. The active (or "bank-
full") channel is defined as the channel that is filled by moderate sized flood events that
typically recur every one to two years. LWD in the active channel is tallied over the entire
length of the reach, including the area between the channel cross-section transects. As in
the thalweg profile, LWD measurements in the channel segment between each cross
section transect and the next one upstream are recorded on the first 10 thalweg profile and
woody debris forms (Figure 7-2). The location of the large end of each piece of LWD
determines the segment to which it is assigned.
First, tally all the pieces of LWD that are at least partially in the bankfull channel
(Figure 7-3, Zones 1 or 2). Then tally all the pieces of LWD that are not actually within the
bankfull channel, but are at least partially spanning (bridging) the bankfull channel (Figure
7-3, Zone 3). For both the Zone 1-2 wood and the Zone 3 LWD, the field form (Figure 7-2)
provides 12 entry boxes for tallying debris pieces visually estimated within three length and
four diameter class combinations. Each LWD piece is tallied in only one box. Pieces of
LWD that are not at least partially within Zones 1, 2, or 3 are not tallied.
For each LWD piece, first visually estimate its length and its large and small end
diameters in order to place it in one of the diameter and length categories. The diameter
class on the field form (Figure 7-2) refers to the large end diameter. Sometimes LWD is not
cylindrical, so it has no clear "diameter". In these cases visually estimate what the diameter
would be for a piece of wood with a circular cross section that would have the same volume.
When evaluating length, include only the part of the LWD piece that has a diameter greater
than 10 cm (4 in). Count each of the LWD pieces as one tally entry and include the whole
piece when assessing dimensions, even if part of it is in Zone 4 (outside of the bankfull
channel). For both the Zone 1-2 wood and the Zone 3 LWD, the field form (Figure 7-2)
provides 12 entry boxes for tallying debris pieces visually estimated within three length and
92
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 17 of 42
TABLE 7-4. PROCEDURE FOR TALLYING LARGE WOODY DEBRIS
Note: Tally pieces of large woody debris (LWD) within each segment of stream at the same time the
thalweg profile is being determined. Include all pieces whose large end is located within the seg-
ment in the tally.
1. Scan the stream segment between the two cross-section transects where thalweg profile
measurements are being made.
2. Tally all LWD pieces within the segment that are at least partially within the bankfull channel.
Determine if a piece is LWD (small end diameter ;>10 cm [4 in.]; length 2:1,5 m [5 ft.])
3. For each piece of LWD, determine the class based on the diameter of the large end (0.1 m
to < 0.3 m, 0.3 m to <0.6 m, 0.6 m to <0.8 m, or >0.8 m, and the class based on the length of
the piece (1.5m to <5.0m, 5m to <15m, or >15m).
• If the piece is not cylindrical, visually estimate what the diameter would be 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
greater than 10 cm (4 in)
4. Place a tally mark in the appropriate diameter * 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>15m).
6. Place a tally mark for each piece in the appropriate diameter * length class tally box in the
"PIECES BRIDGE ABOVE BANKFULL CHANNEL" section of the Thalweg Profile and Woody Debris
Form. .
7. After all pieces within the segment have been tallied, write the total number of pieces for each
diameter x length class in the small box at the lower right-hand corner of each tally box.
8. Repeat Steps 1 through 7 for the next stream segment, using a new Thalweg Profile and
Woody Debris Form.
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EMAP-SW-Streams Field Operations Manual. Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 18 of 42
BANKFULL CHANNEL WIDTH
ZONE
ZONE 4
WATER SURFACE AT
BANKFULL FLOW
WATER SURFACE
ZONE 2 AT BASEFLOW
Figure 7-3. Large woody debris influence zones (modified from Robison and Beschta, 1990)
four diameter class combinations. Each LWD piece is tallied in only one box. There are 12
size classes for wood at least partially in Zones 1 and 2, and 12 for wood partially within
Zone 3. Wood that is not at least partially within those zones is not tallied.
7.5 CHANNEL AND RIPARIAN MEASUREMENTS AT CROSS-SECTION TRANSECTS
7.5.1 Slope and Bearing
The slope, or gradient, of the stream reach is useful in three different ways. First,
the overall stream gradient is one of the major stream classification variables, giving an
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 19 of 42
indication of potential water velocities and stream power, which are in turn important con-
trols on aquatic habitat and sediment transport within the reach. Second, the spatial vari-
ability of stream gradient is a measure of habitat complexity, as reflected in the diversity of
water velocities and sediment sizes within the stream reach. Lastly, using methods de-
scribed by Stack (1989) and Robison and Kaufmann (1994), the water surface slope will
allow us to compute residual pool depths and volumes from the multiple depth and width
measurements taken in the thalweg profile (Section 7.4.1). Compass bearings between
cross section stations, along with the distance between stations, will allow us to estimate the
sinuosity of the channel (ratio of the length of the reach divided by the straight line distance
between the two reach ends).
Measure slope and bearing by "backsiting" downstream between transects (e.g.,
transect "B" to "A", "C" to "B", etc.) as shown in Figure 7-4. To measure the slope and
bearing between adjacent stations, use a clinometer, bearing compass, tripod, tripod exten-
sion, and flagging, following the procedure presented in Table 7-5. Record slope and
bearing data on the Slope and Bearing Form as shown in Figure 7-5.
Slope can also be measured by two people, each having a pole that is marked at the
same height. Alternatively, the second person can be "flagged" at the eye level of the
person doing the backsiting. Be sure that you mark your eye level on the other person or
on a separate pole beforehand while standing on level ground. Site to your eye level when
backsiting on your co-worker. If two marked poles are used, site from the mark on one pole
to the mark on the other. Also, be sure that the second person is standing (or holding the
marked pole) at the water's edge or in the same depth of water as you are. The intent is to
get a measure of the water surface slope, which may not necessarily be the same as the
bottom slope. The clinometer reads both percent slope and degrees of the slope angle; be
careful to read and record percent slope. Percent slope is the scale on the right-hand side
as you look through most clinometers. If using an Abnev Level, insure that you are reading
the scale marked "PERCENT." With the clinometer or the Abney level, verify this by com-
paring the two scales. Percent slope is always a higher number than degrees of slope
angle (e.g., 100% slope=45° angle). For slopes > 2%, read the clinometer to the nearest
0.5%. For slopes < 2%, read to the nearest 0.25%. If the clinometer reading is 0%, but
water is moving, record the slope as 0.1%. If the clinometer reading is 0% and water is not
moving, record the slope as 0%.
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EMAP-SW-Streams Field Operations Manual. Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 20 of 42
Slope (gradient) Measurement
Downstream Transect
Upstream Transect
!
ArtoA *
Tripod with flagging
at eye level
Stand at transect in same water
depth as tripod (may have to
move to side of channel as
shown here)
Bearing Measurement Between Transects
Supplemental
bearing point
Figure 7-4. Channel slope and bearing measurements.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 21 of 42
TABLE 7-5. PROCEDURE FOR OBTAINING SLOPE AND BEARING DATA
1. Stand in the center of the channel at the downstream cross-section transect. Determine if you
can see the center of the channel at the next cross-section transect upstream. If not, you will
have to take supplementary slope and bearing measurements.
2. Set up the tripod in shallow water or at the water's edge at the downstream cross-section
transect (or at a supplemental point). Standing tall in a position with your feet as near as
possible to the water surface elevation, set the tripod extension and mark it with a piece of
flagging at your eye level. Remember the depth of water in which you are standing when you
adjust the flagging to eye level.
On gradually sloped streams, it is advisable to use two people, each holding a pole
marked with flagging at the same height on both poles.
3. Walk upstream to the next cross-section transect. Find a place to stand at the upstream
transect (or at a supplemental point) that is at the same depth as where you stood at the
downstream transect when you set up the eye-level flagging.
If you have determined in Step 1 that supplemental measurements are required for this
segment, walk upstream to the furthest point where you can still see the center of the
channel at the downstream cross-section transect from the center of the channel. Mark
this location with a different color flagging than that used to mark the cross-section tran-
sects.
4. With the clinometer, site back downstream on your flagging at the downstream transect (or at
the supplementary point). Read and record the percent slope in the "MAIN" section on the
Slope and Bearing Form. Record the "PROPORTION" as 100%.
If two people are involved, place the base of each pole at the water level (or at the same
depth at each transect). Then site with the clinometer (or Abney level) from the flagged
height on upstream pole to the flagged height on the downstream pole.
If you are backsiting from a supplemental point, record the slope (%) and proportion (%)
of the stream segment that is included in the measurement in the appropriate "SUPPLE-
MENTAL" section of the Slope and Bearing Form.
5. Stand in the middle of the channel at upstream transect (or at a supplemental point), and site
back with your compass to the middle of the channel at the downstream transect (or at a
supplemental point). Record the bearing (degrees) in the "MAIN" section of the Slope and
Bearing Form.
If you are backsiting from a supplemental point, record the bearing in the appropriate
"SUPPLEMENTAL" section of the Slope and Bearing Form.
6. Retrieve the tripod from the downstream cross section station (or from the supplemental point)
and set it up at the next upstream transect (or at a supplemental point) as described in Step 2.
7. When you get to each new cross-section transect (or to a supplementary point), backsite on
the previous transect (or the supplementary point), repeat Steps 2 through 6 above.
97
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98
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 23 of 42
For bearing measurements, it does not matter whether or not you adjust your com-
pass bearings for magnetic declination, but it is important that you are consistent in the
use of magnetic or true bearings throughout all the measurements you make on a given
reach. Note in the comments section of the Slope and Bearing Form which type of bearings
you are taking. 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 ("supplementary") slope
and bearing points between a pair of cross-section transects if you do not have direct line-
of-site along (and within) the channel between stations (see Figure 7-4). This can happen if
brush is too heavy, or if there are sharp slope breaks or tight meander bends. Mark these
intermediate station locations with a different color of plastic flagging than used for the
cross-section transects to avoid confusion. Record these supplemental slope and bearing
measurements, along with the proportion of the stream segment between transects in-
cluded in each supplemental measurement, in the appropriate sections of the Slope and
Bearing Form (Figure 7-5). Note that the main slope and bearing observations are always
downstream of supplemental observations. Similarly, first supplemental observations are
always downstream of second supplemental observations.
7.5.2 Substrate Size and Channel Dimensions
Substrate size is one of the most important determinants of habitat character for fish
and macroinvertebrates in streams. Along with bedform (e.g., riffles and pools), substrate
influences the hydraulic roughness and consequently the range of water velocities in the
channel. It also influences the size range of interstices that provide living space and cover
for macroinvertebrates, salamanders, and sculpins. Substrate characteristics are often
sensitive indicators of the effects of human activities on streams. Decreases in the mean
substrate size and increases in the percentage of fine sediments, for example, may de-
stabilize channels and indicate changes in the rates of upland erosion and sediment supply
(Dietrich et al, 1989; Wilcock, 1998).
In the EMAP protocol, substrate size and embeddedness are evaluated at each of
the 11 cross-section transects (refer to Figure 7-1) using a combination of methods adapted
from those described by Wolman (1954), Bain et al. (1985), Platts et al. (1983), and Plafkin
et al. (1989). The basis of the protocol is a systematic selection of 5 substrate particles
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 24 of 42
from each of 11 cross-section transects (Figure 7-6). In the process of measuring substrate
particle sizes at each channel cross section, you also measure the wetted width of the
channel and the water depth at each substrate sample point. If the wetted channel is split
by a mid-channel bar (see Section 7.4.1), the five substrate points are centered between
the wetted width boundaries regardless of the mid-channel bar in between. Consequently,
substrate particles selected in some cross-sections may be "high and dry". For dry chan-
nels, make cross-section measurements across the unvegetated portion of the channel.
The distance you record to the right bank is the same as the wetted channel width.
(NOTE: this is the same value that is also recorded under "BANK MEASUREMENTS" on the
cross-section and thalweg profile data form [Section 7.5.3]). 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 7-6.
Record these measurements on the Channel/Riparian Cross-section and Thalweg Profile
Form as shown in Figure 7-7. 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 7-7) based on the
middle dimension of its length, width, and depth. This "median" dimension determines the
sieve size through which the particle can pass. Always distinguish "hardpan" from "fines",
coding hardpan as "HP". Similarly, always distinguish concrete or asphalt from bedrock;
denote these artificial substrates as "other" ("OT") and describe them in the comments
section of the field data form. Code and describe other artificial substrates (including metal,
tires, car bodies, etc.) in the same manner. When you record the size class as "OT" (other),
assign an "F"-series flag on the field data form (Figure 7-7) and describe the substrate type
in the comments section of the field form, as shown in Figure 7-2.
Examine particles larger than sand for surface stains, markings, and algal coatings
to estimate embeddedness of all particles in the 10 cm diameter circle around the substrate
sampling point. Embeddedness is the fraction of a particle's surface that is surrounded by
(embedded in) sand or finer sediments on the stream bottom. By definition, the embedded-
ness of sand, silt, clay, and muck is 100 percent, and the embeddedness of hardpan and
bedrock is 0 percent.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 25 of 42
Right
Bank
Figure 7-6. Substrate sampling cross-section.
7.5.3 Bank Characteristics
The procedure for obtaining bank and channel dimension measurements is pre-
sented in Table 7-7. Data are recorded in the "Bank Measurements" section of the Chan-
nel/Riparian Cross-section and Thalweg Profile Form as shown in Figure 7-7. Bank angle
and bank undercut distance are determined on the left and right banks at each cross sec-
tion transect. Other features include the wetted width of the channel (as determined in
Section 7.5.2), the width of exposed mid-channel bars of gravel or sand, estimated incision
height, and the estimated height and width of the channel at bankfull stage as described in
Section 7.4.2 and Figure 7-3. The "bankfull" or "active" channel is defined as the channel
that is filled by moderate-sized flood events that typically occur every one or two years.
Such flows do not generally overtop the channel banks to inundate the valley floodplain,
and are believed to control channel dimensions in most streams.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 26 of 42
TABLE 7-6. SUBSTRATE MEASUREMENT PROCEDURE
1. Fill in the header information on page 1 of a Channel/Riparian Cross-section and Thalweg
Profile Form. Indicate the cross-section transect. At the transect, extend the surveyor's rod
across the channel perpendicular to the flow, with the "zero" end at the left bank (facing down-
stream). If the channel is too wide for the rod, stretch the metric tape in the same manner.
2. Divide the wetted width of the channel by 4 to obtain the locations of the substrate measure-
ment points along the cross-section. In the "DisrLB" fields of the form, record the distances
corresponding to 0% (L.FT), 25% (LCiR), 50% (CTR), 75% (Rcra), and 100% (RGT) of the
measured wetted width.
3. Place your sharp-ended meter stick or calibrated pole at the "|_FT" location (0 m). Measure the
depth and record it on the field data form.
Entries for the water's edge 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, 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.
Code Size Class
RS Bedrock (Smooth)
RR Bedrock (Rough)
HP Hardpan
BL Boulders
CB Cobbles
GC Gravel (Coarse)
GF Gravel (Fine)
SA Sand
FN Fines
WD Wood
OT Other
Size Range (mm)
>4000
>4000
>250 to 4000
>64 to 250
>16to64
> 2 to 16
>0.06 to 2
<0.06
Regardless of Size
Regardless of Size
Description
Smooth surface rock bigger than a car
Rough surface rock bigger than a car
Firm, consolidated fine substrate
Basketball to car size
Tennis ball to basketball size
Marble to tennis ball size
Ladybug to marble size
Smaller than ladybug size, but visible as
particles - gritty between fingers
Silt Clay Muck (not gritty between fingers)
Wood & other organic particles
Concrete, metal, tires, car bodies etc.
(describe in comments)
5. For particles larger than sand, examine the surface for stains, markings, and algae. Estimate
the average percentage embeddedness of particles in the 10 cm circle around the measuring
rod. Record this value on the field data form. By definition, sand and fines are embedded 100
percent; bedrock and hardpan are embedded 0 percent.
6. Move successively to the next location along the cross section. Repeat steps 4 through 6 at
each location.
7. Repeat Steps 1 through 6 at each new cross section transect.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization). Rev. 4, Sept. 1998 Page 27 of 42
Figure 7-7. Channel/Riparian Cross-section and Thalweg Profile Form.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 28 of 42
TABLE 7-7. PROCEDURE FOR MEASURING BANK CHARACTERISTICS
1. To measure bank angle, lay the surveyor's rod or your meter ruler down against the left bank
(determined as you face downstream), with one end at the water's edge. Lay the clinometer
on the rod, 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 and Thalweg Profile Form.
A vertical bank is 90 degrees; undercut banks have angles >90 degrees approaching
180 degrees, and more gradually sloped banks have angles <90 degrees. To measure
bank angles >90 degrees, turn the clinometer (which only reads 0 to 90 degrees) over
and subtract the angle reading from 180 degrees.
2. If the bank is undercut, measure the horizontal distance of the undercutting to the nearest
0.01 m. Record the distance on the field data form. The undercut distance is the distance
from the water's edge out to the point where a vertical plumb line from the bank would hit the
water's surface.
Measure submerged undercuts by thrusting the rod into the undercut and reading the
length of the rod that is hidden by the undercutting.
3. Repeat Steps 1 and 2 on the right bank.
4. Hold the surveyor's rod vertical, with its base planted at the water's edge. Using the surveyor's
rod as a guide while examining both banks, estimate (by eye) 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). Record this value in the "INCISED HEIGHT" field
of the bank measurement section on the field data form.
5. Still holding the surveyor's rod as a guide, examine both banks to estimate 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.
transition from flood- and scour-tolerant vegetation to that which is relatively intolerant of
these conditions.
6. Record the wetted width value determined when locating substrate sampling points in the
"WETTED WIDTH" field in the bank measurement section of the field data form. Also determine
the bankfull channel width and the width of exposed mid-channel bars (if present). Record
these values in the "BANK MEASUREMENT" section of the field data form.
7. Repeat Steps 1 through 6 at each cross-section transect. Record data for each transect on a
separate field data form.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 29 of 42
If the channel is not greatly incised, bankfull channel height and incision height will
be the same. However, if the channel is incised greatly, the bankfull level will be below the
level of the first terrace of the valley floodplain, making bankfull channel height smaller than
incision height (Figure 7-8). You may need to look for evidence of recent flows (within
about one year) to distinguish bankfull and incision heights. In cases where the channel is
cutting a valley sideslope and has oversteepened and destabilized that slope, the bare
"cutbank" is not necessarily an indication of recent incision. Examine both banks to more
accurately determine channel downcutting.
Spotting the level of bankfull flow during baseflow conditions requires judgement and
practice; even then it remains somewhat subjective. In many cases there is an obvious
slope break that differentiates the channel from a relatively flat floodplain terrace higher
than the channel. Because scouring and inundation from bankfull flows are often frequent
enough to inhibit the growth of terrestrial vegetation, the bankfull channel may be evident by
a transition from exposed stream sediments to terrestrial vegetation. Similarly, it may be
identified by noting moss growth on rocks along the banks. Bankfull flow level may also be
seen by the presence of drift material caught on overhanging vegetation. However, in years
with large floods, this material may be much higher than other bankfull indicators. In these
cases, record the lower value, flag it, and also record the height of drift material in the
comments section of the field data form.
7.5.4 Canopy Cover Measurements
Riparian canopy cover over a stream is important not only in its role in moderating
stream temperatures through shading, but also as an indicator of conditions that control
bank stability and the potential for inputs of coarse and fine particulate organic material.
Organic inputs from riparian vegetation become food for stream organisms and structure to
create and maintain complex channel habitat.
Canopy cover over the stream is determined at each of the 11 cross-section tran-
sects. A Convex Spherical Densiometer (model B).is used (Lemmon, 1957). The densi-
ometer must be taped exactly as shown in Figure 7-9 to limit the number of square grid
intersections to 17. Densiometer readings can range from 0 (no canopy cover) to 17 (maxi-
mum canopy cover). Six measurements are obtained at each cross-section transect (four
measurements in four directions at mid-channel and one at each bank). The mid-channel
measurements are used to estimate canopy cover over the channel. The two bank mea-
surements complement your visual estimates of vegetation structure and cover within the
105
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EMAP-SW-Streams Field Operations Manual. Section 7 (Physical Habitat Characterization), Rev. 4. Sept. 1998 Page 30 of 42
A. Channel not "Incised"
Downcutting over Geologic Time
Stream - No recent
incision. Bankfull
Level at Valley
Bottom
First Terrace on Valley Bottom
Second Terrace
B. Channel "Incised"
Downcutting over Geologic Time
Recent incision: Bankfull Level
below first terrace of Valley
Bottom
First Terrace on
Valley Bottom
Second Terrace !
Valley Fill
Figure 7-8. Schematic showing bankfull channel and incision for channels. (A) not recently
incised, and (B) recently incised into valley bottom. Note level of bankfull stage relative to
elevation of first terrace on valley bottom (Stick figure included for scale).
riparian zone itself (Section 7.5.5), and are particularly important in wide streams, where
riparian canopy may not be detected by the densiometer when standing midstream.
The procedure for obtaining canopy cover data is presented in Table 7-8. Densi-
ometer measurements are taken at 0.3 m (1 ft) above the water surface, rather than at
waist level, to (1) avoid errors because people differ in height; (2) avoid errors from standing
in water of varying depths; and (3) include low overhanging vegetation more consistently in
the estimates of cover. Hold the densiometer level (using the bubble level) 0.3 m above the
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization). Rev. 4, Sept. 1998 Page 31 of 42
TAPE
BUBBLE LEVELED*
Figure 7-9. Schematic of modified convex spherical canopy densiometer (From Mulvey et al.,
1992). In this example, 10 of the 17 intersections show canopy cover, giving a densiometer reading
of 10. Note proper positioning with the bubble leveled and face reflected at the apex of the "V."
water surface with your face reflected just below the apex of the taped "V", as shown in
Figure 7-9. Concentrate on the 17 points of grid intersection on the densiometer that lie
within the taped "V". If the reflection of a tree or high branch or leaf overlies any of the
intersection points, that particular intersection is counted as having cover. For each of the
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EMAP-SW-Streatns Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 32 of 42
TABLE 7-8. PROCEDURE FOR CANOPY COVER MEASUREMENTS
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. Hold the densiometer level
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 "CENlIp" 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. Repeat Steps 2 and 3 again, this time facing the bank while standing first at the left bank, then
the right bank. Record the values in the "L.FT" and "RGT" fields of the field data form.
7. Repeat Steps 1 through 6 at each cross-section transect. Record data for each transect on a
separate field data form.
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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 and Thalweg Profile Form as shown in (Figure 7-7).
7.5.5 Riparian Vegetation Structure
The previous section (7.5.4) described methods for quantifying the cover of canopy
over the stream channel. The following visual estimation procedures supplement those
measurements with a semi-quantitative evaluation of the type and amount of various types
of riparian vegetation. These data are used to evaluate the health and level of disturbance
of the stream corridor. They also provide an indication of the present and future potential
for various types of organic inputs and shading.
Observations to assess riparian vegetation apply to the riparian area upstream 5
meters and downstream 5 meters from each of the 11 cross-section transects (refer to
Figure 7-1). They include the visible area from the stream back a distance of 10m (-30 ft)
shoreward from both the left and right banks, creating a10mx10m riparian plot on each
side of the stream (Figure 7-10). The riparian plot dimensions are estimated, not mea-
sured. On steeply sloping channel margins, the 10 m * 10m plot boundaries are defined
as if they were projected down from an aerial view. If the wetted channel is split by a mid-
channel bar, the bank and riparian measurements are made at each side of the channel,
not the bar.
Table 7-9 presents the procedure for characterizing riparian vegetation structure and
composition. Figure 7-7 illustrates how measurement data are recorded in the "VISUAL
RIPARIAN ESTIMATES" section of the Channel/Riparian Cross-section and Thalweg Profile
Form. Conceptually divide the riparian vegetation into three layers: a CANOPY LAYER
(> 5 m high), an UNDERSTORY (0.5 to 5 m high), and a GROUND COVER layer (< 0.5 m
high). Note that several vegetation types (e.g., grasses or woody shrubs) can potentially
occur in more than one layer. Similarly note that some things other than vegetation are
possible entries for the "Ground Cover" layer (e.g.,. barren ground).
. Before estimating the areal coverage of the vegetation layers, record the type of
vegetation (Deciduous, Coniferous, broadleaf Evergreen, Mixed, or None) in each of the two
taller layers (Canopy and Understory). Consider the layer "Mixed" if more than 10% of the
areal coverage is made up of the alternate vegetation type.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 34 of 42
10m
Flow
10m
10m
RIPARIAN
PLOT
(Left Bank)
Cross-section Transect
5m I
5 m
Instream Fish
Cover Plot
RIPARIAN
PLOT
(Right Bank)
10m
Figure 7-10. Boundaries for visual estimation of riparian vegetation, fish cover, and human
influences.
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%),
110
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TABLE 7-9. PROCEDURE FOR CHARACTERIZING RIPARIAN VEGETATION STRUCTURE
1. Standing in mid-channel at a cross-section transect, 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 three layers: a
CANOPY LAYER (>5m high), an UNDERSTORY (0.5 to 5 m high), and a GROUND COVER
layer (<0.5 m high).
4. Within this 10 m x 10 m area, determine the dominant vegetation type for the CANOPY LAYER
(vegetation > 5 m high) as either Deciduous, Coniferous, broadleaf Evergreen, Mixed, or None.
Consider the layer "Mixed" if more than 10% of the areal coverage is made up of the alternate
vegetation type. Indicate the appropriate vegetation type in the "VISUAL RIPARIAN ESTIMATES"
section of the Channel/Riparian Cross-section and Thalweg Profile Form.
5. Determine separately the areal cover class of large trees (> 0.3 m [1 ft] diameter at breast
height [DBH]) and small trees (< 0.3 m DBH) within the canopy layer. Estimate areal cover as
the amount of shadow that would be cast by a particular layer alone if the sun were directly
overhead. Record the appropriate cover class on the field data form ("0"=absent: zero cover,
"1"=sparse: <10%, "2"=moderate: 10-40%, "3"=heavy: 40-75%, or "4"=very heavy: >75%).
6. Look at the UNDERSTORY layer (vegetation between 0.5 and 5 m high). Determine the
dominant vegetation type for the understory layer as described in Step 4 for the canopy layer.
7. Determine the areal cover class for woody shrubs and saplings separately from non-woody
vegetation within the understory, as described in Step 5 for the canopy layer.
8. Look at the GROUND COVER layer (vegetation < 0.5 m high). Determine the areal cover
class for woody shrubs and seedlings, non-woody vegetation, and the amount of bare ground
present as described in Step 5 for large canopy trees.
9. Repeat Steps 1 through 8 for the right bank.
10. Repeat Steps 1 through 9 for all cross-section transects, using a separate field data form for
each transect.
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and "very heavy" (>75%). These cover classes and their corresponding codes are shown
on the field data form (Figure 7-6). When rating vegetation cover types, 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").
Two heavy classes with 40-75% cover can both be rated "3".
7.5.6 Instream Fish Cover, Algae, and Aquatic Macrophytes
This portion of the EMAP physical habitat protocol is a visual estimation procedure
that semi-quantitatively evaluates the type and amount of important types of cover for fish
and macroinvertebrates. Alone and in combination with other metrics, this information is
used to assess habitat complexity, fish cover, and channel disturbance.
The procedure to estimate the types and amounts of instream fish cover is outlined
in Table 7-10. Data are recorded in the "Fish Cover/Other" section of the Channel /Riparian
Cross-section and Thalweg Profile Form as shown in Figure 7-7. Estimate the areal cover
of all of the fish cover and other listed features that are in the water and on the banks 5
meters upstream and downstream of the cross-section (see Figure 7-10). The areal cover
classes of fish concealment and other features are the same as those described for riparian
vegetation (Section 7.5.5).
The entry "Filamentous algae" refers to long streaming algae that often occur in slow
moving waters. "Aquatic macrophytes" are water-loving plants, including mosses, in the
stream that could provide cover for fish or macroinvertebrates. If the stream channel con-
tains live wetland grasses, include these as 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 7.4]). "Brush/woody debris" refers
to smaller wood pieces that primarily affect cover but not morphology. "Overhanging vegeta-
tion" includes tree branches, brush, twigs, or other small debris that is not in the water but is
close to the stream (within 1 m of the surface) and provides potential cover. "Boulders" are
typically basketball- to car-sized particles. "Artificial structures" include those designed for
fish habitat enhancement, as well as in-channel structures discarded (e.g., cars or tires) or
purposefully placed for diversion, impoundment, channel stabilization, or other purposes.
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TABLE 7-10. PROCEDURE FOR ESTIMATING INSTREAM FISH COVER
1. Standing mid-channel at a cross-section transect, estimate a 5m distance upstream and
downstream (10m total length).
2. Examine the water and the banks within the 10-m segment of stream for the following features
and types offish cover: filamentous algae, aquatic macrophytes, large woody debris, brush
and small woody debris, 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 and Thalweg Profile Form
("0"=absent: zero cover, "1"=sparse: <10%, "2"=moderate: 10-40%, "3"=heavy: 40-75%, or
"4"=very heavy: >75%).
4. Repeat Steps 1 through 3 at each cross-section transect, recording data from each transect on
a separate field data form.
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7.5.7 Human Influence
The field evaluation of the presence and proximity of various important types of
human land use activities in the stream riparian area is used in combination with mapped
watershed land use information to assess the potential degree of disturbance of the sample
stream reaches.
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 7-11. Relate your observations and proxim-
ity evaluations to the stream and riparian area within 5 m upstream and 5 m downstream
from the station (Figure 7-10). Four proximity classes are used: In the stream or on the
bank within 5 m upstream or downstream of the cross-section transect, present within the
10 m * 10 m riparian plot but not in the stream or on the bank, present outside of the ripar-
ian plot, and absent. Record data on the Channel/Riparian Cross-section and Thalweg
Profile Form as shown in Figure 7-6. If a disturbance is within more than one proximity
class, record the one that is closest to the stream (e.g., "C" takes precedence over "P").
A particular influence may be observed outside of more than one riparian observa-
tion plot (e.g., at both transects "D" and "E"). Record it as present at every transect where
you can see it without having to site through another transect or its 10 m * 10 m riparian
plot.
7.6 EQUIPMENT AND SUPPLIES
Figure 7-11 lists the equipment and supplies required to conduct all the activities
described for characterizing physical habitat. This checklist is similar to the checklist pre-
sented in Appendix A, which is used at the base location (Section 3) to ensure that all of the
required equipment is brought to the stream. Use this checklist to ensure that equipment
and supplies are organized and available at the stream site in order to conduct the activities
efficiently.
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TABLE 7-11. PROCEDURE FOR ESTIMATING HUMAN INFLUENCE
1. Standing mid-channel at a cross-section transect, look toward the left bank (left when facing
downstream), and estimate a 5m distance upstream and downstream (10m total length).
Also, estimate a distance of 10 m back into the riparian zone to define a riparian plot area.
2. Examine the channel, bank and riparian plot area adjacent to the defined stream segment
for the following human influences: (1) walls, dikes, revetments, riprap, and dams; (2) build-
ings; (3) pavement (e.g., parking lot, foundation); (4) roads or railroads, (5) inlet or outlet
pipes; (6) landfills or trash (e.g., cans, bottles, trash heaps); (7) parks or maintained lawns;
(8) row crops; (9) pastures, rangeland, or hay fields; (10) logging; and (11) mining (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 site through another
transect or its 10 m *10 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 and
Thalweg Profile Form. Proximity classes are:
B ("Bank") Present within the defined 10m stream segment and lo-
cated in the stream or on the stream bank.
• C ("Close") Present within the 10 x 10 m riparian plot area, but away from the
bank.
P ("Present") Present, but outside the riparian plot area.
O ("Absent") Not present within or adjacent to thelO 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, recording data for each transect
on a separate field form.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 40 of 42
EQUIPMENT AND SUPPLIES FOR PHYSICAL HABITAT
QTY.
1
1
1
1
1
2
1
1 roll ea.
1
1
1 or 2
2 pair
11 plus
extras
1 plus
extras
1 copy
1 set
Item
Surveyor's telescoping leveling rod (round profile, metric scale, 7.5m extended)
50-m fiberglass measuring tape & reel
Hip chain (metric) for measuring reach lengths (Optional)
Clinometer (or Abney level) with percent and degree scales.
Lightweight telescoping camera tripod (necessary only if slope measurements
are being determined by one person)
14-inch diameter PVC pipe, 2-3 m long, each marked at the same height (for use
in slope determinations involving two persons)
Meter stick. Alternatively, a short (1-2 m) rod or pole (e.g., a ski pole) with cm
markings for thalweg measurements, or the PVC pipe described for slope deter-
minations can be used
Colored surveyor's plastic flagging (2 colors)
Convex spherical canopy densiometer (Lemmon Model B), modified with taped
"V"
Bearing compass (Backpacking type)
Fisherman's vest with lots of pockets and snap fittings. Used at least by person
conducting the in-channel measurements to hold the various measurement
equipment (densiometer, clinometer, compass, etc.). Useful for both team mem-
bers involved with physical habitat characterization.
Chest waders with felt-soled boots for safety and speed if waders are the neo-
prene "stocking" type. Hip waders can be used in shallower streams.
Covered clipboards (lightweight, with strap or lanyard to hang around neck)
Soft (#2) lead pencils (mechanical are acceptable)
Channel/Riparian Cross-section & Thalweg Profile Forms
Slope and Bearing Forms
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for physical
habitat characterization
Figure 7-11. Checklist of equipment and supplies for physical habitat.
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4. Sept. 1998 Page 41 of 42
7.6 LITERATURE CITED
Bain, M.B., J.T. Finn, and H.E. Booke. 1985. Quantifying stream substrate for habitat
analysis studies. North American Journal of Fisheries Management 5:499-500.
Bisson, P.A., J.L. Neilsen, R.A. Palmason, and L.E. Grove. 1982. A system of naming
habitat types in small streams, with examples of habitat utilizations by salmonids dur-
ing low stream flow. pp. 62-73 ]N: N.B. Armantrout (ed.). Acquisition and Utilization of
Aquatic Habitat Inventory Information. Symposium Proceedings, October 18-30, 1981,
Portland, Oregon. The Hague Publishing, Billings, Montana.
Dietrich, W.E., J.W. Kirchner, H. Ikeda, and F. Iseya. 1989. Sediment supply and the
development of the coarse surface layer in gravel bed rivers. Nature 340:215-217.
Frissell, C.A., W.J. Liss, C.E. Warren, and M.D. Hurley. 1986. A hierarchical framework for
stream habitat classification: viewing streams in a watershed context. Environmental
Management 10(2): 199-214.
Kaufmann, P.R. (ed.). 1993. Physical Habitat, pp. 59-69 IN: R.M. Hughes (ed.). Stream
Indicator and Design Workshop. EPA/600/R-93/138. U.S. Environmental Protection
Agency, Corvallis, Oregon.
Kaufmann, P.R., P. Levine, E.G. Robison, C. Seeliger, and D.V. Peck. In preparation.
Quantifying Physical Habitat in Wadeable Streams. Environmental Monitoring and
Assessment Program, U.S. Environmental Protection Agency, Corvallis, Oregon.
Lemmon, P.E. 1957. A new instrument for measuring forest overstory density. Journal of
Forestry 55(9):667-669.
Linsley, R.K., M.A. Kohler, and J.LH. Paulhus. 1982. Hydrology for Engineers. McGraw-
Hill Book Co. New York, NY. 508 p.
Mulvey, M., L. Caton, and R. Hafele. 1992. Oregon Nonpoint Source Monitoring Protocols
Stream Bioassessment Field Manual for Macroinvertebrates and Habitat Assessment.
Oregon Department of Environmental Quality, Laboratory Biomonitoring Section. 1712
S.W. 11thAve. Portland, Oregon, 97201. 40 p.
117
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EMAP-SW-Streams Field Operations Manual, Section 7 (Physical Habitat Characterization), Rev. 4, Sept. 1998 Page 42 of 42
Plafkin, J.L., M.T. Barbour, K.D. Porter, S.K. Gross, R.M. Hughes. 1989. Rapid Bioassess-
ment Protocols for Use in Streams and Rivers: Benthic Macroinvertebrates and Fish.
EPA/440/4-89/001. U.S. Environmental Protection Agency, Assessment and Water-
shed Protection Division, Washington, D.C.
Platts, W.S., W.F. Megahan, and G.W. Minshall. 1983. Methods for Evaluating Stream,
Riparian, and Biotic Conditions. USDA Forest Service General Technical Report
INT-183. 71 p.
Robison, E.G. and R.L. Beschta. 1990. Characteristics of coarse woody debris for several
coastal streams of southeast Alaska, USA. Canadian Journal of Fisheries and Aquatic
Sciences 47(9): 1684-1693.
Robison, E.G. and P.R. Kaufmann. 1994. Evaluating two objective techniques to define
pools in small streams, pp. 659-668, ]N: R.A. Marston and V.A. Hasfurther (eds.).
Effects of Human Induced Changes on Hydrologic Systems. Summer Symposium
proceedings, American Water Resources Association,. June 26-29, 1994, Jackson
Hole, Wyoming. 1182 p.
Stack, B.R. 1989. Factors Influencing Pool Morphology in Oregon Coastal Streams. M.S.
Thesis, Oregon State University. 109 p.
Wilcock, P.R. 1988. Two-fraction model of initial sediment motion in gravel-bed rivers.
Science 280:410-412.
Wolman, M.G. 1954. A method of sampling coarse river-bed material. Transactions of the
American Geophysical Union 35(6):951-956.
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SECTION 8
PERIPHYTON
by
Brian H. Hill1
Periphyton are algae, fungi, bacteria, protozoa, and associated organic matter
associated with channel substrates. Periphyton are useful indicators of environmental
condition because they respond rapidly and are sensitive to a number of anthropogenic
disturbances, including habitat destruction, contamination by nutrients, metals, herbicides,
hydrocarbons, and acidification.
The "biomorphs" (refer to Figure 2-1) collect periphyton samples after completing
activities pertaining to water chemistry (Section 5) and discharge (Section 6). Periphyton
samples are collected from erosional and depositional habitats located at each of the nine
interior cross-section transects (transects "B" through "J") established within the sampling
reach (Section 4). Periphyton samples are collected at each transect at the same time as
sediment samples (Section 9) and benthic macroinvertebrate samples (Section 11). At
each stream, composite "index" samples of periphyton are prepared for erosional and
depositional habitats. At the completion of the day's sampling activities, but before leaving
the stream, four types of laboratory samples are prepared from each composite index
sample.
8.1 SAMPLE COLLECTION
The general scheme for collecting periphyton samples from the sampling reach at
each stream is illustrated in Figure 8-1. The procedure for collecting periphyton samples is
presented in Table 8-1. At each transect, samples are collected from an assigned sam-
pling point (left, center, or right). Sampling points at each transect may have been assigned
when the sampling reach was laid out (Figure 8-1; refer also to Section 4; Table 4-3). If not,
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin L. King Dr.,
Cincinnati, OH 45268.
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton), Rev. 4, September 1998 Page 2 of 14
CROSS SECTION TRANSECTS (A to K)
TRANSECT SAMPLES (9 total)
Collected from assigned sampling point (left, center, or right) on each transect
EROSIONAL SAMPLE
Attached periphyton collected from
12 cm2 area of rock(s) by scrubbing
and/or scraping
DEPOSITIONAL SAMPLE
Top 1 cm of sediment from a 12 cm2
area collected in 60-mL syringe
COMPOSITE TRANSECT SAMPLES
BY HABITAT TYPE
COMPOSITE INDEX SAMPLES
(Erosional and Depositional)
ID/ENUMERATION SAMPLE
50-mL aliquot
Preserve with 10% formalin (2 mL)
ACID/ALKALINE
PHOSPHATASE ACTIVITY
(APA) SAMPLE
BIOMASS SAMPLE
Filter 25-mL aliquot (pre-weighed
glass-fiber filter)
Store filter at -20 C
CHLOROPHYLL SAMPLE
Filter 25-mL aliquot
(glass-fiber filter)
Store filter at -20 C
Figure 8-1. Index sampling design for periphyton.
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EMAP-SW-Streams Field Operations Manual. Section 8 (Periphyton), Rev. 4. September 1998 Page 3 of 14
TABLE 8-1. PROCEDURE FOR COLLECTING COMPOSITE INDEX SAMPLES
OF PERIPHYTON
1. . Starting with Transect "B", determine if the assigned sampling point (Left, Center, or Right) is
located in an erosional (riffle) habitat or a slack water (pool) habitat. Collect a single sample at
the point using the appropriate procedure in Step 2 below.
If the sampling points were not assigned previously when laying out the sampling reach,
proceed to Transect "B". Roll a die to determine if it is a left (L), center (C), or right (R) sam-
pling point for collecting periphyton and benthic macroinvertebrate samples. A roll of 1 or 2
indicates L, 3 or 4 indicates C, and 5 or 6 indicates R (or use a digital wristwatch and glance at
the last digit (1-3=L, 4-6=C, 7-9=R). Mark L, C, or R on the transect flagging. Assign sampling
points at each successive transect in order as L, C, R after the first random selection.
2A. Erosional habitats:
(1) Collect a sample of substrate (rock or wood) that is small enough (< 15 cm diameter) and
can be easily removed from the stream. Place the substrate in a plastic funnel which
drains into a 500-mL plastic bottle with volume graduations marked on it and labeled
"EROSIONAL."
(2) 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.
(3) Fill a wash bottle with stream water. Using a minimal volume of water from this bottle,
wash the dislodged periphyton from the funnel into the 500-mL bottle.
2B. Depositional habitats:
(1) Use the area delimiter to confine a 12-cm2 area of soft sediments.
(2) Vacuum the top 1 cm of sediments from within the delimited area into a 60-mL syringe.
(3) Empty the syringe into a 500-mL plastic bottle with volume graduations marked on it and
labeled "DEPOSITIONAL."
3. Repeat Steps land 2 for transects "C" through "J". Place the sample collected at each sam-
pling site into its appropriate 500-mL bottle ("EROSIONAL" or "DEPOSITIONAL") to produce
the composite index sample for each habitat type:
4. After samples have been collected from all nine transects, mix each 500-mL bottle thoroughly.
For each composite sample, place an "X" in the appropriate habitat type box ("riffle" for ero-
sional; "pool" for depositional) and record the total estimated volume of the composite sample
in the periphyton section of the Sample Collection Form.
121
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton), Rev. 4, September 1998 Page 4 of 14
the sampling point at Transect "B" is assigned at random using a die or other suitable
means (e.g., digital watch). Once the first sampling point is determined, depositional sam-
ple, depending on whether the habitat at the site is flowing water (e.g., a riffle or run) or
slack water (e.g., a pool). Composite samples for erosional and depositional habitats are
prepared by combining the individual transect samples as they are collected from each
habitat into separate plastic bottles. The habitat type and volume of each composite sam-
ple are recorded on the Sample Collection Form as shown in Figure 8-2.
8.2 PREPARATION OF LABORATORY SAMPLES
Four different types of laboratory samples are prepared from each of the two com-
posite index samples: an ID/enumeration sample (to determine taxonomic composition and
relative abundances), a chlorophyll sample, a biomass sample (for ash-free dry mass
[AFDM]), and an acid/alkaline phosphatase activity (APA) sample. All the sample contain-
ers required for an individual stream should be sealed in plastic bags until use (see Section
3) to avoid external sources of contamination (e.g., dust, dirt, or mud) that are present at
streamside.
A set of completed periphyton sample labels is shown in Figure 8-3. All labels in a
set have the same sample ID number. Circle the habitat type of the composite index sam-
ple and the appropriate type of sample (chlorophyll, biomass, etc.) on each label. Attach
completed labels to the appropriate containers and cover with clear tape. When attaching
the completed labels, avoid covering any volume graduations and markings on the con-
tainer.
8.2.1 ID/Enumeration Sample
Prepare the ID/Enumeration samples as 50-mL aliquots from each composite index
sample, following the procedure presented in Table 8-2. Preserve each sample with 2 ml_
of 10% formalin. For each habitat type (riffle and pool), record the ID number (barcode)
from each sample container label and the total volume of the sample in the appropriate
fields on the Sample Collection Form as shown in Figure 8-2. Store the preserved samples
upright in a container containing absorbent material, according to the guidelines provided
for handling formalin-preserved samples.
122
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton), Rev. 4, September 1998 Page 5 of 14
Reviewed by (initial):
SAMPLE COLLECTION FORM - STREAMS
SITE NAME: Ml LI, CjfeffK
DATE: 7 / //"/ 97 VISIT: 01 D2
SITEID: MAIA97-_2__2__2 TEAMID(X): H1 D2 D3 D4 D5 D6 D7
D8
COMPOSITE BENTHOS SAMPLES
SAMPLE ID
(BARCODE)
HABITAT
(XONE)
R P
3. jL 1 O O ^L *•'
A a. ? o o _g, *
STATION A
RIFFLE OR POOL -
{XONE>-
LEFT, CENTER, OR
RIGHT -
(XONE)-
B C
No.
OF
JARS
A
/
FLAG COMMENTS
D E F G H
SIR OR DR HR DR DR DR
DP HP HP DP HP HP HP
DL EL. DL DL HL DL DL
DC DC HC DC DC HC DC
BR DR DR HR DR DR HR
I J
DR HR
BP DP
HL DL
DC HC
DR DR
K
COMPOSITE PERIPHYTON SAMPLES HABITAT TYPE (X)- H RIFFLE D POOL D OTHER
SAMPLE ID (BARCODE) -
ASSEMBLAGE ID
(50-MLTUBE)
SUB. SAMPLE VOL.
5~ O ML
_5,_3_ _?.._£ J?L-iL- COMPOSITE VOLUME - S. O O ML
CHLOROPHYLL
(GF/F FILTER)
VOL. FILTERED
A_j
L_ML
COMPOSITE PERIPHYTON SAMPLES
SAMPLE ID (BARCODE) -
ASSEMBLAGE ID
(50-ML TUBE)
SUB. SAMPLE VOL.
S~ O ML
A.-2_j2_je.o__l_
CHLOROPHYLL
(GF/F FILTER)
VOL. FILTERED
A.1
:LML
BlOMASS
(TARED FILTER)
FILTER No. VOL. FILTERED
979 a f ML
APA SAMPLE
(50-ML TUBE)
SUB. SAMPLE VOL.
S~ O ML
HABITAT TYPE (X)- n RIFFLE BPOOL D OTHER
• COMPOSITE VOLUME - "3 O & ML
BlOMASS
(TARED FILTER)
FILTER No. VOL. FILTERED
lOOt Ufa.
APA SAMPLE
(50-ML TUBE)
SUB.SAKPLEVOL.
STJ2.ML
COMMENTS:
.
Flag codes: K= Samplo not collected; U= Suspect sample; F1, F2, etc.= misc. flag assigned by field crew. Explain all flags In Comment sections.
Rev. 06/02/97 (st_saco.97)
SAMPLE COLLECTION FORM - STREAMS -1
Figure 8-2. Sample Collection Form (pagel) showing data recorded for periphyton samples.
123
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EMAP-SW-Streams Field Operations Manual. Section 8 (Periphyton), Rev. 4, September 1998 Page 6 of 14
PERIPHYTON
APA d5TOMASSl> CHLA ID
SITE ID: MAIA_9_J3L-JL 5-
DATE: 7 / if 198
HABITAT: POOL <^RIFFLE/RUN>
SUBSAMPLE VOLUME: £/" mL
COMPOSITE VOLUME:
PERIPHYTON
<20S> BIOMASS CHLA
SITEID:MAIA ? 7 -
DATE: 7 l/fj98
HABITAT: POOL ^RIFFI
SUBSAMPLE VOLUME: S~O
COMPOSITE VOLUME:
ID
F/RLMsL?
229004
229004
mL
mL
PERIPHYTON
APA BIOMASS CHLA Q
SITE ID: MAIA J?. _2_ - -L JL "
DATE:
HABITAT:
POOL ^RIFFLE/RUN)
SUBSAMPLE VOLUME: JTQ mL
COMPOSITE VOLUME: &.OC) _ mL
PERIPHYTC
APA BIOMASS <
SITE ID: MAIAJ?_..3_-.
DATE:
HABITAT: POOL
SUBSAMPLE VOLUME: 2f
COMPOSITE VOLUME:
_mL
mL
229004
229004
Figure 8-3. Completed set of periphyton sample labels.
8.2.2 Chlorophyll Sample
Prepare chlorophyll samples by filtering a 25-mL aliquot of each composite index
sample through a glass fiber filter (0.4 to 0.6 urn nominal pore size). The procedure for
preparing chlorophyll samples is presented in Table 8-3. Chlorophyll can degrade rapidly
when exposed to bright light. If possible, prepare the samples in subdued light (or shade),
filtering as quickly as possible after collection to minimize degradation. The filtration appar-
atus is illustrated in Figure 8-4. Rinse the filtration chamber with deionized water each day
before use at the base site and then seal in a plastic bag until use at the stream (see Sec-
tion 3). 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 no exceed 7 pounds per square inch (psi) to
avoid rupturing cells. If the vacuum pressure exceeds 7 psi, prepare a new sample. If the
124
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton), Rev. 4, September 1998 Page 7 of 14
TABLE 8-2. PREPARATION OF ID/ENUMERATION SAMPLES FOR PERIPHYTON
1. Thoroughly mix the bottle containing the "EROSIONAL" composite index sample.
2. Prepare a barcoded sample label. Circle the sample type ("ID") and habitat type ("RIFFLE/RUN"
for EROSIONAL. "POOL" for DEPOSITIONAL) on the label. Record the volume of the sub-
ample (typically 50 mL) and the volume of the composite index sample on the label. Attach the
completed label to a 50-mL centrifuge tube; avoid covering the volume graduations and mark-
ings. Cover the label completely with a clear tape strip.
3. Place and "X" in the appropriate "HABITAT TYPE" box (riffle or pool) in the first "COMPOSITE
PERIPHYTON SAMPLE" section of the Sample Collection Form. Record the sample ID number
(barcode) of the label and the total volume of the composite index sample on the form.
4. Rinse a 60-mL syringe with deionized water.
5. Withdraw 50 mL of the composite index sample into the syringe. Place the contents of the
syringe sample into the labeled 50-mL centrifuge tube.
6. Wearing gloves and safety glasses, use a syringe or bulb pipette to add 2 mL of 10% formalin
solution to the tube. Cap the tube tightly and seal with plastic electrical tape. Shake gently to
distribute preservative.
7. Record the volume of the sample in the centrifuge tube (excluding the volume of preservative)
in the "ASSEMBLAGE ID SUBSAMPLE VOL." field of the Sample Collection Form.
8. Repeat Steps 1 through 7 above for the DEPOSITIONAL composite index sample. Record
information in the second "COMPOSITE PERIPHYTON SAMPLE" section of the Sample Collection
Form.
125
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton), Rev. 4, September 1998 Page 8 of 14
TABLE 8-3. PROCEDURE FOR PREPARING CHLOROPHYLL SAMPLES FOR PERIPHYTON
Mix the "EROSIONAL" composite index sample bottle thoroughly.
2. Using clean forceps, place a glass fiber filter on the filter holder. Use a small amount of deion-
ized water from a wash bottle to help settle the filter properly. Attach the filter funnel to the
filter holder and filter chamber, then attach the hand vacuum pump to the chamber.
4. Rinse the sides of the filter funnel and the filter with a small volume of deionized water.
5. Rinse a 25-mL or 50-mL graduated cylinder three times with small volumes of deionized water.
Measure 25 mL (±1 ml_) of sample into the graduated cylinder.
NOTE: For composite samples containing fine sediment, (e.g., the "DEPOSITIONAL"
sample), allow grit to settle before pouring the sample into the graduated cylinder.
6. Pour the 25-mL aliquot into the filter funnel, replace the cap, and pump the sample through the
filter using the hand pump. NOTE: Vacuum pressure from the pump should not exceed 7
psi to avoid rupture of fragile algal cells.
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 sam-
ple, measured to ±1 mL. Be sure to record the actual volume sampled on the sample
label and the Sample Collection Form.
7. Remove both plugs from the filtration chamber and pour out the filtered water in the chamber.
Remove the filter funnel from the filter holder. Remove the filter from the holder with clean
forceps. Avoid touching the colored portion of the filter. Fold the filter in half, with the colored
side folded in on itself. Wrap the folded filter in a small piece of aluminum foil.
9. Complete a periphyton sample label for chlorophyll, including the type of composite index
sample and the volume filtered, and attach it to the foil. Cover the label completely with a strip
of clear tape. Place the foil packet into a self-sealing plastic bag.
10. Place and "X" in the appropriate "HABITAT TYPE" box (riffle or pool) in the first "COMPOSITE
PERIPHYTON SAMPLE section of the Sample Collection Form. Record the sample ID number
(barcode) of the label and the total volume of the composite index sample on the form. Record
the volume filtered in the "CHLOROPHYLL" field" on the Sample Collection Form. Double check
that the volume recorded on the collection form matches the total volume recorded on the
sample label.
11. Place the plastic bag containing the filter into a portable freezer, a cooler containing dry ice, or
between two sealed plastic bags of ice in a cooler.
12. Rinse the filter funnel, filter holder, filter chamber, and graduated cylinder thoroughly with
deionized water.
13. Repeat Steps 1 through 12 for the "DEPOSITIONAL" composite index sample. Record
information in the second "COMPOSITE PERIPHYTON SAMPLE" section on the Sample Collection
Form. __^^_
126
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EMAP-SW-Streams Field Operations Manual. Section 8 (Periphyton), Rev. 4. September 1998 Page 9 of 14
HAND
VACUUM PUMP
FILTER
HOLDER yv
FILTER
CHAMBER
CLEAR
PLASTIC
TUBING
Figure 8-4. Filtration apparatus for preparing chlorophyll and biomass subsamples for peri-
phyton. Modified from Chaloud et al. (1989).
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. Rinse the
filtration unit and the graduated cylinder thoroughly with deionized water between the two
composite index samples.
After filtering each sample, wrap the filter in aluminum foil. Complete a sample label
(Figure 8-3) and check it to ensure that all written information is complete and legible. Affix
the label to the foil packet and cover it completely with a strip of clear tape. Record the
barcode assigned to the sample on the Sample Collection Form (Figure 8-2). Make sure
the volume recorded on each sample label matches the corresponding volume recorded on
the Sample Collection Form. Record a flag and provide comments on the Sample Collec-
127
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton), Rev. 4, September 1998 Page 10 of 14
tion Form if there are any problems in collecting the sample or if conditions occur that may
affect sample integrity. Store each foil packet in a self-sealing plastic bag. Store the sam-
ple frozen until shipment to the laboratory (Section 3).
8.2.3 Biomass Sample
Prepare the biomass samples from 25-mL aliquots of each composite index sample.
The filters for the biomass samples (the same type as is used for chlorophyll) may be
provided in a sealed, numbered container. These filters have been prepared by combusting
(30 min at 525 °C), desiccating, re-hydrating, drying (60 °C for 24 hours), then weighed to
the nearest 0.01 mg. Prepare each sample according to the procedure presented in Table
8-4. Take extra care in handling the filters, as they may be very fragile as a result of their
preparation. As with the chlorophyll sample, it is important to measure the volume to be
filtered accurately (±1 mL). Rinse the filter chamber components (Figure 8-4) and the
graduated cylinder thoroughly between the two composite index samples with deionized
water.
After filtering each sample, do not fold the filter (as was done for the chlorophyll
sample). Place the unfolded filter back into its numbered container. Complete a sample
label as shown in Figure 8-3. Check each sample label to ensure that all written information
is complete and legible. Affix the label to the filter container and cover it completely with
clear tape. Record the bar code assigned to the sample, the container number, and the
volume filtered on the Sample Collection Form as shown in Figure 8-2. Make sure the
information recorded on each sample label and filters container matches the corresponding
values recorded on the Sample Collection Form. Record a flag and provide comments on
the Sample Collection Form if there are any problems in collecting the sample or if condi-
tions occur that may affect sample integrity. Store each labeled filter container frozen until
shipment to the laboratory (Section 3).
8.2.4 Acid/Alkaline Phosphatase Activity Sample
The Acid/Alkaline phosphatase activity (APA) samples are prepared from 50-mL
subsamples of each composite index sample. Table 8-5 presents the procedure for prepar-
ing APA samples. No field treatment (i.e., filtration, preservation) of the APA sample is
necessary. Complete a label for each sample as shown in Figure 8-3 and affix it to a 50-mL
centrifuge tube. Record the ID number (barcode), and the volume of the subsample on the
Sample Collection Form (Figure 8-2). Check to ensure that the information recorded on the
128
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton). Rev. 4, September 1998 Page 11 of 14
TABLE 8-4. PROCEDURE FOR PREPARING BIOMASS SAMPLES FOR PERIPHYTON
1. Mix the "EROSIONAL" composite index sample bottle thoroughly.
2. Using clean forceps, remove a pre-leached, pre-weighed glass-fiber filter from its numbered
container and place it on the filter holder. Usexa 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 cham-
ber, then attach the hand vacuum pump to the chamber.
3. Rinse the filter chamber and filter with a small volume of deionized water.
4. Rinse a 25-mL or 50-mL graduated cylinder three times with small volumes of deionized water.
Measure 25 ml_ (±1 ml_) of composite index sample into the graduated cylinder.
NOTE: For composite samples containing fine sediment, (e.g., the "DEPOSITIONAL"
sample), allow grit to settle before pouring the sample into the graduated cylinder.
5. Pour the 25-mL aliquot into filter funnel, replace the cap, and pump the sample through the filter
using the hand pump. NOTE: Filtration pressure should not exceed 7 psi to avoid rupture
of fragile algal cells.
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 sam-
ple, measured to +1 mL. Be sure to record the actual volume filtered on the sample label
and the Sample Collection Form.
6. Remove both plugs from the filtration chamber and pour out the filtered water in the chamber.
Remove the filter funnel from the filter holder. Remove the filter from the holder with clean
forceps. Avoid touching the colored portion of the filter.
8. Place the unfolded filter back into its numbered container. Complete a periphyton sample
label for biomass, including the type of index sample, the container number, and the volume
filtered. Affix the label to the filter container and cover the label completely with a strip of clear
tape.
9. Place and "X" in the appropriate "HABITAT TYPE" box (riffle or pool) in the first "COMPOSITE
PERIPHYTON SAMPLE" section of the Sample Collection Form. Record the sample ID number
(barcode) of the label and the total volume of the composite index sample on the form. Record
the number from the filter container ("FILTER No.") and the volume filtered in the "BIOMASS"
portion on the Sample Collection Form. Double check that the volume recorded on the collec-
tion form matches the total volume recorded on the sample label.
10. Place the labeled filter container into a portable freezer, a cooler containing dry ice, or between
two sealed plastic bags of ice in a cooler.
11. Rinse the filter funnel, filter holder, filter chamber, and graduated cylinder thoroughly with
deionized water.
12. Repeat Steps 1 through 11 for the "DEPOSITIONAL" composite index sample. Record infor-
mation in the second "COMPOSITE PERIPHYTON SAMPLE" section on the Sample Collection Form.
129
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EMAP-SW-Streams Field Operations Manual. Section 8 (Periphyton), Rev. 4, September 1998 Page 12 of 14
TABLE 8-5. PROCEDURE FOR PREPARING ACID/ALKALINE PHOSPHATASE ACTIVITY
SAMPLES FOR PERIPHYTON
1. Thoroughly mix the bottle containing the "EROSIONAL" composite index sample.
2. Prepare a barcoded sample label. Circle the sample type ("ID") and habitat type
("RIFFLE/RUN" for the "EROSIONAL" sample; "POOL" for the "DEPOSITIONAL" sample) on
the label. Record the volume of the sample (typically 50 mL) and the volume of the compos-
ite index sample on the label. Attach the completed label to a 50-mL centrifuge tube; avoid
covering the volume graduations and markings. Cover the label completely with a clear
tape strip.
3. Rinse a 60-mL syringe with deionized water.
4. Withdraw 50 mL of the composite index sample into the syringe. Place the contents of the
syringe sample into the labeled 50-mL centrifuge tube. Cap the tube tightly and seal with
plastic tape.
5. Place and "X" in the appropriate "HABITAT TYPE" box (riffle or pool) in the first "COMPOSITE
PERIPHYTON SAMPLE" section of the Sample Collection Form. Record the sample ID number
(barcode) of the label and the total volume of the composite index sample on the form.
7. Record the volume of the sample in the centrifuge tube in the "APA SAMPLE" field of the
Sample Collection Form.
8. Repeat Steps 1 through 7 above for the "DEPOSITIONAL" composite index sample. Record
information in the second "COMPOSITE PERIPHYTON SAMPLE" section of the Sample Collection
Form.
130
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton), Rev. 4, September 1998 Page 13 of 14
Sample Collection Form matches the corresponding information recorded.on the sample
label. Store APA samples frozen until shipment to the laboratory (Section 3).
8.3 EQUIPMENT AND SUPPLIES
Figure 8-5 is a checklist of equipment and supplies required to conduct periphyton
sample collection and processing activities. This checklist is similar to the checklist pre-
sented in Appendix A, which is used at the base location (Section 3) to ensure that all of the
required equipment is brought to the stream. Use this checklist to ensure that equipment
and supplies are organized and available at the stream site in order to conduct the activities
efficiently.
8.4 LITERATURE CITED
Chaloud, D.J., J.M. Nicholson, B.P. Baldigo, C.A. Hagley, and D.W. Sutton. 1989. Hand-
book of Methods for Acid Deposition Studies: Field Methods for Surface Water Chem-
istry. EPA 600/4-89-020. U.S. Environmental Protection Agency, Washington, D.C.
131
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EMAP-SW-Streams Field Operations Manual, Section 8 (Periphyton), Rev. 4, September 1998 Page 14 of 14
EQUIPMENT AND SUPPLIES FOR PERIPHYTON
1
1
1
1
1
2
1
4
1 box
1 pair
1
1
1
2
2
2
4mL
1
1 pair
1 pair
2 sets
1
1 pkg.
1
1 copy
1 set
Large funnel (15-20 cm diameter)
12-cm2 area delimiter (3.8 cm diameter pipe, 3 cm tall)
Stiff-bristle toothbrush with handle bent at 90° angle
1-L wash bottle for stream water
1-L wash bottle containing deionized water
500-mL plastic bottles for composite index samples, labeled "EROSIONAL"
and "DEPOSITIONAL"
60 mL plastic syringe with 3/8" hole bored into the end
50-mL screw-top centrifuge tubes (or similar sample vials)
Glass-fiber filters for chlorophyll samples
Forceps for filter handling.
25-mL or 50-mL graduated cylinder
Filtration unit, including filter funnel, cap, filter holder, and receiving chamber
Hand-operated vacuum pump and clear plastic tubing
Pre-leached, pre-ashed, weighed glass-fiber filters in numbered containers for
biomass sample
Aluminum foil squares (3" x 6")
Self-sealing plastic bags for chlorophyll samples
10% formalin solution for ID/Enumeration samples
Small syringe or bulb pipette for dispensing formalin
Chemical-resistant gloves for handling formalin
Safety glasses for use when handling formalin
Sample labels (4 per set) with the same barcode ID number
Sample Collection Form for stream
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
Portable freezer, cooler with dry ice, or cooler with bags of ice to store frozen
samples
Field operations and method manual
Laminated sheets of procedure tables and/or quick reference guides for peri-
phvton
Figure 8-5. Checklist of equipment and supplies for periphyton.
132
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SECTION 9
SEDIMENT COMMUNITY METABOLISM
by
Brian H. Hill1
This section describes procedures to collect a composite sediment sample from the
sampling reach. The "biomorphs" (refer to Figure 2-1) collect sediment samples from each
transect at the same time as periphyton samples (Section 8) and benthic macroinvertebrate
samples (Section 11). At each stream, a composite "index" sample of sediment is pre-
pared. A portion of this composite sample is used in the determination of sediment commu-
nity metabolism. The remaining composite sample is prepared for use in toxicity testing, if
necessary (see Section 1.3.8 and Section 10).
The method outlined here for determining sediment community metabolism is de-
signed for headwater to mid-order streams, though it may be adapted for larger rivers or
lakes. The method measures changes in dissolved oxygen (DO) concentrations of the
overlying water within microcosms containing small amounts (ca. 10 mL) of sediments as a
means of assessing benthic microbial community activity. Sediments are collected from
depositional habitats along a study reach defined by 40 times the channel width. Following
incubation, the DO is remeasured and the sediments are saved for ash-free dry mass
(AFDM) analysis. Respiration rate, estimated as the change in DO concentration per hour
within each microcosm, is adjusted for AFDM, yielding a measure of community respiration
per gram of AFDM. Organic carbon turnover time can be calculated from the empirical
relationship between the organic carbon content of the sediment (estimated as 0.5 x AFDM)
and oxygen consumption.
9.1 SAMPLE COLLECTION
Table 9-1 describes the procedure for collecting the composite sediment sample.
Collect sediment from depositional areas (e.g., pools, eddies, and backwaters) located at or
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin Luther King
Dr., Cincinnati, OH 45268.
133
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EMAP-SW-Streams Field Operations Manual. Section 9 (Sediment Community Metabolism ). Rev. 4, Sept. 1998 Page 2 of 8
TABLE 9-1. SEDIMENT COLLECTION PROCEDURE
1. At cross-section transect "B", locate a depositional habitat (a pool, eddy, or backwater).
If soft sediments are scarce, collect them wherever you can within the reach
2, Use a plastic scoop to collect a sample of surficial sediment (top 2 cm). Remove any visible
organisms from the sediment. Place the sample in a plastic jar with volume graduations
labeled "SEDIMENT SAMPLE".
Approximately 3 L of sediment (~ 400 mL of sediment per transect) is required for both
sediment metabolism and sediment toxicity. If a sediment toxicity sample is not required,
250 mL of sediment (~ 30 mL per transect) is sufficient.
3. Repeat Steps 1 through 2 for Transects "C" through "J".
134
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EMAP-SW-Streams Field Operations Manual, Section 9 (Sediment Community Metabolism ), Rev. 4, Sept. 1998 Page 3 of 8
near each of the nine interior cross-section transects ("B" through "J") within the sampling
reach. If soft sediments are scarce, collect them from wherever you can within the sam-
pling reach. At each sampling point, use a small plastic scoop to collect the top 2 cm (~ 1
inch) of soft surface sediment. Combine sediments from different sampling points into a
single jar or self-sealing plastic bag to prepare a single composite index sample for the
stream reach. A composite sample volume of 250-mL is sufficient to prepare sediment
metabolism samples. An additional 1 to 2 L of sediment is required for a sediment toxicity
sample.
9.2 DETERMINING SEDIMENT RESPIRATION
The procedure to measure sediment respiration in presented in Table 9-2. A dis-
solved oxygen meter, equipped with a biological oxygen demand (BOD) probe and stirrer, is
used for the determination of respiration rates. This may or may not be the same meter
used to determine in situ dissolved oxygen concentration (Section 5). If a separate meter is
used to measure sediment respiration, check the probe membrane and the meter's batter-
ies and electronics according to the instrument's operating manual (see Sections 3 and 5,
also). Calibrate the meter as directed in the instrument's operating manual.
A small cooler filled with stream water is used as an incubation chamber. The initial
dissolved oxygen concentration and temperature of the water in the cooler are measured
and recorded on the Field Measurement Form as shown in Figure 9-1. This concentration
is assumed to be the initial concentration of all subsamples. Five subsamples (10-mL ±1
mL) are prepared from the composite sediment sample. A set of completed sample labels
for these subsamples is shown in Figure 9-2. A 10-mL subsample of water from the incuba-
tion cooler is used as a control for changes in ambient conditions during the incubation.
The subsamples are incubated in the cooler for 2 hours. After the incubation, the final DO
concentration of each tube is determined and recorded on the Field Measurement Form
(Figure 9-1). The sediment in each tube is retained and stored frozen until it can be
shipped to the laboratory (Section 3) to determine the AFDM.
9.3 EQUIPMENT AND SUPPLIES
Figure 9-3 is a checklist of equipment and supplies required to conduct sediment
sampling and to determine sediment community respiration. This checklist is similar to the
checklist presented in Appendix A, which is used at the base location (Section 3) to ensure
135
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EMAP-SW-Streams Field Operations Manual, Section 9 (Sediment Community Metabolism ), Rev. 4, Sept. 1998 Page 4 of 8
TABLE 9-2. PROCEDURE TO MEASURE SEDIMENT RESPIRATION
1. Inspect the probe of the dissolved oxygen meter for outward signs of fouling and for an intact
membrane. Do not touch the electrodes inside the probe with any object. Always keep the
probe moist by keeping it inside its calibration chamber. Check the batteries and electronic
functions of the meter and stirrer unit as described in the meter's operating manual.
2. Calibrate the oxygen probe in water-saturated air as described in the operating manual. Allow
at least 15 minutes for the probe to equilibrate before attempting to calibrate.
NOTE: Try to perform the calibration as close to stream temperature as possible (not air
temperature) by using stream water to fill the calibration chamber prior to equilibration.
NOTE: For doing the elevation correction, the elevation of the sample site is provided on
the site information sheet in the dossier for the site. Alternatively, obtain the elevation
from a topographic map.
3. Prepare a set of five sediment metabolism sample labels. Note that each label will have a
different sample ID number (barcode). Attach each completed label to a 50-mL screw-cap
centrifuge tube.
NOTE: Avoid covering volume gradations on the tube with the label. Cover each label
with a strip of clear tape.
4. Fill a small insulated cooler % full with streamwater. Measure the dissolved oxygen and
temperature of the water in the cooler. Record the values in the "INITIAL O2" and "INITIAL
INCUBATION TEMP." fields in the metabolism section of the Field Measurement Form.
5. Thoroughly mix the composite sediment sample. Use a small plastic spoon to transfer
10 mL of sediment from the composite sample container to each of the five labeled
tubes.
6. Fill each tube to the top (no head space) with stream water from the cooler and seal the
tube. Fill a centrifuge tube labeled "BLANK" with stream water from the cooler and seal.
This tube serves as a control for changes in ambient conditions during the incubation
period.
7. Place the six tubes in a 1-L plastic beaker and place the beaker inside the cooler. Record the
start time in the "INCUBATION TIME" area of the Field Measurement Form. Close the cooler and
incubate the sediment samples for 2 hours.
~~~ (continued)
136
-------
EMAP-SW-Streams Field Operations Manual, Section 9 (Sediment Community Metabolism ), Rev. 4, Sept. 1998 Page 5 of 8
TABLE 9-2 (Continued)
8. If necessary, re-calibrate the oxygen probe (i.e., the meter was turned off or you have moved
to a different elevation during the incubation) before the end of the incubation period.
9. At the end of the incubation period, record the end time in the "INCUBATION TIME" area of the
Field Measurement Form. Measure the DO in each tube, including the blank. Record the
sample ID number of each tube and its measured DO concentration on the Field Measurement
Form.
10. Decant the overlying water from each labeled tube, retaining the sediment. Tightly seal each
tube and place in a portable freezer, a container with dry ice, or in a cooler with bags of ice as
soon as possible. Keep the samples frozen until they can be shipped. Discard the water from
the "BLANK" tube.
11. If the remaining composite sediment sample will not be used for a sediment toxicity sample,
discard the sample and rinse the composite sample container thoroughly with stream water
and/or deionized water.
137
-------
EMAP-SW-Streams Field Operations Manual. Section 9 (Sediment Community Metabolism ), Rev. 4, Sept. 1998 Page 6 of 8
Reviewed by (initial):
FIELD MEASUREMENT FORM - STREAMS/RIVERS
SITE NAME: ft} ILL CKfffK
SITE ID: MAIA97- 7 ? *
DATE: 1 1 AS"/ 97 VISIT: 01 D2
TEAM ID (X): B1 D2 D3 Q4 D5 D6 D7 D8
WEATHER CONDITIONS 00
CLOUD COVER
PRECIPITATION
PREVIOUS PRECIPITATION (24 H)
AIR TEMPERATURE XX
H<5%
HNONE
DNONE
a. ? .c
IN SITU MEASUREMENTS
QCCS COND MS/CM
STREAM/RIVER COND X/S/OM
STREAM/RIVER DO MG/L
STOEAM/RIVER TEMP °C
.2. A.
5" o c?
V. £
2 c? . o
D5-25% D 25-50% d 50-75% D>75%
O LIGHT D MODERATE D HEAVY
H LIGHT D MODERATE D HEAVY
| STATION ID: Jg_ \ Assume X-alte unless marked
FLAG COMMENTS
STREAM/RIVER METABOLISM DETERMINATION
,.._,.. n INITIAL
INITIAL Oi iMi-imA-nnni
(MO/L) TEMP.(°C)
"/ .*L 2. c?. e?
SAMPLE ID FINAL O,
(BARCODE) (MG/L)
a.JLJ.JLJLJ- _A.S.
1AJLJ1.1A £-.£
±.fL-t_l-2.-2. L.S-
±3_J_JLJL1L -L.3-
±3.±1-JL£- L.-L
AJ=.JL^JL —±L.£.
INCUBATION TIME
START FINISH
LJL.3-f i-.V-i
FLAG
DURATION OF
— INCUBATION FLAG COMMENTS
(HH:HM)
7 _3_-.e_£L
COMMENTS
OXYGEN METER CALIBRATION INFORMATION
MEMORAHE CHECK fxl
ELECTRONIC ZERO |}?| RED LINE: ^tj
CAUBnATlOHCKAIWERTEUPEBATURE: St. 7. *7 *C SATURATED 0, 9 TEMP.: *7. 93 MGfl.
STATIONELEVATIONiFROMTOPO.MAPORALTIMETER): / O
Tt<» aHartton v«ta h obulf»d by miMplylng Ita utmkd DO co(K«itrttoo Om
•Wittn OOCTKBOO f«dor (obUi»d from th. tabm on U» luck o< ft« YSI imtar).
nwtar rMdlng to tt» canntkn v ihw.
"fO FT ELEVATION CORRECTION FACTOR: x <9.9£
»«n CALIBRATION VALUE: tj, f 2. MG/L
ld(urttt>«
COMMENTS:
Flag Codec K » no measurement or observation made; U= suspect measurement or observation; Q = unacceptable QC check associated wlm
moMurement F1, F2, etc.» miscellaneous (lags assigned by each Held crew. Explain all flags In comments section.
Rev. OS/DZ/97 (strvfldm.97)
FIELD MEASUREMENT FORM - STREAMS/RIVERS -1
Figure 9-1. Field Measurement Form (page 1), showing data for sediment metabolism sam-
ples.
138
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EMAP-SW-Streams Field Operations Manual, Section 9 (Sediment Community Metabolism ), Rev. 4, Sept. 1998 Page 7 of 8
SEDIMENT METABOLISM
. SITE ID:MAIA _2. 3- - -3_ -?_ J_
DATE: 7 //r/98
SAMPLE TYPE: Rl fc) R3 R4 R5
SEDIMENT METABOLISM
SITE ID:MAIA _£ i - _
DATE: 7 //r/98
SAMPLE TYPE: (Ry R2 R3 R4 R5
229007
229006
SEDIMENT METABOLISM
SITE ID:MAIA ^L 1_ -3_S_3_
DATE: 7 //J"/98
SAMPLE TYPE: R1 R2(^>R4 R5
SEDIMENT METABOLISM
SITE ID:MA!A _?_ _2_ -
229008
DATE: 7 /
SAMPLE TYPE: R1 R2 R3(R4)R5
229009
SEDIMENT METABOLISM
SITE ID:MAIA
DATE: 7 MTV 98
SAMPLE TYPE: R1 R2 R3
229010
Figure 9-2. Completed sample labels for sediment metabolism.
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 stream site in order to conduct
the activities efficiently.
139
-------
EMAP-SW-Streams Field Operations Manual, Section 9 (Sediment Community Metabolism •), Rev. 4, Sept. 1998 Page 8 of 8
EQUIPMENT AND SUPPLIES FOR SEDIMENT METABOLISM
QTY
1
1
1
1 set
1
5
1
1
1
5
1
1 pkg
1
1 copy
1 set
ITEM
Small scoop sampler for sediments
Wide-mouthed plastic jar labeled "COMPOSITE SEDIMENT SAMPLE". If
sediment is only being collected for metabolism samples, a 250-mL jar is
sufficient. If metabolism and toxicity samples are being prepared, use a 1-
gallon jar
YSI Model 58 Dissolved Oxygen meter with Model 5730 Stirring BOD probe
Spare batteries for DO meter
Small plastic spoon or spatula to transfer sediment from the composite sam-
ple container to respiration tubes
50-mL, screw-top, centrifuge tubes
50-mL screw-cap centrifuge tube labeled "BLANK"
Small cooler used as incubation chamber
1,000-mL plastic beaker to holding centrifuge tubes during incubation
Sediment metabolism sample labels (each with different ID number)
Field Measurement Form
Soft (#2) lead pencils to fill in field data forms
Fine tip indelible markers for preparing labels
Clear tape strips for covering labels
Portable freezer, or cooler with bags of ice or dry ice to store sediment metab-
olism samples.
Field operations and methods manual
Laminated sheets'of procedure tables and/or quick reference guides for sedi-
ment community metabolism
Figure 9-3. Checklist of equipment and supplies for sediment metabolism.
140
-------
SECTION 10
SEDIMENT TOXICITY
by
James M. Lazorchak1 and Mark E. Smith2
This section describes procedures to prepare a sediment sample for shipment to a
laboratory for use in toxicity testing (see Section 1.3.8). The "biomorphs" (refer to Figure 2-
1) collect sediment samples from each transect at the same time as periphyton samples
(Section 8) and benthic macroinvertebrate samples (Section 11). At each stream, a com-
posite "index" sample of sediment is prepared. A portion of this composite sample is used
in the determination of sediment community metabolism (Section 9).
10.1 SAMPLE COLLECTION AND PREPARATION
The composite sediment sample remaining after the sediment respiration sub-
samples have been prepared is used to prepare a sediment toxicity sample. The procedure
to prepare the sediment toxicity sample is presented in Table 10-1. A completed sample
label for the sediment toxicity sample is shown in Figure 10-1. Record the sample ID num-
ber on the Sample Collection Form as shown in Figure 10-2. Use a heavy-duty self-sealing
plastic bag as a sample container. Double-bag the sample and place it a suitably sized
hard plastic container with a snap-on lid for transport and storage. Keep the sample chilled
(but not frozen) until it can be shipped to the laboratory (Section 3).
10.2 EQUIPMENT AND SUPPLIES
Figure 10-3 is a checklist of equipment and supplies required to prepare the sed-
iment toxicity sample. This checklist is similar to the checklist presented in Appendix A,
which is used at the base location (Section 3) to ensure that all of the required equipment is
brought to the
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin Luther King
Dr., Cincinnati, OH 45268.
2
SoBran Environmental, Inc., c/o U.S. EPA, 26 W. Martin Luther King Dr., Cincinnati, OH 45268.
141
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EMAP-SW-Streams Field Operations Manual, Section 10 (Sediment Toxicity). Rev. 4, September 1998 Page 2 of 5
TABLE 10-1. PROCEDURE FOR PREPARING SEDIMENT TOXICITY SAMPLES
1. Complete a sediment toxicity sample label with the stream ID and the date of collection. If
sediment for the composite sample was collected at several cross-section transects, write
"ALL" in the "STATION" field.
2. Record the sample ID number (barcode) printed on the label in the "SEDIMENT TOXICITY SAM-
PLES" section of the Sample Collection Form (page 2).
3. Attach the completed label to a 2-gallon polyethylene (4-mil) bag. Cover the label with a strip
of clear (ape.
4. Mix sediment well with a stainless steel or plastic mixing spoon, or gloved hand. Transfer at
least 1 L of sediment from the composite sediment index sample container to the labeled
plastic bag. Close bag, squeeze air out and tie a knot in the remaining portion of the bag to
seal. Seal the bag.
5. Place the labeled bag with the sample inside a second 2-gallon polyethylene bag and tie off
the top to seal. Place the sample into a hard plastic container with a snap-on lid (if available),
to further protect the sample.
6. Place the sample inside a cooler containing bags of ice that is used only for sediment sam-
ples in them. Store the sediment toxicity sample chilled, but not frozen, until it can be shipped
to the laboratory.
142
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EMAP-SW-Streams Field Operations Manual. Section 10 (Sediment Toxicity), Rev. 4, September 1998 Page 3 of 5
SEDIMENT TOXICITY
SITE ID: MAI A 3- _2. - ? ? 9
DATE: 7 / AT798
STATION:
229011
Figure 10-1. Completed sample label for sediment toxicity.
stream. Use this checklist to ensure that equipment and supplies are organized and avail-
able at the stream site in order to conduct the activities efficiently.
143
-------
EMAP-SW-Streams Field Operations Manual, Section 10 (Sediment Toxicity), Rev. 4. September 1998 Page 4 of 5
Reviewed by (initial):
SAMPLE COLLECTION FORM - STREAMS (continued)
SITE NAME: A)/AL
DATE: 7
VISIT: H1 D2
SITEID: MAIA97-_2 _3__?
TEAM ID (X): B1 D2 D3 D4 D5 D6 D7 D8
CHEMISTRY AND MICROBIAL WATER SAMPLE (Chem: 4-L Cubitainer and 2 Syringes, Micro: Glass Bottle)
SAMPLE ID (BARCODE)
TRANSECT
FLAG
COMMENTS
CHEMISTRY
o i SL
MICROBIAL
SEDIMENT TOXICITY SAMPLES
SAMPLE ID (BARCODE) FLAG
COMMENTS
FISH TISSUE SAMPLES - PRIMARY SAMPLE (mln. 50g total wgt)
SAMPLE ID (BARCODE) -
LINE
SPECIES CODE
COMMON NAME
NUMBER OF
INDIVIDUALS
FLAG
P1
IS COMPOSITE SAMPLE COMPOSED OF INDIVIDUALS COLLECTED FROM THROUGHOUT REACH? (X) -
BYES D NO
IF No, EXPLAIN:
FISH TISSUE SAMPLES - SECONDARY SAMPLE (where available; 5 Individuals)
SAMPLE ID (BARCODE) -
2. JL O JL
LINE
SPECIES CODE
COMMON NAME
TOTAL LENGTH (MM)
FLAG
S1
C A TO C o
svc.tr
S2
T O CO
S3
S4
55
er
IS COMPOSITE SAMPLE COMPOSED OF INDIVIDUALS COLLECTED FROM THROUGHOUT REACH? (X) -
BYES DNO
IF No, EXPLAIN:
LINE
COMMENT OR FLAG EXPLANATION FOR FISH TISSUE
Ft s It /V^ltw^../, i^.
Flag codes: Id Sampla not collected; U= Suspect sample; F1, F2,eto.= misc. flag assigned by field crew. Explain all flags In Comments sections.
(8t_s«co.97)
SAMPLE COLLECTION FORM - STREAMS - 2
Figure 10-2. Sample Collection Form (page 2), showing information recorded for a sediment
toxicity sample.
144
-------
EMAP-SW-Streams Field Operations Manual, Section 10 (Sediment Toxicity), Rev. 4, September 1998 Page 5 of 5
EQUIPMENT AND SUPPLIES FOR SEDIMENT TOXICITY
OTY.
1
1
1
1
2
1 pkg
1
1
1 copy
1 set
ITEM
Small scoop sampler for sediments
Wide-mouthed plastic jar labeled "COMPOSITE SEDIMENT SAMPLE". If
sediment is only being collected for metabolism samples, use a 250-mL jar is
sufficient. If metabolism and toxicity samples are being prepared, use a 1-
gallon jar
Sediment toxicity sample label
Sample Collection Form
Soft (#2) lead pencils to fill in field data forms
Fine tip indelible markers for preparing labels
1 -gallon heavy-duty self-sealing plastic bags for the sediment toxicity sample
Clear tape strips for covering labels
Plastic container with snap-on lid to hold sediment toxicity sample
Cooler with bags of ice to store the sediment toxicity sample
Field operations and methods manual
Laminated sheets with procedure tables and/or quick reference guides for
sediment toxicity
Figure 10-3. Checklist of equipment and supplies for sediment toxicity.
145
-------
-------
SECTION 11
BENTHIC MACROINVERTEBRATES
by
Donald J. Klemm1, James M. Lazorchak1, and Philip A. Lewis1-2
Benthic invertebrates inhabit the sediment or live on the bottom substrates of
streams. Benthic macroinvertebrate assemblages in streams reflect overall biological
integrity of the benthic community. Monitoring these assemblages is useful in assessing the
status of the water body and detecting trend in ecological condition. Benthic communities
respond to a wide array of stressors in different ways so that it is often possible to deter-
mine the type of stress that has affected a macroinvertebrate community (e.g., Klemm et
al., 1990). Because many macroinvertebrates have relatively long life cycles of a year or
more and are relatively immobile, macroinvertebrate community structure is a function of
present or past conditions.
The EMAP-SW benthic macroinvertebrate protocol is intended to evaluate the
biological integrity of wadeable streams in the United States for the purpose of detecting
stresses on community structure and assessing the relative severity of these stresses. It is
based on the "Rapid Bioassessment Protocol III - Benthic Macroinvertebrates" published, by
the U.S. Environmental Protection Agency (Plafkin et al., 1989) and adopted for use by
many states. The two man kick net procedure of the Rapid Bioassessment Protocol (RBP)
is replaced in the EMAP-SW protocol with a kick net modified for use by one person (Figure
11-1), as is used by the U.S. Geological Survey for their National Water-Quality Assess-
ment Program (NAWQA; Cuffney et al., 1993). This protocol requires only one person and
is the preferred macroinvertebrate collecting method for streams with flowing water (a
second person is often used for water safety and to keep time and record information on the
field forms).
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin Luther King
Dr., Cincinnati, OH 45268.
2 Current address: 1037 Wylie Road, RR #2, Seaman, OH 45679.
147
-------
r
EMAP-SW-Streams Field Operations Manual. Section 11 (Benthic Macroinvertebrates). Rev. 4..Sept. 1998 Page 2 of 14
1.5 m long, 2-piece detachable handle
30cm
Sewed End
Canvas Bottom Panel
Figure 11-1. Modified kick net. (Not drawn to scale.)
The "biomorphs" (refer to Figure 2-1) collect kick net samples for benthic macro-
invertebrate at sampling points located on each cross-section transect. Kick net samples
are collected at the same time as periphyton samples (Section 8) and sediment samples
(Section 9). Kick net samples collected from flowing water habitats (e.g., riffles, runs) are
combined into a single composite sample for the stream reach. Kick net samples collected
from pool habitats are combined into a separate composite sample.
148
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EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates). Rev. 4. Sept. 1998 Page 3 of 14
11.1 SAMPLE COLLECTION
The index sample design for benthic macroinvertebrates is shown in Figure 11-2.
This design is used in the EMAP and R-EMAP stream studies in the mid-Atlantic region
(refer to Section 1 for project descriptions). A modified index sample design was developed
and implemented in some studies conducted in the western U.S. In the modified design, an
equal number of kick net samples are collected from available riffle and pool habitats lo-
cated within the sampling reach. This modified index sampling design is described in more
detail in Appendix E.
A kick net sample is collected from each of the nine interior cross-section transects
(Transects "B" through "J") at an assigned sampling point (Left, Center, or Right). These
points may have been assigned when the sampling reach was laid out (Figure 11-2; refer
also to Section 4; Table 4-3). If not, the sampling point at Transect "B" is assigned at
random using a die or other suitable means (e.g., digital watch). Once the first sampling
point is determined, points at successive transects are assigned in order (Left, Center,
Right). These are the same sampling points as those used for periphyton samples (Section
8). 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".
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. To collect a kick net sample from a sampling point classified as
"riffle/run" habitat, follow the procedure presented in Table 11-1. To collect a kick net
sample from a sampling point classified as a "pool/glide" habitat, follow the procedure
presented in Table 11-2. Record the habitat type and sampling point for each kick net
sample collected on the Sample Collection Form as shown in Figure 11-3. As you proceed
upstream from transect to transect, combine all kick net samples collected from "riffle/run"
habitats into a bucket or similar container labeled "RIFFLE". Combine kick net samples
collected from "pool/glide" habitats into a second bucket labeled "POOL". Fill in the check-
list shown in Figure 11-4 as individual activities are completed.
If it is impossible to sample at the sampling point with the modified kick net following
either procedure, spend about 60 seconds hand picking a sample from about 0.25 m2 of
149
-------
EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates), Rev. 4, Sept. 1998 Page 4 of 14
CROSS SECTION TRANSECTS (A to K)
TRANSECT SAMPLES (1 per transect)
Sampling point of esch transect (1/4,1/2, 3/4) selected at random
• Modified kick net (595 |jm mesh)
Combine all kick net
samples collected from
riffles and runs
Combine all kick net
samples collected from
pools
500-mL or 1-L aliquots
Preserve with 70% ethanol
Figure 11-2. Index sampling design for benthic macroinvertebrates.
150
-------
EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates), Rev. 4, Sept. 1998 Page 5 of 14
TABLE 11-1. PROCEDURE TO COLLECT KICK NET SAMPLES FROM
RIFFLE AND RUN HABITATS '
At each cross-section transect, beginning with Transect "B", locate the assigned sampling point
(Left, Center, or Right as you face downstream) as 25%, 50%, and 75% of the wetted width,
respectively.
If the sampling points were not assigned previously when laying out the sampling reach,
proceed to Transect "B". Roll a die to determine if it is a left (L), center (C), or right (R) sam-
pling point for collecting periphyton and benthic macroinvertebrate samples. A roll of 1 or 2
indicates L, 3 or 4 indicates C, and 5 or 6 indicates R (or use a digital wristwatch and glance at
the last digit (1-3=L, 4-6=C, 7-9=R). Mark L, C, or R on the transect flagging. Assign sampling
points at each successive transect in order as L, C, R after the first random selection.
Attach the 4-ft handle to the kick net. Make sure that the handle is on tight or the net may
become twisted in a strong current, causing the loss of part of the sample.
Determine if there is sufficient current in the area at the sampling point 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.
With the net opening facing upstream, position the net quickly and securely on the stream
bottom to eliminate gaps under the frame. Avoid large rocks that prevent the sampler from
seating properly on the stream bottom.
• NOTE: If there is too little water to collect the sample with the kick net, randomly pick up
10 rocks from the riffle and pick and wash the organisms off them into a bucket labeled
"RIFFLE" which is half-full of water.
Holding the net in position on the substrate, visually define a rectangular quadrat that is one
net width wide and two net widths long upstream of the net opening. The area within this
quadrat is -0.5 m2.
\
Check the quadrat for heavy organisms, such as mussels and snails. Remove these organ-
isms from the substrate by hand and place them into the net.
Hold the net securely in position while kicking the substrate within the quadrat vigorously for 20
seconds (use a stopwatch).
After 20 seconds, hold the net in place with your knees and pick up any loose rocks within the
quadrat. Use your hands to rub any clinging organisms off the rocks (especially those covered
with algae or other debris) in front of the net. Also, place any additional mussels and snails
found into the net. Remove the net from the water with a quick upstream motion to wash the
organisms to the bottom of the net.
(continued)
151
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EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates), Rev. 4, Sept. 1998 Page 6 of 14
TABLE 11-1. (Continued)
8. Invert the net into a bucket labeled "RIFFLE", which is about half full of water, to rinse organ-
isms out of the net. Inspect the net for clinging organisms. Use watchmakers' forceps to
remove any organisms from the net and place them in the bucket. 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 any organisms.
9. Place an "X" in the appropriate habitat type and sampling point boxes for the transect on the
Sample Collection Form.
10. Proceed upstream to the next transect and repeat Steps 1 through 9. Combine all kick net
samples from "riffle/run" habitats into the "RIFFLE" bucket.
152
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EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates), Rev. 4. Sept. 1998 Page 7 of 14
TABLE 11-2. PROCEDURE TO COLLECT KICK NET SAMPLES FROM
POOL AND GLIDE HABITATS
1. At each cross-section transect, beginning with Transect "B", locate the assigned sampling point
(Left, Center, or Right as you face downstream) as 25%, 50%, and 75% of the wetted width,
respectively.
If the sampling points were not assigned previously when laying out the sampling reach,
proceed to Transect "B". Roll a die to determine if it is a left (L), center (C), or right (R) sam-
pling point for collecting periphyton and benthic macroinvertebrate samples. A roll of 1 or 2
indicates L, 3 or 4 indicates C, and 5 or 6 indicates R (or use a digital wristwatch and glance at
the last digit (1-3=L, 4-6=C, 7-9=R). Mark L, C, or R on the transect flagging. Assign sampling
points at each successive transect in order as L, C, R after the first random selection.
2. Attach the 4-ft handle to the kick net. Make sure that the handle is on tight or the net may
become twisted during the collection process, causing the loss of part of the sample.
3. Determine if there is sufficient current in the area at the sampling point to fully extend the net. If
so, use the sampling procedure described for "Riffle/run" habitats. If not, classify the habitat as
"Pool/glide" and proceed to Step 4. NOTE: If the pool is too deep (much more than 1 m) to
sample safely at the designated spot, move downstream till a safe sampling spot is found.
4. Visually define a rectangular quadrat that is one net width wide and two net widths long at the
sampling point. The area within this quadrat is -0.5 m2.
4. Inspect the stream bottom within the quadrat for any heavy organisms, such as mussels and
snails. Remove these organisms by hand and place them into the net or into a bucket labeled
"POOL".
5. Vigorously kick the substrate within the quadrat with your feet while dragging the net repeat-
edly 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 20 seconds. NOTE: If there is too little water to use the kick net, stir up the
substrate with your gloved hands and use the U.S. Standard #30 sieve to collect the organisms
from the water in the same way the net is used in larger pools.
6. After 20 seconds, hold the net between your legs and partially submerged. Pick up any loose
rocks within the quadrat. Rub or brush any organisms found on them into the net. Also re-
check the area for any additional snails or clams and place them in the net.
7. Invert the net into a bucket labeled "POOL", which is about half full of water, to rinse organisms
out of the net. Inspect the net for clinging organisms. Use watchmakers' forceps to remove
any organisms from the net and place them in the bucket. 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 any organisms.
8. Place an "X" in the appropriate habitat type and sampling point boxes for the transect on the
Sample Collection Form.
9. Proceed upstream to the next transect and repeat Steps 1 through 8. Combine all kick net
samples from "pool/glide" habitats into the "POOL" bucket.
153
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EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates), Rev. 4, Sept. 1998 Page 8 of 14
SAMPLE COLLECTION FORM - STREAMS
SITE NAME: MlLL CjtffK.
SITE ID: MAIA97-_?._2_-2
DATE: 7 / tfl 97 VISIT: 01 D2
TEAM ID,(X):
mi D2 D3 D4 D5 D6 D7
D8
COMPOSITE BENTHOS SAMPLES
SAMPLE ID
(BARCODE)
HABITAT
(XONE)
R P
J- A.3L O-Q.-L *
A a. ? o o _£. *
STATION A
RIFFLE OR POOL-
PC ONE) -
LEFT, CENTER, OR
RIGHT -
(XONE)-
B C
OF
JARS
z
1
FLAG
COMMENTS
D E F
HR DR DR HR DR
DP HP HP DP HP
DL HL DL DL HL
DC DC HC DC DC
HR DR DR HR DR
G H
DR DR
HP HP
DL DL
HC DC
DR HR
1 J
DR HR
HP DP
HL DL
DC HC
DR DR
K
COMPOSITE PERIPHYTON SAMPLES HABITAT TYPE (X) - @JR|FFLE D POOL D OTHER
SAMPLE ID (BARCODE) -
ASSEMBLAGE ID
(50-MLTUBE)
SUB. SAMPLE VOL.
f O ML
JLA_?._£
__0.jt_ COMPOSITE VOLUME - 3- O O ML
CHLOROPHYLL
(GF/F FILTER)
VOL. FILTERED
A. _r_ ML
COMPOSITE PERIPHYTON SAMPLES
SAMPLE ID (BARCODE) -
ASSEMBLAGE ID
(50-MLTUBE)
SUB. SAMPLE VOL.
5~ O ML
_*._2__2__e..0-_3.
CHLOROPHYLL
(GF/F FILTER)
VOL. FILTERED
A f uL
BlOMASS
(TARED FILTER)
FILTER No.
VOL. FILTERED
?99 A-JLui.
APA SAMPLE
(50-MLTUBE)
SUB. SAMPLE VOL.
£_Q_ ML
HABITAT TYPE (X)- D RIFFLE HPOOL D OTHER
COMPOSITE VOLUME - 3 O & ML
BlOMASS
(TARED FILTER)
FILTER No.
VOL. FILTERED
/OOI Z f ML
APA SAMPLE
(50-MLTUBE)
SUB. SAMPLE VOL.
5" O ML
COMMENTS:
Fligcodn: K» Samplo not collected; U=Suspoctsamplo; F1,F2,eto.= misc. flag assigned by Reid crew. Explain all flags In Comment sections.
R«v. 06/02/97 (*t_«aco.97)
SAMPLE COLLECTION FORM - STREAMS -1
Figure 11-3. Sample Collection Form (page 1), showing information for benthic macro-
invertebrate samples.
154
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EMAP-SW-Slreams Field Operations Manual, Section 11 (Benthic Macroinvertebrates), Rev. 4, Sept. 1998 Page 9 of 14
Date:
Time:
Site No.:
MACROINVERTEBRATE SAMPLING ACTIVITIES CHECKLIST
Stream Name and Location:
Crew ID: 123456
Collector:
1.
Initial observations, if any, on the Sample Collection Form - Streams.
2.
Composite riffle/run sample collected with a label inside the jar.
3.
Composite pool/glide sample collected with a label inside the jar.
4.
Correct barcode and label on all jars and sealed with clear, waterproof tape.
5.
All samples preserved.
6.
With a grease pencil write site number, sample type (Riffle or Pool), and number
of transects sampled for sample type on the cap. If two jars are used be sure to
mark them as such.
7.
Caps are sealed with plastic electrical tape.
8.
Photos of the site.
9.
Sample jars in cooler or otherwise secured.
10. All equipment accounted for and secured in the vehicle.
Signature:
Time sampling completed:
Figure 11-4. Checklist for benthic macroinvertebrate sampling activities.
155
-------
EMAP-SW-Streams Field Operations Manual. Section 11 (Benthic Macroinvertebrates), Rev. 4, Sept. 1998 Page 10 of 14
substrate at the sampling point. Place the contents of this hand-picked sample into either
the "RIFFLE" or "POOL" bucket.
11.2 SAMPLE PROCESSING
After collecting kick net samples from all transects, prepare two composite index
samples from the contents of the "RIFFLE" and "POOL" buckets as described in Table 11-3.
Record tracking information for each composite sample on the Sample Collection Form as
shown in Figure 11-3. A set of completed sample labels, including the label that is used if
more than one jar is required for a single composite sample, is shown in Figure 11-5. Note
that each composite sample has a different sample number (barcode). The ID number is
also recorded on a waterproof label that is placed inside the jar (Figure 11-5, lower right). If
more than one jar is used for a composite sample, a special label (Figure 11-5, lower left) is
used to record the ID number assigned to the sample. DO NOT use two different barcode
numbers on two jars containing one single sample. Blank labels for use inside of sample
jars are presented in Figure 11-6. These can be copied onto waterproof paper.
Complete the check-off sheet (Figure 11-4). Check to be sure that the prenumbered
adhesive barcoded label is on the jar and covered with clear tape, and that the waterproof
label is in the jar and filled in properly. Be sure the inside label and outside label describe
the same sample. Replace the cap on each jar and seal them with plastic electrical tape.
Check to make sure the cap is properly marked with site number, habitat type (pool or
riffle), and number of transects sampled. Record any additional pertinent information in the
"Field Notes" section of the checklist. Place the samples in a cooler or other secure con-
tainer for transporting and/or shipping the laboratory (see Section 3). The container and
absorbent material should both be suitable for transporting ethanol. Check to see that all
equipment is in the vehicle.
11.3 EQUIPMENT AND SUPPLY CHECKLIST
Figure 11-7 shows the checklist of equipment and supplies required to complete the
collection of benthic macroinvertebrates from streams. This checklist is similar to the
checklist presented in Appendix A, which is used at the base location (Section 3) to ensure
that all of the required equipment is brought to the stream. Use this checklist to ensure that
equipment and supplies are organized and available at the stream site in order to conduct
the activities efficiently.
156
-------
EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates). Rev. 4, Sept. 1998 Page 11 of 14
TABLE 11-3. PROCEDURE FOR PREPARING COMPOSITE SAMPLES FOR
BENTHIC MACROINVERTEBRATES
~ Pour the entire contents of the "RIFFLE" bucket through a U.S. Standard #30 sieve (or sieve-
bottomed bucket with 595 urn mesh size). Remove any large objects and wash off any cling-
ing organisms back into the sieve before discarding.
!. Using a wash bottle filled with stream water, rinse all the organisms from the bucket into the
sieve. This is the composite sample for that habitat (riffle or pool) for the site.
!. Estimate the total volume of the sample in the sieve and determine how large a jar will be
needed for the sample (half gallon or gallon). Do not use more than one jar for each of the
samples unless it cannot be avoided.
k Fill in a "Composite Benthos" sample label with the stream ID and date of collection. Circle the
habitat type (Riffle or Pool). Attach the completed label to the jar and cover it with a strip of
clear tape.
k 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 VA full, or use a second jar if a larger one is not available. Carefully examine the sieve for
any remaining organisms and use watchmakers' forceps to place them into the sample jar.
If a second jar is needed, fill in a sample label that does not have a pre-printed barcode
number on it. Record the barcode number from the pre-printed label prepared in Step 4
in the "BARCODE" field of the label. Attach the label to the second jar and cover it with a
strip of clear tape.
>. Add 95% ethanol to each jar so that the final concentration of ethanol is at least 70%. If there
is a small amount of water in the sample, it may not be necessary to fill the jar entirely full to
reach a 70% concentration. It is very important that sufficient ethanol be used to reach a 70%
concentration. Otherwise, the organisms will not be properly preserved.
NOTE: Prepared composite samples can be transported back to the vehicle before
adding ethanol if necessary.
5. Place a waterproof label with the following information inside each jar:
7.
8.
9.
Stream Number
Type of sampler and mesh size used
Habitat type (riffle or pool)
Name of stream
Date of collection
Collectors initials
Number of transect samples
composited
Replace the cap on each jar. Seal each jar with plastic tape. Use a grease pencil to write the
site number, sample type (Riffle or Pool), and number of transects on the cap of each jar.
Repeat Steps 1 through 7 for the "POOL" bucket.
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. -
157
-------
EMAP-SW-Streams Field Operations Manual. Section 11 (Benthic Macroinvertebrates), Rev. 4, Sept. 1998 Page 12 of 14
COMPOSITE BENTHOS
SITE ID: MAIA_17--_2212__
DATE: 7 I /*"/ 98
HABITAT: (Tliffle) Pool
229001
COMPOSITE BENTHOS
SITE ID: MAIA ?7 •
DATE: _
HABITAT: (Riffle Pool
BARCODE:
COMPOSITE BENTHOS
SITE ID: MAIA 97 -
DATE: 7 / If I 98
HABITAT: Riffle <^ooT}
229002
BENTHOS IDENTIFICATION
Site number A>/}
-------
EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates). Rev. 4. Sept. 1998 Page 13 of 14
BENTHOS IDENTIFICATION
Site Number
Stream
Collection Date
Sampler
Habitat Type
Collector(s)
Number of Transects
BENTHOS IDENTIFICATION
Site Number '
Stream
Collection Date
Sampler
Habitat Type
Collector(s)
Number of Transects
BENTHOS IDENTIFICATION
Site Number
Stream
Collection Date
Sampler
Habitat Type
Collector(s)
Number of Transects
BENTHOS IDENTIFICATION
Site Number
Stream
Collection Date
Sampler
Habitat Type
Collector(s)
Number of Transects
BENTHOS IDENTIFICATION
Site Number
Stream
Collection Date
Sampler
Habitat Type
Collector(s)
Number of Transects
BENTHOS IDENTIFICATION
Site Number •
Stream
Collection Date
Sampler
Habitat Type
Collector(s)
Number of Transects
Figure 11-6. Blank labels for benthic invertebrate samples.
159
-------
EMAP-SW-Streams Field Operations Manual, Section 11 (Benthic Macroinvertebrates), Rev. 4, Sept. 1998 Page 14 of 14
EQUIPMENT AND SUPPLIES FOR BENTHIC MACROINVERTEBRATES
OTY ITEM
1
1
2
1
1
2pr.
1
1
1
4 to 6
each
2 gal
2pr.
1
2
4
6
1
1
1 pkg.
4 rolls
1
1
1
1 pkg.
1 copy
1 set
Modified kick net ( closed bag with 595/600 urn mesh) and 4-ft handle (Wildco
#425-C50)
Spare net(s) for the kick net sampler or extra sampler
Watch with timer or a stopwatch
Buckets, plastic, 8- to 1 0-qt capacity, labeled "RIFFLE" and "POOL"
Sieve, U.S. Standard 30
Sieve-bottomed bucket, 595-um mesh openings
Watchmakers' forceps
Wash bottle, 1-L capacity labeled "STREAM WATER"
Small spatula, spoon, or scoop to transfer sample
Funnel, with large bore spout
Sample jars, plastic with screw caps, Vz and 1 gallon capacity, suitable for use
with ethanol
95% ethanol, in a proper container
Rubber gloves, heavy rubber
Cooler (with suitable absorbent material) for transporting ethanol and samples
Composite Benthic sample labels, with preprinted ID numbers (barcodes)
Composite Benthic sample labels without preprinted ID numbers
Blank labels on waterproof paper for inside of jars
Sample Collection Form for site
Field check list sheet
Soft (#2) lead pencils
Fine-tip indelible markers
Grease pencils
Clear tape strips
Plastic electrical tape
Knife, pocket, with at least two blades
Scissors
Pocket-sized field notebook (optional)
Kim wipes in small self-sealing plastic bag
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for benthic
macroinvertebrates
Figure 11-7. Equipment and supply checklist for benthic macroinvertebrates.
160
-------
SECTION 12
AQUATIC VERTEBRATES
by
Frank H. McCormick1 and Robert M. Hughes2
Sampling amphibian and fish species to determine their proportionate abundances
and the presence of external anomalies is conducted after all other field sampling and
measurement activities are completed. The objective is to collect a representative sample
of all except very rare species in the assemblage. Backpack electrofishing equipment is
used as the principal sampling gear (Section 12.1.1), supplemented by block netting (when
necessary) and seining (Section 12.1.2) in habitats where flow, substrate and structure
affect capture of benthic species. All team personnel are involved in collecting aquatic
vertebrates. In addition to gathering data on the assemblage, fish specimens are retained
for analysis of tissue contaminants (Section 13).
12.1 SAMPLE COLLECTION
The entire channel within the sampling reach is sampled. Complex, very large, or
wide systems without clearly-defined habitat types are sampled through use of transects so
that effort is distributed along the entire reach relative to the mean width of each transect,
as illustrated in Figure 12-1. Fish and other aquatic vertebrates are collected according to
time and distance criteria. Collection time should continue for not less than 45 minutes and
not longer than 3 hours within the defined sampling reach (Section 4) to obtain a represen-
tative sample. Sampling information is recorded on the Vertebrate Collection Form (Figure
12-2). Record general comments (perceived fishing efficiency, missed fish, gear operation,
suggestions) on the blank lines of the form.
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin Luther King
Dr., Cincinnati, OH 45268.
Dynamac International Corp., 200 SW 35th St., Corvallis, OR 97333
161
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates). Rev. 4, September 1998 Page 2 of 22
Stippled areas represent habitats too hazardous to sample.
A-B B-C
J-K
Transects correspond to cross-section transects established every 10 channel widths or 15 m
Transect
A-B
B-C
C-D
D-E
E-F
F-G
G-H
H-l
I-J
J-K
Mean
Transect
Width (m)
8m
9m
12m
12m
15m
15m
15m
15m
10m
8m
Time
Allotment
(sec)
8*84= 672
9*84= 756
12*84=1008
12*84=1008
15*84=1260
15*84=1260
15*84=1260
15*84=1260
10*84=840
8*84= 672
Shock
Time
(estimated)
350
400
600
600
800
800
800
800
500
350
Example Calculation
Sum of Mean Transect Widths
8 + 9 + 12 + 12 + 15 + 15 + 15 + 15 + 10 + 8 = 128m.
3 hrs = 10800s sampling time.
10800s / 128m = 84 s/m of mean width.
Multiply mean width by # units to calculate time in sec-
onds to be spent in each transect.
Depending on complexity of habitat, actual shock time
may vary from 50% - 75% of fishing time.
Figure 12-1. Index sample design for allocating aquatic vertebrate sampling effort in very
complex or very large wadeable streams. Note distribution of effort in narrow and wide
sections.
162
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates). Rev. 4, September 1998 Page 3 of 22
12.1.1 Eiectrofishing
Because fishes and amphibians are collected using portable electrofishing units,
safety procedures must be followed meticulously at all times (refer to Section 2). Primary
responsibility for safety while electrofishing rests with the team leader. Electrofishing units
have a high voltage output and may deliver a dangerous electrical shock. While
electrofishing, avoid contact with the water unless sufficiently insulated against electrical
shock. Use chest waders with nonslip soles and watertight rubber (or electrician's) gloves
that cover to the elbows. If they become wet inside, stop fishing until they are thor-
oughly dry. Avoid contact with the anode and cathode at all times due to the poten-
tial shock hazard. If you perspire heavily, wear polypropylene or some other wicking and
insulating clothing instead of cotton. While electrofishing avoid reaching into the water. If it
is necessary for a team member to reach into the water to pick up a fish or something that
has been dropped, do so only after the electrical current has been interrupted and the
anode is removed from the water. Do not resume electrofishing until all individuals are
clear of the electroshock hazard. The 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 turning off the electricity.
Avoid operating electrofishing equipment near unprotected people, pets, or livestock.
Discontinue activity during thunderstorms or heavy rain. Team members should keep each
other in constant view or communication while electrofishing. For each site, know the
location of the nearest emergency care facility. Although the team leader has authority,
each team member has the responsibility to question and modify an operation or decline
participation if it is unsafe. Use hand signals to communicate direction and power on or off
when using generators.
Gasoline is extremely volatile and flammable. Its vapors readily ignite on contact
with heat, spark or flame. Never attempt to refill the generator while it is running. Always
allow the generator to cool before refilling. Keep gasoline out of direct sunlight to re-
duce volatilization and vapor release. Always wear gloves and safety glasses when han-
dling gasoline. Keep gasoline only in approved plastic containers and store in a tightly
closed container in safety cabinet or cooler lined with vermiculite.
163
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 4 of 22—
The procedure to sample with the backpack electrofisher unit is presented in Table
12-1. Record information on the Vertebrate Collection Form as shown in Figure 12-2.
If the stream cannot be sampled by either electrofishing or seining, complete the "NOT
FISHED" field on the form. Select the initial voltage based on the measured conductivity of
the stream (see Section 5). Select the initial frequency based on the expected size of fish.
If fishing success is poor, increase the pulse width first and then the voltage. Increase the
frequency last to minimize mortality or injury to large fish.
Determine that all team members are wearing waders and gloves and are clear of
the anode. Wear polarized sunglasses to aid vision. Start the electrofisher, set the timer to
zero, and depress the switch to begin fishing. Starting at the bottom of the reach, fish in an
upstream direction. Adjust voltage and waveform output according to sampling effective-
ness and incidental mortality to specimens. The backpack unit is equipped with an audio
alarm that sounds when the output voltage exceeds 30 V. It also serves as an input current
indicator for pulse cycles greater than 5Hz. It begins as a strong continuous tone and
begins to beep slowly at currents of 1.25 amps. It beeps faster as input current increases.
In case of an overload (in excess of 3 amps), the beep becomes very rapid and the over-
load indicator comes on. Release the anode switch and adjust voltage and waveform and
continue fishing.
When fishing, slowly sweep the electrode wand from side to side in the water in
riffles and pools. Sample available cut-bank and snag habitat areas as well as riffles and
pools. Move the wand in and out of large snags or deep cuts or release the electrode
switch, move the wand away slightly, depress the switch again and sweep the wand away
from the cover to draw fish out into open. In fast, shallow water, it may be more effective to
use a seine as a block net; sweep the anode and fish downstream into the net.
In extremely wide streams, it may be necessary to work from the midline of the
stream channel to the banks. Be sure that deep, shallow, fast, slow, complex, and simple
habitats are all sampled. In stretches with deep pools, fish the margins of the pool as much
as possible, being extremely careful not to step into deep water. In larger or more complex
streams, allocate the fishing time between transects based on differences in the mean
wetted width of the stream (Figure 12-1).
One or two netters follow along beside or slightly behind the person operating the
electrofisher (on the anode side). Each netter uses an insulated dip net to retrieve stunned
individuals, which are then deposited into a bucket for later processing (Section 12.3).
164
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 5 of 22
TABLE 12-1. PROCEDURE TO COLLECT AQUATIC VERTEBRATES BY ELECTROFISHING
Survey the sampling reach and set block nets at each end (Transects "A" and "K") if necessary
(e.g., the majority of the reach is one large, continuous pool). If necessary, allocate the total
shocking time among transects based on mean stream widths.
2.
3.
4.
6.
9.
Complete the header information on a copy of the Vertebrate Collection Form. Indicate that all
transects are being sampled in the "TRANSECT" field on the form.
NOTE: Make an effort to search and sample for aquatic vertebrates at all streams, even if
the stream is extremely small, and it appears that sampling may not collect any speci-
mens. If no specimens are collected, complete the "NONE COLLECTED" field on the
Vertebrate Collection Form. Provide an explanation in the comments section of the form.
If the conductivity measured during the water chemistry sampling is less than 10 uS/cm, or if
the depth or velocity make electrofishing unsafe, sample by seining if possible, otherwise do
not sample. If you do not sample, complete the "NOT FISHED" field on the Vertebrate Collec-
tion Form. Provide an explanation in the comments section of the form.
Set unit to 300 volt-amperes (VA) and pulsed DC. Select initial voltage setting (150-400 V for
high conductivity [>300 uS/cm]; 500-800 V for medium conductivity [100 to 300 uS/cm]; 900-
1100 V for low conductivity [<100 uS/cm] waters). In waters with strong-swimming fish (length
>200 mm), use a frequency of 30 Hz with a pulse width of 2 msec. If mostly small fish are
expected, use a frequency of 60-70 Hz. Start the generator, set the timer, and depress the
switch to begin fishing.
Beginning at the downstream end of the reach (Transect "A"), fish in an upstream direction,
parallel to the current. Depress the switch and sweep the electrodes from side to side in the
water. Sample available cut-bank and snag habitats as well as riffles and pools.
The netters follow the operator and net stunned aquatic vertebrates. Deposit individuals in
buckets for processing. If necessary, use seines to block riffles, pools and snags. The opera-
tor should adjust voltage and waveform output according to sampling effectiveness and the
mortality of fish specimens.
Continue upstream until the next transect is reached. In large or complex streams, allocate the
fishing time between the two transects as calculated based on the mean transect widths (Step
1). Process fish after each transect to reduce mortality.
Repeat Steps 5 through 7 until Transect "K" is reached. Record the following on the Verte-
brate Collection Form:
The reading from the electrofisher timer in the "TOTAL SHOCK TIME" field on the
Vertebrate Collection Form.
The total distance sampled by electrofishing
The total fishing time, if no additional sampling is conducted (e.g., by seining) once
electrofishing is completed. Total sampling time should be between 45 minutes
and 3 hours.
If no aquatic vertebrates were collected, complete the "NONE COLLECTED" field on the
Vertebrate Collection Form.
165
-------
EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 6 of 22
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166
-------
EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates). Rev. 4, September 1998 Page 7 of 22
Change the water in the bucket periodically to minimize mortality prior to processing. If
individuals show signs of stress (loss of righting response, gaping, gulping air, excessive
mucus), stop and process them. This should only be necessary on very warm days, in long
reaches, or if very large numbers of aquatic vertebrates are collected. Electrofishing may
also need to cease at times to immediately process and release specimens (e.g., listed
species or large game fish) as they are netted (see Section 12.2). If periodic processing is
required, be sure to release individuals downstream to reduce the likelihood of collecting
them again.
At the completion of electrofishing, record the total operating time (shock time)
shown on the electrofisher timer and the distance sampled by electrofishing on the Verte-
brate Collection Form (Figure 12-2). if sampling activities (electrofishing and seining) are
completed, also record the total fishing time on the Vertebrate Collection Form. If no
aquatic vertebrates were collected, indicate this on the form as shown in Figure 12-2.
12.1.2 Seining
Seining may be used in conjunction with electrofishing to ensure sampling of those
species which may otherwise be underrepresented by an electrofishing survey alone (e.g.,
darters, sculpins, madtoms, and benthic cyprinids). Seining may also be used in sites
where the stream is too deep for electrofishing to be conducted safely or in turbid, simple,
soft-bottomed streams where it is more effective.
Seining procedures are presented in Table 12-2. Depending on the particular use
(block netting vs. active seining) and the habitat, different sizes of seines are used. In riffle
habitats, the seine is held stationary while team members disturb the substrate immediately
upstream of the net. In pools, the seine is pulled back and forth across the pool, using the
shore and other natural habitat breaks as barriers, or pulled rapidly downstream through the
pool and then swept toward the shore. Block nets may be used in very large pools to limit
escape or as seines. Large nets are typically deployed parallel to the current and swept to
shore.
Proceed upstream through the reach, allocating the seining effort among habitat
areas (riffles and pools) so that the entire reach is sampled within the required sampling
time (45 minutes to 3 hours). Deposit aquatic vertebrates collected by seining into a bucket
for later processing as described in Section 12.1. At the completion of sampling activities
(electrofishing and/or seining), record the total fishing time on the Vertebrate Collection
167
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates), Rev. 4. September 1998 Page 8 of 22
TABLE 12-2. PROCEDURES TO COLLECT AQUATIC VERTEBRATES BY SEINING
NOTE: Seining is used in place of electrofishing where a stream is too deep for electrofishing to be
conducted safely, or in turbid, simple, soft-bottomed streams where it is more effective.
1. Survey the sampling reach and set block nets at each end (Transects "A" and "K") if necessary
(e.g., the majority of the reach is one large, continuous pool). Allocate the sampling effort
throughout the sampling reach so that the total fishing time will be between 45 minutes (small
stream) and 3 hours (large stream).
2. Complete the header information on a copy of the Vertebrate Collection Form. Indicate that all
transects are being sampled in the "TRANSECT" field on the form.
NOTE: Make an effort to search and sample for aquatic vertebrates at all streams, even if
the stream is extremely small, and it appears that sampling may not collect any speci-
mens. If no specimens are collected, complete the "NONE COLLECTED" field on the
Vertebrate Collection Form. Provide an explanation in the comments section of the form.
3. Begin at the downstream end of the sampling reach (Transect "A"). Proceed upstream, sam-
pling available riffle and pool habitats using the appropriate method below:
3A. Riffle habitats- Use a small minnow seine (2 m long * 1.25 m wide; 0.6 cm mesh size).
1. One or two persons place the seine perpendicular to the current across the down-
stream end of the riffle. Ensure that the lead line is on the bottom. Tilt the net
slightly backward to form a pocket to trap aquatic vertebrates.
2. Starting about 2 m upstream, the other two team members disturb the substrate in
front of the net by kicking through the substrate and overturning rocks, and proceed
downstream toward the nets.
3. Raise the net and examine it carefully for aquatic vertebrates.
3B. Pool habitats: Use a larger seine (3 m long * 2 m wide; 0.6 cm mesh size).
1. Two people pull the seine back and forth across the pool, using the shore and other
natural habitat breaks as barriers.
2. Alternatively (in areas with some current), pull the net along in a downstream direc-
tion and then sweep toward the shore.
3. Pull the net onto the shore and examine it carefully for aquatic vertebrates.
4. Deposit individuals in buckets for processing, and continue upstream to the next habitat area.
5. Repeat Steps 3 and 4 for successive habitat areas until Transect "K" is reached. If no aquatic
vertebrates were collected by either seining or electrofishing, complete the "NONE COL-
LECTED" box on the Vertebrate Collection Form.
6. Record the total fishing time on the Vertebrate Collection Form. Include any time spent
electrofishing in the total fishing time. Total fishing time should be between 45 minutes and 3
hours
168
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 9 of 22
Form (Figure 12-2). If no aquatic vertebrates were collected, indicate this on the form as
shown in Figure 12-2.
12.2 SAMPLE PROCESSING
Sample processing involves tallying and identifying fish and amphibians, examining
individual specimens for external anomalies, obtaining length measurements from selected
specimens, preparing voucher specimens for taxonomic confirmation and archival at a
museum, and selecting specimens to prepare samples for fish tissue contaminants (see
Section 13). Process collections as quickly as possible to minimize stress to live speci-
mens. All team members can work to separate aquatic vertebrates into families or obvious
"morphotypes". Alternatively, 1 or 2 persons can process fish from one bucket while the
other team members continue to collect fish and deposit them into a second bucket. Once
the rough sort has been completed, one person can identify, measure, and examine individ-
uals while another person may record information on the field data forms.
12.2.1 Taxonomic Identification and Tally
Table 12-3 presents the procedure for identifying and tallying aquatic vertebrates.
Record identification and tally data for each species on the Vertebrate Collection Form as
shown in Figure 12-2. Record comments and data for additional species on page 2 of the
Vertebrate Collection Form (Figure 12-3). Each team needs to be provided with a list of
standardized names (required) and species codes (optional) for aquatic vertebrate species
that are expected to be collected (see Appendix D for an example).
Sort aquatic vertebrates by species into small buckets and containers. Taxonomic
identification should be performed only by trained ichthyologists familiar with the fish spe-
cies and other aquatic vertebrate taxa of the region. Use taxonomic reference books and
other materials that contain species descriptions, ranges, and identification keys to make
species identifications in the field. Try to process one species completely before going on
to the next. However, where there are many individuals of easily identified species, pro-
cessing may be facilitated by keeping a tally count of the number of individuals of each
species and totaling the tally once processing is complete.
To minimize handling, process threatened and endangered species first, and
immediately return all individuals to the stream. If conditions permit and stress to
individuals will be minimal, photograph such fish for voucher purposes (Section 12.2.3).
169
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 10 of 22
TABLE 12-3. PROCEDURE TO IDENTIFY, TALLY, AND EXAMINE AQUATIC VERTEBRATES
1. Separate aquatic vertebrates retained in collecting buckets or live wells into families or obvious
"morphotypes" (e.g., two dorsal fins vs. one, a sucker mouth, catfish, trout, etc.). Place each
group into a separate bucket or similar container. All team members can participate in this
"rough" sort. Alternatively, identify and process each individual completely, thus handling it
only once.
2. Sort each group created in Step 1 by species into separate containers. This should be done
only by team members who are trained ichthyologists familiar with the fish species and other
aquatic vertebrate taxa of the region.
3. Select a container and record the common name (from a standardized list) and species code (if
required) on the first blank line in the "SPECIMENS" section of the Vertebrate Collection Form.
If a species cannot be positively identified, assign it an "unknown" species code from the list
provided.
NOTE: Process species listed as threatened and endangered first and return individuals
immediately to the stream. Photograph specimens for voucher purposes if conditions
permit and stress to individuals will be minimal. Indicate if photographed on Vertebrate
Collection Form. If individuals have died, prepare them as voucher specimens and pre-
serve in formalin. Notify the appropriate state officials as soon as possible.
4. Tally the number of individuals collected (use the "TALLY" box on the Vertebrate Collection Form
if necessary) and record the total number in the "COUNT" field on the form.
5. Measure the total length of the largest and smallest individual to provide a size range for the
species. Record these values in the "LENGTH" area of the Vertebrate Collection Form.
6. If the container has sport fish and other very large specimens, or if 3 or fewer species are
captured at the stream, prepare a Vertebrate Length Recording Form.
A. Complete the header information on the form, then enter the common name (from a
standardized list) and the species code (if required) in the first blank line.
B. Measure the total length of each individual (up to 30) and record the lengths in the boxes
on the form (2 lines of boxes per species). For smaller species, measure and record
lengths of a random set (up to 30) of the individuals collected.
7. Examine each individual for external anomalies and note the types of anomalies observed. After
all of the individuals of a species have been processed, record the anomaly code and the total
number of individuals affected in the "ANOMALIES" area of the Vertebrate Collection Form.
8. Record the total number of mortalities due to electrofishing or handling on the Vertebrate Collec-
tion Form.
9. Follow the appropriate procedure to prepare voucher specimens and/or to select specimens for
tissue samples. Release all remaining individuals into the stream. If there is still a portion of the
sampling reach that has not been sampled, release fish downstream to avoid their recapture.
10 Rpneat Steos 3 throuah 9 for all other soecies.
170
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates), Rev. 4. September 1998 Page 11 of 22
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171
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 12 of 22—
Indicate if photographed with an "F" series flag for the species on page 1 of the Vertebrate
Collection Form (Figure 12-2) and record a notation in the comments section on page 2 of
the form (Figure 12-3). If protected fish have died, they should be prepared as voucher
specimens and preserved in formalin. Notify the appropriate state officials as soon as
possible.
If a species cannot be confidently identified in the field (e.g., small individuals or
suspected hybrids), record it as an "unknown" species on the Vertebrate Collection Form,
using one of the names (and code, if required) provided for unknowns from the standard-
ized list (see Figure 12-2 for an example). If possible, flag unknown species with an "F"
series flag and provide your best guess at an identification in the comments section of the
Vertebrate Collection Form (Figure 12-3).
12.2.2 External Examination and Length Measurements
During the tallying procedure for each species (Table 12-3), examine each individual
for the presence of external anomalies. External anomalies may result from sublethal
environmental or behavioral stress, diseases, and toxic chemicals. Readily identified exter-
nal anomalies include deformities, eroded fins, lesions, tumors, diseases and parasites.
Codes for different types of anomalies are presented in Table 12-4. Record the types of
anomalies observed and the number of individuals affected on the Vertebrate Collection
Form as shown in Figure 12-2.
Blackening and exopthalmia may occasionally result from electrofishing. Injuries
due to sampling are not included in the tally of external anomalies, but should be noted in
the comments section of the Vertebrate Collection Form (Figure 12-3). Care should be
taken in the early stages of electrofishing to use the most effective combination of voltage
and pulse width while minimizing injury to fish. Blackening from electrofishing usually
follows the myomeres or looks like a bruise. If fish die due to the effects of sampling or
processing, record the number for each species on the Vertebrate Collection Form (Figure
12-2).
For each species, use a measuring board or ruler to determine the total length
(Figure 12-4) of the largest and smallest individuals. Measure individuals on right side, and
slide fish to touch the "Bump Board" on the measuring board. Measure total length to the
nearest millimeter (mm) and record these values on the Vertebrate Collection Form as
shown in Figure 12-2. For sport fish and other larger species, measure the total lengths of
172
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 13 of 22
TABLE 12-4. EXTERNAL ANOMALY CATEGORIES AND CODES
Categories
Absent
Blisters
Blackening
Extensive black spot
disease
Cysts
Copepqd
Deformities
Eroded fins
Eroded gills
Fungus
Fin anomalies
Grubs
Hemorrhaging
Ich
Lesions
Lice
Mucus
None
Other
Scale anomalies
Shortened operculum
Tumors
Leeches
Exophthalmia
Code
AB
BL
BK
BS
CY
CO
DE
EF
EG
FU
FA
GR
HM
1C
LE
LI
MU
NO
OT
SA
SO
TU
WR
EX
Definition
Absent eye, fin, tail.
In mouth, just under skin.
Tail or whole body with darkened pigmentation.
Small black cysts (dots) all over the fins and body.
Fluid-filled swellings; may be either small or large dots.
A parasitic infection characterized by a worm-like copepod embed-
ded in the flesh of the fish; body extends out and leaves a
sore/discoloration at base, may be in mouth gills, fins, or any-
where on body.
Skeletal anomalies of the head, spine, and body shape; amphibi-
ans may have extra tails, limbs, toes.
Appear as reductions or substantial fraying of fin surface area.
Gill filaments eroded from tip.
May appear as filamentous or "fuzzy" growth on the fins, eyes, or
body.
Abnormal thickenings or irregularities of rays
White or yellow worms embedded in muscle or fins.
Red spots on mouth, body, fins, fin bases, eyes, and gills.
White spots on the fins, skin or gills.
Open sores or exposed tissue; raised, granular, or warty out-
growths.
Scale-like, mobile arthropods.
Thick and excessive on skin or gill, or as long cast from vent.
No anomalies present.
Anomalies or parasites not specified.
Missing patches, abnormal thickenings, granular skin
Leaves a portion of the gill chamber uncovered
Areas of irregular cell growth which are firm and cannot be easily
broken open when pinched. (Masses caused by parasites can
usually be opened easily.)
Annelid worms which have anterior and posterior suckers. They
may attach anywhere on the body.
Bulging of the eye.
173
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 14 of 22
Standard Length
Figure 12-4. Fish length measurements.(modified from Lagler, 1956).
174
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates). Rev. 4, September 1998 Page 15 of 22
up to 30 individuals and record these values on the Vertebrate Length Recording Form as
shown in Figure 12-5. If less than four species (large or small) are collected, randomly
select up to 30 individuals of each species and determine the total length of each individual.
Record these length measurements on the length recording form (Figure 12-5).
12.2.3 Preparing Voucher Specimens
With the exception of very large individuals of easily identified species, voucher
collections of up to 25 individuals (where allowed by collecting permits) of all species are
made to provide a permanent, archived, historical record offish collections. Prepare the
voucher sample for a site according to the procedure presented in Table 12-5. Retain
additional specimens of the appropriate species for the fish tissue contaminants sample
(Section 13). For each species, voucher specimens take priority over specimens for
the tissue contaminants sample.
The number of voucher specimens and the method of vouchering varies with spe-
cies. Large, easily identified species, larger species that are difficult to identify in the field,
or species that are uncommon in the region require a few specimens of both adults and
juveniles, if both were collected. Very large specimens, especially of easily identified game
fish, are "vouchered" by photographing them and then releasing them alive. A larger num-
ber of voucher specimens are required for smaller species, which are typically more difficult
to identify in the field. As stated previously, species of "special concern" (state and federally
protected species), are processed first, vouchered by photography, and released alive.
Include any individuals of protected species that die before they can be processed and
released as part of the preserved voucher sample for the stream. In some cases, special
restrictions may apply to protected species (e.g., sampling may have to cease upon collect-
ing an individual). These restrictions will be stipulated on the scientific collecting permits
issued by state and federal agencies.
Individuals selected as voucher specimens are first anaesthetized in a concentrated
solution of carbon dioxide. Voucher specimens for each species are counted and placed
into individual nylon mesh bags (1 bag per species). Nylon stockings or panty hose may be
substituted in place of nylon bags. Each bag contains a numbered tag (Figure 12-6). Re-
cord the tag number and the number of individuals vouchered for each species on the
Vertebrate Collection Form as shown in Figure 12-2.
175
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates). Rev. 4, September 1998 Page 16 of 22
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176
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 17 of 22
TABLE 12-5. GUIDELINES AND PROCEDURES FOR PREPARING
AQUATIC VERTEBRATE VOUCHER SPECIMENS
1. Determine the voucher category of a species and the number of specimens to include in the
voucher sample based on the following guidelines. NOTE: Category 3 species should be
processed first.
A Category 1 — Large easily identified species OR adults may be difficult to identify OR the
species is uncommon in that region. Examples include:
American Eel
Sturgeon
Paddlefish
Gars
Bowfin
Mooneye and
Goldeye
White Sucker
Longnose Sucker
Hogsucker
Quillback
Carpsuckers
Moxostoma spp.
Buffalo fishes
Bullhead catfish
Channel catfish
Esocids
Morone spp
Shads
Drum
Carp
Salmonids
Crappies
Micropterus spp.
Walleye and Sauger
B.
1. Preserve 1-2 small (<150 mm total length) adult individuals per site plus 2-5 juve-
niles. If only large adults are collected, reserve smallest individuals until voucher
procedure is complete and preserve ONLY if space is available.
• NOTE: Individuals with a total length > 160 mm should be slit on the lower
abdomen of the RIGHT side before placing them into the container.
2. Photograph if considered too large for the jar. All photographs should include (1) a
card with the stream ID, date, species code, and common name, and (2) a ruler or
some other object of known length to provide some indication of the size of the
specimen.
3. Retain additional individuals of primary and secondary target species for the tissue
contaminant sample.
Category 2 — Small to moderate-sized fish OR difficult to identify species. Examples
include:
Lampreys
Cyprinids
Darters
Troutperch
Chubsucker
Topminnows
Sculpins
Sunfish
Silversides
Madtoms
Sticklebacks
Mudminnows
Preserve 25 adults and juveniles. If fewer than 25 individuals are collected,
voucher all of them. Voucher samples take priority over tissue contaminant
sample.
Retain additional individuals of primary and secondary target species for tissue
contaminants sample.
(continued)
177
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 18 of 22
TABLE 12-5 (Continued)
C. Category 3 — Species of "special concern." These are state or federally listed species.
1. Photograph as in Step 1 .A.2 and then release immediately.
2. If specimens have died, proceed to Step 2 and include them as part of the voucher
sample. Flag the species with an "F" series flag on the Vertebrate Collection Form
and note it is a listed species in the comments section of the form. Notify the ap-
propriate state officials as soon as possible.
Place the voucher specimens in a bucket with two carbon dioxide tablets (e.g., Alka Seltzer®)
and a small volume of water. When specimens are anaesthetized, transfer them to a nylon
mesh bag. Record the number of individuals included in the voucher sample in the
"VOUCHERED COUNT" field for the species on the Vertebrate Collection Form.
3. Select a "FISH-BAG" tag that has the same ID number (barcode) as the voucher sample jar
(Step 3). Record the tag number in the "TAG No." field on the corresponding line for the spe-
cies on the Vertebrate Collection Form. Place the tag into the mesh bag and seal.
4. Immediately place the bag into a container (Yz or 1 gal plastic jar) large enough to hold all
voucher specimens. Add a volume of 10% formalin solution equal to the volume of fish.
5. Repeat Steps 1 through 4 for all species collected.
Add additional 10% formalin solution as bags are added so that the final volume of for-
malin solution is equal to the total volume of fish specimens. Use additional jars if
necessary to avoid tight packing and bending of voucher specimens.
6. Prepare two "FISH-JAR" labels (each having the same ID number [barcode]) by filling in the
stream ID and the date of collection. Place one label into the sample jar. Cap tightly and seal
with plastic electrical tape.
7. Attach the second label to the outside of the sample container by covering it with a strip of
clear tape. Record the voucher sample ID number (barcode) on page 1 of the Vertebrate
Collection Form. NOTE: If more than one jar is required, use labels that have the same ID
number printed on them.
8. Place the preserved sample in a suitable container with absorbent material. Store the con-
tainer in a well-ventilated area during transport. Follow all rules and regulations pertaining to
the transport and shipment of samples containing 10% formalin.
2.
178
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates), Rev. 4. September 1998 Page 19 of 22
FISH - JAR
SITE ID: MAIA 160 mm) should be
slit on the lower abdomen of the RIGHT side to allow for complete fixation of internal tissues
and organs. Start with a concentrated solution of formaldehyde and dilute to the final
volume with water. The final volume of 10% formalin in the sample container should equal
the total volume of specimens. Use additional containers if necessary and avoid tight
packing of specimen bags. Delays in carrying out the anaesthetization and preservation
procedures, overpackinq a sample container, or an inadequate volume of preservative will
result in unidentifiable specimens.
Formaldehyde (37%) and formalin (10% formaldehyde by volume) are extremely
caustic agents and may cause severe irritation on contact of vapors or solution with skin,
eyes or mucus membranes. It is a potential carcinogen. Contact with vapors or solution
should be avoided. Wear gloves and safety glasses and always work in a well-ventilated
area. In case of contact with skin or eyes, rinse immediately with large quantities of water.
Store stock solution in sealed containers in safety cabinet or cooler lined with vermiculite. If
possible, transport outside of the passenger compartment of a vehicle.
A set of two sample labels is completed for each sample container as shown in
Figure 12-6. Place one label inside each sample container, and attach the second label to
the outside of the jar with clear tape. Record the sample ID number on the Vertebrate
Collection Form as shown in Figure 12-2. Some museums may also require that a separate
collection card be completed and inserted into each jar of voucher specimens.
179
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 20 of 22
12.3 EQUIPMENT AND SUPPLIES
Figure 12-7 is a checklist of equipment and supplies required to conduct protocols
described in this section. This checklist may differ from the checklists presented in Appen-
dix A, which are used at a base site to ensure that all equipment and supplies are brought
to the stream site. Field teams are required to use the checklist presented in this section to
ensure that equipment and supplies are organized and available to conduct the protocols
efficiently.
12.4 LITERATURE CITED
Lagler, K.R. 1956. Freshwater Fishery Biology. 2nd. Edition. William C. Brown Co.,
Dubuque, Iowa.
McCormick, F.H. 1993. Fish. pp. 29-36 IN: R.M. Hughes (ed.). Stream Indicator Work-
shop. EPA/600/R-93/138. U.S. Environmental Protection Agency, Corvallis, Oregon.
180
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EMAP-SW-Streams Field Operations Manual, Section 12 (Aquatic Vertebrates), Rev. 4, September 1998 Page 21 of 22
EQUIPMENT AND SUPPLIES FOR AQUATIC VERTEBRATES
QTY.
1
3pr
3pr
3pr
2
1
1
2
2
1 set
1
1
1
15-20
1
2ea.
2 gal
1
1pr
1pr
Item
Gasoline or battery-powered backpack electrofishing unit with netted anode
(electrode wand)
Extra battery (charged) or gasoline
Heavy-duty rubber gloves
Chest waders with non-slip soles
Polarized sunglasses
Long-handled dip nets (0.6 cm mesh) with insulated handles
Watch or stopwatch to track elapsed fishing time
Collapsible buckets for holding and processing aquatic vertebrates
Minnow seine (2m x 1 .25 m, 0.6 cm mesh) with brailles
Large seines (3 m x 2 m, 0.6 cm mesh) with brailles
Larger sized seines for block nets (if necessary)
Taxonomic reference books and keys for fishes and amphibians of the region
Fish measuring board or ruler
List of vertebrate species common names (and species codes, if required)
List of external anomaly codes
Small nylon mesh bags for holding voucher specimens (bags can also be con-
structed from sections of nylon stockings or panty hose)
Small fillet knife or scalpel for preparing larger voucher specimens for preserva-
tion
1/2- or 1 -gallon screw-top plastic jars for voucher sample
10% (buffered) formalin solution OR 0.2 gal buffered formaldehyde solution.
Alternatively, fill each voucher sample jar one-half full of 1 0% formalin
Container to hold formalin solution and preserved voucher sample jars
Safety glasses
Chemical-resistant gloves
(continued)
Figure 12-7. Equipment and supplies checklist for aquatic vertebrates.
181
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EMAP-SW-Streams Field Operations Manual. Section 12 (Aquatic Vertebrates), Rev. 4. September 1998 Page 22 of 22
EQUIPMENT AND SUPPLIES FOR AQUATIC VERTEBRATES (Continued)
QTY.
1
4
1
1pr
1 roll
1 pkg.
1 + extras
1 + extras
1
1set
Item
Plastic bucket for anesthetization
Carbon dioxide tablets (Alka-Seltzer® or equivalent)
Sheet of pre-printed jar labels (4) and voucher bag tags (36), all with same
preprinted sample ID number (barcode)
Scissors for cutting labels
Plastic electrical tape
Clear tape strips
Soft lead pencils for recording data and completing tags
Fine-tipped indelible markers for completing sample labels
Vertebrate Collection Form
Vertebrate Length Recording Form
Field operations manual
Laminated sheets of aquatic vertebrate procedure tables and/or quick refer-
Figure 12-7. (Continued).
182
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SECTION 13
FISH TISSUE CONTAMINANTS
by
Roger B. Yeardley1, James M. Lazorchak2, and Frank H. McCormick2
In addition to gathering data on the aquatic vertebrate assemblage (Section 12),
certain specimens of fish are retained for analysis of fish tissue contaminants. In general,
the focus is on fish species that commonly occur throughout the region of interest, and that
are sufficiently abundant within a sampling reach. Two types of composite samples of fish
are prepared at each site (if possible). One composite sample is prepared using individuals
of a Primary Target Species. Primary target species include species of fish whose adults
are small (e.g., small minnows, sculpins, or darters). The second composite sample is
prepared using individuals of a Secondary Target Species. Secondary target species
include species whose adults are of larger size (e.g., suckers, bass, trout, sunfish, carp).
13.1 PREPARING COMPOSITE SAMPLES FOR PRIMARY AND SECONDARY
TARGET SPECIES
To determine the proper quantity for each composite sample, weight is used for the
primary target species and the number of individuals of sufficient size is used for the sec-
ondary target species. Prepare each composite sample using similar sized individuals if
possible. The general rule-of-thumb for "similar size" is that the smallest individual in the
sample should be at least 75% of the total length of the largest individual. Keep this crite-
rion in mind while selecting the final samples. Do not include any obviously small or large
individuals if there is a sufficient sample (weight or number of individuals) without them. If
there is a conflict between criteria, getting a sufficient sample is a higher priority than getting
similar-sized individuals.
SoBran Environmental.lnc., c/o. U.S. EPA, 26 W. Martin Luther King Dr., Cincinnati, OH 45268.
2
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin Luther King
Dr., Cincinnati, OH 45268.
183
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EMAP-SW-Streams Field Operations Manual. Section 13 (Fish Tissue Contaminants), Rev. 4. Sept. 1998 Page 2 of 9
Prepare a composite sample of a primary target species, as described in Table 13-1.
For the primary species composite sample, choose the highest priority target species that
has at least enough individuals to attain the minimum weight (50 g). Get as much weight of
fish as possible within the desired weight range (50-400 g). Do not settle for the minimum
amount (weight) if more fish are present, but instead send as many fish as possible up to
the 400 g weight goal. If there are no primary species with enough individuals available to
meet the desired weight goal, prepare the primary composite sample using individuals of a
small nontarget species for which there are enough individuals available (after vouchering)
to prepare a sample of at least 50 g.
Prepare a composite sample of a secondary target species as described in Table
13-2. For the secondary species composite sample, choose the highest priority target
species that has the desired number (5) of similar-sized individuals (minimum total
length=120 mm) available. If, for any secondary target species, you did not collect 5 individ-
uals of the desired size, prepare the composite sample from a species having 5 individuals
available (including smaller sized individuals). If fewer than 5 fish of any size for any sec-
ondary species are available, prepare the composite sample using as few as 3 fish that are
at least at or near the minimum desired size. If an acceptable secondary target species
sample (by the above criteria) is not available, send only the primary target species sample.
If neither a primary nor secondary species sample that meets these criteria is avail-
able, use your best judgement in preparing some type of fish tissue sample from the avail-
able species collected. Use the procedure for either primary or secondary species, depend-
ing upon the species used and the size range of individuals selected.
Individuals comprising the primary composite sample are wrapped together in alumi-
num foil and placed into a single plastic bag. Each individual comprising the secondary
composite sample is wrapped separately, but all individuals are placed into a single plastic
bag. Each composite sample is labeled as shown in Figure 13-1. Prepare two identical
labels for each composite sample. Double-bag each sample, and place a label on each
bag. Record information about each composite sample on page 2 of the Sample Collection
Form as shown in Figure 13-2. Make sure the sample ID numbers (barcodes) recorded on
the collection form match those on the sample labels.
Tissue samples are stored frozen, using either a portable freezer, a container with
dry ice, or a cooler with several bags of ice. When using ice, double bag the ice and tape
the last bag shut to prevent contamination of samples by melting ice. Store tissue samples
184
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EMAP-SW-Streams Field Operations Manual, Section 13 (Fish Tissue Contaminants), Rev. 4, Sept. 1998 Page 3 of 9
TABLE 13-1. PROCEDURE TO PREPARE THE PRIMARY COMPOSITE SAMPLE FOR
FISH TISSUE CONTAMINANTS
NOTE: If neither a primary nor secondary species sample is available, use your best judge-
ment in sending some type of composite fish tissue sample.
1. After all voucher specimens have been prepared, choose the highest priority primary target
species from the list below that has at least enough individuals to attain the minimum weight
(50 g). Include as many individuals as possible to attain a maximum sample weight of 400 g.
PRIMARY TARGET SPECIES (small adult fish)*
1) The most common minnow spe-
cies in the region (e.g., blacknose
dace)
2) Another dace species
3) Another common minnow (e.g.,
creek chub or fallfish)
4) The most common sculpin spe-
cies in the region (e.g., Slimy scul-
pin or mottled sculpin)
5) Another common minnow species (e.g.,
stoneroller)
6) A darter species
7) A shiner species
8) If less than the desired weight of any pri-
mary target species is collected, send individu-
als of a small nontarget species if 50 g or more
are available.
* The smallest individual in the sample should be at least 75% of the length of the largest individual. If there is a
conflict between criteria, getting a sufficient sample is a higher priority than getting similar-sized individuals.
2. Prepare a clean work surface to prepare the primary composite sample. Keep hands, work
surfaces, and wrapping materials clean and free of potential contaminants (mud, fuel, formalin,
sun screen, insect repellant, etc.)
3. Rinse the teflon weighing beaker (to be used ONLY for weighing fish) with deionized water or
stream water. Line the beaker with a sufficiently large piece of aluminum foil. Place the dull
side of the foil toward the inside so it will be in contact with the fish. Place the beaker with foil
on the scale and tare it.
4. If not done previously during the preparation of voucher specimens, place the individuals for
the primary composite sample (Step 1) into a bucket with two carbon dioxide tablets (e.g.,
"Alka Seltzer®") and a small volume of water. After the individuals have been anaesthetized,
use clean hands to transfer them into the beaker with foil.
5. Measure the total weight to the nearest 5g. Record the common name (from a standardized
list) of the primary target species, its species code (if required), and the number of individuals
in the sample in the appropriate fields on line "P1" in the primary tissue sample section of the
Sample Collection Form. Enter an "F" series flag in the "Flag" field. Record the total weight of
the sample in the comment/flag explanation section of the Sample Collection Form (if neces-
sary).
(continued)
185
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EMAP-SW-Streams Field Operations Manual, Section 13 (Fish Tissue Contaminants), Rev. 4, Sept. 1998 Page 4 of 9
TABLE 13-1 (Continued)
6. If the individuals included in the composite sample were collected from throughout the sam-
pling reach, place an "X" in the "YES" box in the primary sample section of the Sample Collec-
tion Form. If the individuals were only collected from a limited segment of the sampling reach,
place an "X" in the "No" box and explain in the "EXPLAIN" field on the form.
7. Wrap the fish in the aluminum foil from the beaker. Make sure the dull side of the aluminum
foil is in contact with the fish.
8. Place the sample in a self-sealing plastic bag. Expel excess air and seal the bag(s). Wrap
clear tape around the bag to seal and make a surface for each sample label.
9. Prepare two Fish Tissue sample labels (each having the same sample ID number) by filling in
the stream ID and the date of collection. Circle "PRIMARY" on each label. Record the sample
ID number (barcode) in the primary sample section of the Sample Collection Form. Attach one
label to the tape surface of the bag. Cover the label with a strip of clear tape.
10. Place the labeled bag into a second self-sealing plastic bag. Seal and attach the second label
to the outside of the bag. Cover the label with a strip of clear tape.
11. Place the double-bagged sample into a portable freezer, into a container with dry ice, or into a
cooler containing bags of ice until shipment. Keep the sample frozen until shipment.
186
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EMAP-SW-Streams Field Operations Manual, Section 13 (Fish Tissue Contaminants), Rev. 4, Sept. 1998 Page 5 of 9
TABLE 13-2. PROCEDURE TO PREPARE THE SECONDARY COMPOSITE SAMPLE FOR
FISH TISSUE CONTAMINANTS
NOTE: If neither a primary nor secondary species sample is available, use your best judge-
ment in sending some type of composite fish tissue sample.
1. After all voucher specimens have been prepared, select the highest priority secondary target
species from the list below that has at least 5 individuals of the desired size (>120 mm) is
available. Include similar sized individuals if available.
SECONDARY TARGET SPECIES (Larger adult fish)
1) A regionally common bottom
feeder (e.g., white sucker)
2) Another regionally common bot-
tom feeder (e.g., hogsucker)
3) A regionally common piscivore
(e.g., a bass species)
4) Another regionally common
piscivore (e.g., a trout species)
7) If fewer than 5 individuals of the desired
size are collected for any target species, select
a species having 5 individuals, even if some
individuals are smaller than the desired size.
8) If fewer than 5 individuals of any size are
available for any target species, prepare a
composite sample using as few as 3 fish that
are at least at or near the minimum desired
size (120 mm).
5) Another regionally common
piscivore (e.g., a sunfish species)
6) Carp
9) If an acceptable secondary target species
sample (by the above criteria) is not available,
send only the primary target species sample.
The smallest individual in the sample should be at least 75% of the length of the largest individual. If there is a
conflict between criteria, getting a sufficient sample is a higher priority than getting similar-sized individuals.
2. Prepare a clean work surface to prepare the secondary composite sample. Keep hands, work
surfaces, and wrapping materials clean and free of potential contaminants (mud, fuel, formalin,
sun screen, insect repellant, etc.)
3. Measure the total length (TL) of each individual. Record the common name (from a standard-
ized list) of the secondary target species, its species code (if required), and the total length for
each individual on lines S1 through S5 in the secondary sample section of the Sample Collec-
tion Form.
4. If the individuals included in the composite sample were collected from throughout the sam-
pling reach, place an "X" in the "YES" box in the secondary sample section of the Sample
Collection Form. If the individuals were only collected from a limited segment of the sampling
reach, place an "X" in the "No" box and explain in the "EXPLAIN" field on the form.
5. Wrap each individual separately in aluminum foil, with the dull side of the foil in contact with
the fish. Place all the wrapped individuals into a single self-sealing plastic bag.
(continued)
187
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EMAP-SW-Streams Field Operations Manual, Section 13 (Fish Tissue Contaminants), Rev. 4, Sept. 1998 Page 6 of 9
TABLE 13-2 (Continued)
6. Expel excess air and seal the bag. Wrap clear tape around the bag to seal and make a sur-
face for the sample label.
7. Prepare two Fish Tissue sample labels (each having the same sample ID number) by filling in
the stream ID and the date of collection. Circle "SECONDARY" on each label. Record the
sample ID number (barcode) in the secondary sample section of the Sample Collection Form.
Attach one label to the tape surface of the bag. Cover the label with a strip of clear tape.
8. Place the labeled bag into a second self-sealing plastic bag. Seal and attach the second label
to the outside of the bag. Cover the label with a strip of clear tape.
9. Place the double-bagged sample into a portable freezer, a container with dry ice, or a cooler
containing ice bags until shipment. Keep the sample frozen until shipment.
188
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EMAP-SW-Streams Field Operations Manual. Section 13 (Fish Tissue Contaminants). Rev. 4. Sept. 1998 Page 7 of 9
FISH TISSUE
SITE ID: MAI A JL_2_ - JL JL 3_
DATE: 7//T/98
229013
FISH TISSUE
SITE ID: MAIA J? JJ_ -_?.£. 2_
DATE: 11 Ifl
SAMPLE: PRIMARY /SECONDARY
229014
Figure 13-1. Completed sample labels for fish tissue contaminants.
frozen until they can be shipped (Section 3). Tissue samples can be stored and shipped
with other samples requiring freezing ( periphyton chlorophyll, periphyton biomass, peri-
phyton APA, and sediment metabolism samples). If shipping on dry ice, special containers
and shipping forms will be required.
13.2 EQUIPMENT AND SUPPLIES
Figure 13-3 is a checklist of equipment and supplies required to conduct protocols
described in this section. This checklist may differ from the checklists presented in Appen-
dix A, which are used at a base site to ensure that all equipment and supplies are brought
to and are available at the stream site. Field teams are required to use the checklist pre-
sented in this section to ensure that equipment and supplies are organized and available to
conduct the protocols efficiently.
189
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EMAP-SW-Streams Field Operations Manual, Section 13 (Fish Tissue Contaminants), Rev. 4, Sept. 1998 Page 8 of 9
Reviewed by (initial):
SAMPLE COLLECTION FORM - STREAMS (continued)
SITE NAME: /H/tt Cueeif. DATE: 7//J"/97 VISIT: H1 D2
SITE ID: MA A9 7-_jtJL_2- TEAM ID (X): Hi D2 D3 D4 D5 D6 D7 D8
CHEMISTRY AND MICROBIAL WATER SAMPLE (Chem: 4-L Cubitainer and 2 Syringes
SAMPLE ID (BARCODE) TRANSECT FLAG COMMENTS
CHEMISTRY
JL A T° c o
_£.jLjrr<2. c o
£_AH^_ C 0
2 2._?L o _!__&
COMMON NAME TOT
ic/A»^e" fucker
UJlti^e fucker
UJ A ('/« sue. k e r*
k/Ai'/e fffefee*-
UJ A iT e. StfC ker>
IS COMPOSITE SAMPLE COMPOSED OF INDIVIDUALS COLLECTED FROM THROUGHOUT REACH? (X) -
AL LENGTH (MM) FLAG
IZ&
I3.f
/3V
UK
/*r*
BYES DNo
IF No, EXPLAIN:
LINE
PI
COMMENT OR FLAG EXPLANATION FOR FISH TISSUE
F\ ~ it Ittftlviatt^tr tvii'tlit*! SO a.
J J
Flag code*: KsSxnpIa not collected; U= Suspect samplo; F1, F2, otc.= misc. flag assigned by field crew. Explain all flags In Comments sections.
R«V. 06/02/97 (8<_s«o.97)
SAMPLE COLLECTION FORM - STREAMS - 2
Figure 13-2. Sample Collection Form , showing information recorded for fish tissue samples.
190
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EMAP-SW-Streams Field Operations Manual, Section 13 (Fish Tissue Contaminants), Rev. 4, Sept. 1998 Page 9 of 9
EQUIPMENT AND SUPPLIES FOR FISH TISSUE CONTAMINANTS
QTY.
1
4
1 roll
1
1
'1 roll
4
1
2 sets
1 pkg.
1
1 copy
1 set
Item
Plastic bucket for anesthetization
Carbon dioxide tablets (Alka-Seltzer® or equivalent)
Clear tape for sealing tissue sample bags
Teflon beaker for weighing primary tissue sample
Portable scale, precision ±5g
Aluminum foil
1 -gallon self-sealing plastic bags
Sample Collection Form
Fish tissue sample labels (each set with a different sample ID number [bar-
code])
Clear tape strips
Soft (#2) lead pencils to record data
Fine-point indelible markers to fill out labels
Portable freezer, OR container with dry ice, OR cooler with ice (double-bagged
and taped)
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for fish
tissue contaminants
Figure 13-3. Equipment and supplies checklist for fish tissue contaminants.
191
-------
-------
SECTION 14
RAPID HABITAT AND VISUAL STREAM ASSESSMENTS
by
James M. Lazorchak1, Alan T. Herlihy2, and Jim Green3
After all other samples and field data have been collected, the field team conducts
an overall habitat assessment of the stream, makes a general visual assessment of the
stream, and performs a final check of the data forms and samples before leaving the stream
site (see Section 15). The habitat assessment protocol used is adapted from EPA's "rapid"
bioassessment protocols (Plafkin et al, 1989), and has been refined from various applica-
tions across the country. The approach focuses on integrating information from specific
parameters on the structure of the physical habitat. The objective of the visual stream as-
sessment is to record field team observations of catchment and stream characteristics that are
useful for data validation, future data interpretation, ecological value assessment, development
of associations, and verification of stressor data. The observations and impressions of field
teams are extremely valuable.
14.1 RAPID HABITAT ASSESSMENT
Based on the perception gained from collecting samples and measurements from
throughout the sampling reach, classify the stream as either "Riffle/run" or "Pool/glide"
prevalent based on your visual impression of the dominant habitat type. Choose the preva-
lent habitat type based on which habitat type occupies the majority of the length of the sam-
pling reach. A different field data form is completed depending upon the prevalent habitat
type.
For each prevalent habitat type, twelve characteristics (termed "parameters") of hab-
itat are considered and evaluated as part of the rapid habitat assessment. These parame-
1 U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin Luther King
Dr., Cincinnati, OH 45268.
2
Dept. of Fisheries and Wildlife, Oregon State University, c/o U.S. EPA, 200 SW 35th St., Corvallis, OR 97333.
3 U.S. EPA Region 3, Wheeling Office, 303 Methodist Bldg., 11th and Chaplin Streets, Wheeling, WV 26003.
193
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EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
^_ Rev. 4, September 1998 Page 2 of 17 . .
ters are described in Table 14-1. Most of the parameters are evaluated similarly for both
types of prevalent habitats. In four cases, the same parameter is evaluated differently, or a
different (but ecologically equivalent) parameter is evaluated in riffle/run prevalent versus
pool/glide prevalent streams. Epifaunal substrates are evaluated differently in riffle/run and
pool/glide prevalent streams. Substrate embeddedness is evaluated in riffle/run prevalent
streams, while pool substrate composition is evaluated in pool/glide prevalent streams.
The presence of four potential types of microhabitat types based on combinations of depth
and current velocity is evaluated in riffle/run prevalent streams, while the presence of four
potential types of pool microhabitat based on depth and area are evaluated in pool/glide
prevalent streams. The frequency of riffles is evaluated in riffle/run prevalent streams, while
channel sinuosity is evaluated in pool/glide prevalent streams.
The procedure for conducting the rapid habitat assessment is presented in Table 14-
2. For each of the twelve parameters, rate the overall quality of the sampling reach on a
scale of 0 to 20. For riffle/run prevalent streams, record your scores for each parameter on
the riffle/run version of the Rapid Habitat Assessment Form as shown in Figures 14-1 and
14-2. If the stream is classified as a pool/glide prevalent stream, record your scores for
each parameter on the pool/glide version of the Rapid Habitat Assessment Form as shown
in Figures 14-3 and 14-4. Transfer the scores assigned for each parameter to the box in
the left-hand column of the form. Sum the scores for each parameter and record the total
score in the box at the top of page 1 of the form.
14.2 VISUAL STREAM ASSESSMENT
The objective of the visual stream assessment is to record field crew observations of
catchment/stream characteristics useful for future data interpretation, ecological value as-
sessment, development of associations, and verification of stressor data. Observations and
impressions of field crews are extremely valuable. Thus, it is important that these observa-
tions about stream characteristics be recorded for future data interpretation and validation.
The 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
Comments section of the form.
Complete the assessment form after all other sampling and measurement activities
have been completed. Take into account all observations the sampling team has made
while at the site. The assessment includes the following components: watershed activities
and
194
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EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 3 of 17 •
TABLE 14-1. DESCRIPTIONS OF HABITAT PARAMETERS USED IN THE RAPID
ASSESSMENT OF STREAMS
Habitat
Parameter
Prevalent
Habitat
Type
R=Riffle/run
P=Pool/glide
Description and Rationale
1.
Instream
Cover
(fish)
R
P
Includes the relative quantity and variety of natural structures in the stream (e.g.,
fallen trees, logs, and branches, large rocks, and undercut banks) that are avail-
able for refugia, feeding, or spawning. A wide variety of submerged structures in
the stream provide fish with a large number of niches, thus increasing assemblage
diversity.
2.
Epifaunal
Substrate
(benthic
invertebrates)
R
Essentially the amount of niche space or hard substrates (rocks, snags) available
for insects and snails. Numerous types of insect larvae attach themselves to
rocks, logs, branches, or other submerged substrates. As with fish, the greater
the variety and number of available niches or attachments, the greater the variety
of insects in the stream. Rocky-bottom areas are critical for maintaining a healthy
variety of insects in most high gradient streams.
The abundance, distribution, and quality of substrate and other stable colonizing
surfaces (e.g., old logs, snags, aquatic vegetation) that maximize the potential for
colonization.
3A.
Embeddedness
R
The extent to which rocks (gravel, cobble, and boulders) are covered or sunken
into the silt, sand, or mud of the stream bottom. Generally, as rocks become em-
bedded, the surface area available to macroinvertebrates and fish for shelter,
spawning, and egg incubation is decreased. To estimate the percent of
embeddedness, observe the amount of silt or finer sediments overlying and sur-
rounding the rocks. If kicking does not dislodge the rocks or cobble, they may be
greatly embedded. It is useful to observe the extent of the dark area on their un-
derside of a few rocks.
3B.
Pool
Substrate
Characteriza-
tion
Evaluates the type and condition of bottom substrates found in pools. Firmer sedi-
ment types (e.g., gravel, sand) and rooted aquatic plants support a wider variety of
organisms than a pool substrate dominated by mud or bedrock and no plants. In
addition, a stream that has a uniform substrate in its pools will support far fewer
types of organisms than a stream that has a variety of substrate types.
4A.
Velocity and
Depth Regimes
R
There are four primary current and depth combinations: (1) slow-deep, (2) slow-
shallow, (3) fast-deep, and (4) fast-shallow. The best streams in high gradient
•egions will have all four combinations present. The presence or availability of
iiese four habitats relates to the ability of the stream to provide and maintain a
stable aquatic environment. In general use a depth of 0.5 m to separate shallow
'rom deep and a current velocity of 0.3 m/sec to separate fast from slow.
4B.
Pool Variability
Rates the overall mixture of pool types found in streams, according to size and
depth. The four basic types of pools are large-shallow, large-deep, small-shallow,
and small-deep. A stream with many pool types will support a wide variety of
aquatic species. Rivers with low sinuosity (few bends) and monotonous pool
characteristics do not have sufficient quantities and types of habitat to support a
diverse aquatic community. As a general guideline, consider a pool deep if it is
greater than 1 m deep, and large if its length, width, or oblique dimension is
greater than half the stream width.
(continued)
195
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EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 4 of 17
TABLE 14-1 (Continued)
Habitat
Parameter
Prevalent
Habitat
Type
R=Riffle/run
P=Pool/glide
Description and Rationale
5.
Channel
Alteration
R
P
Basically a measure of large-scale changes in the shape of the stream channel.
Many streams in urban and agricultural areas have been straightened, deepened,
or diverted into concrete channels, often for flood control purposes. Such streams
have far fewer natural habitats for fish, macroinvertebrates, and plants than do
naturally meandering streams. Channel alteration is present when the stream
runs through a concrete channel; when artificial embankments, riprap, and other
forms of artificial bank stabilization or structures are present; when the stream is
very straight for significant distances; when dams and bridges are present; and
when other such changes have occurred.
6.
Sediment
Deposition
R
P
The amount of sediment that has accumulated and the changes that have
occurred to the stream bottom as a result of the deposition. Deposition occurs
from large-scale movement of sediment caused by watershed erosion. Sediment
deposition may cause the formation of islands, point bars (areas of increased
deposition usually at the beginning of meanders that increase in size as the chan-
nel is diverted toward the outer bank) or shoals or result in the filling of pools.
Increased sedimentation also results in increased deposition. Usually this is evi-
dent in areas that are obstructed by natural or man-made debris and areas where
the stream flow decreases, such as bends. High levels of sediment deposition
create an unstable and continually changing environment that becomes unsuitable
for many organisms. _^^__^_
7A.
Frequency of
Riffles
R
The sequence of riffles occurring in a stream. Riffles are a source of high-quality
habitat and diverse fauna, therefore, an increased frequency of occurrence greatly
enhances the diversity of the stream community. For areas where riffles are un-
common, a run/bend ratio can be used as a measure of sinuosity. A large degree
of sinuosity provides for diverse habitat and fauna, and the stream is better able to
handle the high energy flows that result from storms than are relatively straight
streams.
7B.
Channel
Sinuosity
Evaluates the meandering or relative frequency of bencls of the stream. Streams
that meander provide a variety of habitats for aquatic organisms, whereas straight
stream segments are characterized by monotonous habitats that are prone to
flooding. A high degree of sinuosity creates a variety of pools and reduces the
energy from surges when the stream flow fluctuates. The absorption of this en-
ergy by bends protects the stream from excessive erosion and flooding. In "ox-
bow" streams of coastal areas and deltas, meanders are highly exaggerated and
transient. Natural conditions are shifting channels and bends. Alteration of these
streams is usually in the form of flow regulation and diversion. ^^
8.
Channel Flow
Status
R
P
The degree to which the channel is filled with water. The flow status will change
as the channel enlarges or as flow decreases as a result of dams and other ob-
structions, diversions for irrigation, or drought. When water does not cover much
of the streambed, the amount of useable substrate for aquatic organisms is lim-
ited.
(continued)
196
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EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 5 of 17 .
TABLE 14-1 (Continued)
Habitat
Parameter
Prevalent
Habitat
Type
R=Riffle/run
P=Pool/glide
Description and Rationale
9.
Condition of
Banks
R
P
The stream banks are eroded (or have the potential for erosion). Steep banks are
more likely to collapse and suffer from erosion than are gently sloping banks and
are therefore considered to be unstable. Signs of erosion include crumbling,
unvegetated banks, exposed tree roots, and exposed soil
10.
Bank
Vegetative
Protection
R
P
The amount of the stream bank that is covered by vegetation. The root systems
of plants growing on stream banks help hold soil in place, thereby reducing the
amount of erosion that is likely to occur. This parameter supplies information on
the ability of the bank to resist erosion, as well as some additional information on
the uptake of nutrients by the plants, the control on instream scouring, and stream
shading. Banks that have full, natural plant growth are better for fish and
macroinvertebrates than are banks without vegetative protection or those shored
up with concrete or riprap.
11.
Grazing or
Disruptive
Pressure
R
P
Disruptive changes to the riparian zone because of grazing or human interference
(e.g., mowing). |n areas of high grazing pressure from livestock or where residen-
tial and urban development activities disrupt the riparian zone, the growth of a
natural plant community is impeded. Residential developments, urban centers,
golf courses, and rangeland are the common causes of anthropogenic effects on
the riparian zone.
12.
Riparian
Vegetated Zone
Width
R
P
The width of natural vegetation from the edge of the stream bank (riparian buffer
zone). The riparian vegetative zone serves as a buffer zone to pollutants entering
a stream from runoff, controls erosion, and provides stream habitat and nutrient
input into the stream. A relatively undisturbed riparian zone reflects a healthy
stream system; narrow, far less useful riparian zones occur when roads, parking
lots, fields, lawns, bare soil, rocks, or buildings are near the stream bank. The
presence of "old fields" (i.e., a previously developed field allowed to convert to
natural conditions) will rate higher than fields in continuous or periodic use. Paths
and walkways in an otherwise undisturbed riparian zone may be judged to be in-
consequential to destruction of the riparian zone.
197
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EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 6 of 17
TABLE 14-2. PROCEDURE FOR CONDUCTING THE RAPID HABITAT ASSESSMENT
1. Based on observations during previous sample collection and field measurement activities,
classify the sampling reach as predominantly flowing water habitat ("Riffle/run") or slow water
habitat ("Pool/glide").
2. Select the appropriate version of the Rapid Habitat Assessment Form ("Riffle/Run Prevalence"
or "Pool/Glide Prevalence") based on the classification in Step 1.
3. For each of the 12 habitat parameters, determine the general "quality" category ("POOR",
"MARGINAL", "SUB-OPTIMAL", or "OPTIMAL") of the entire sampling reach. Assign and circle a
score from the values available within each quality category. For each parameter, the sam-
pling reach can be scored from 0 (worst) to 20 (best).
4. After the sampling reach has been scored for all parameters, transfer the score circled for each
category to the corresponding "SCORE" box in the "HABITAT PARAMETER" column of the
assessment form.
5. Sum the scores recorded in Step 4 over all 12 habitat parameters. Record the total score for
the sampling reach in the "TOTAL SCORE" box on page 1 of the assessment form. The total
score can range from 0 to 240.
198
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 7 of 17
Reviewed by (initial):
RAPID HABITAT ASSESSMENT FORM: Rl FFLE/RUN PREVALENCE - STREAMS
SITE NAME: MtLL Cftf£F<- DATE: 7 Iff 1ST VISIT: B1 D2
SITE ID: M Al A9 7 -_2_j?__3_ TEAMID(X): 01 D2 D3 D4 D5 D6 D7 D8
TOTAL 1 /£3
SCORE | ' »•*•
HABITAT PARAMETER
1.. INSTREAM COVER
(FISH)
SCORE:I_J^
2. EPtFAUNAL
SUBSTRATE
SCOREcJ &
3. EMBEDDEDNESS
SCORE: | /&
4. VELOCITY/DEPTH
REGIMES
SCORE:) 15"
5. CHANNEL
ALTERATION
SCORE j /f
6. SEDIMENT
DEPOSITION
SCORE: 1 /*/
QPTIM*?
Greater than 50% mix of
boulder, cobble,
submerged logs,
undercut banks, or
other stable habitat
20 19 18 17 16
Well-developed riffle
and run; riffle Is as wfila
as stream and Its length
extends two times tha
width of stream;
abundance of cobble.
20 19 18 17 16
Gravel, cobble, and
boulder particles are
between 0 and 25%
surrounded by fine
sediment
20 19 18 17 16
AH four velocity
regimes are present
(slow-deep, slow-
shallow, fast-deep, fast-
shallow).
20 19 18 17 16
No channelization of
dredging present.
20 19^17 16
Little or no enlargement
of Islands or point bars
and less than 5% of tha
bottom Is affected by
sediment deposition.
20 19 18 17 16
CATEGORY
pira-Qpriuai
30 to 50% mix of boulder,
cobble, or other stable
habitat; adequate habitat
15 14 13 <5D 11
Riffle Is as wide as stream,
but Es [ess than two times
wkfth; abundance of cobble;
boulders and gravel common.
15 14 13 12 11
Gravel, cobble, and boulder
particles are between 25 and
50%sUTOunded by fine
sodhnent
15 14 13 (32)11
Only three of the four 'habitat
types am present (If fast-
shalow is missing, score
loner than If other regimes
aramfes&ig).
»4S)14 13 12 11
SotnachanneHzatlon is
press,-.!, usually In areas of
bridge abutments; evidence
of past ehannetization, Le.,
dred^ng (greater than past
28 yr) Bay be present, but
recant channelization is not
ptiBSflflt
15 14 13 12 11
Soma new Increase In bar
formalion, mostly from
coarse gravel; 5 to 30% of the
bottom to affected; slight
disposition In pools.
15 (^ 13 12 11
M^nifjiii,
10 to 30% mix of boulder,
cobble, or other stable
habitat; habitat availability
Is less than desirable.
10 9 8 7 6
Run area may be lacking;
reduced riffle area that does
not extend across entire
cross section and is less
than two times the width;
gravel or large boulders and
bedrock prevalent; cobble
present
10 9 (8^ 7 6
Gravel, cobble, and boulder
particles are between 50 and
75% surrounded by fine
sediment
10 9 8 7 6
Only two of the four habitat
types are present {if fast-
shallow or slow-shallow are
missing, score low).
10 9 8 7 6
New embankments are
present on both banks; and
40 to 80% of the stream
reach Is channelized and
disrupted.
10 9 8 7 6
Moderate deposition of new
gravel or coarse sand on
old and new bars; 30 to 50%
of the bottom Is affected;
sediment deposits at
obstructions, constrictions,
and bends; moderate
deposition of pools
prevalent
10 9 8 7 6
POOR
Less than 10% of boulder,
cobble, or other stable
habitat; lack of habitat is
obvious.
543210
Riffles or run virtually non-
existent; gravel or large
boulders and bedrock
prevalent; cobble lacking.
543210
Gravel, cobble, and boulder
particles are over 75%
surrounded by fine
sediment
543210
Dominated by one
velocity/depth regime
(usually slow-deep).
543210
Banks shored with gabion or
cement; over 80% of the
stream reach is channelized
and disrupted.
543210
Heavy deposits of fine
material; Increased bar
development; more than 50%
of the bottom Is changing
Frequently; pools almost
absent due to substantial
sediment deposition.
543210
Rev. 06/02/97 <»txxrhrr.97) RAPID HABITAT ASSESSMENT FORM: RIFFLE/RUN - STREAMS -1
Figure 14-1. Rapid Habitat Assessment Form for riffle/run prevalent streams (page 1).
199
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 8 of 17
Reviewed by (initial):
RAPID HABITAT ASSESSMENT FORM: RIFFLE/RUN - STREAMS (continued)
SITE NAME: M\LL CREGK. DATE: 7 //*""/ 97 VISIT: 01 D2
SITE ID: MAIA97- t 5_i_ TEAM ID (X): 01 D2 Q3 D4 D5 D6 D7 D8
HABITAT PARAMETER
7. FREQUENCY OF
RIFFLES
SCORE:! 13
B. CHANNEL FLOW
STATUS
SCORE: 1 /?
9. CONDITION OF BANKS
SCORE: I If
10. BANK VEGETATIVE
PROTECTION
SCORE: \ /*>
11. GRAZING OR OTHER
DISRUPTIVE
PRESSURE
SCORE: I //
12. RIPARIAN
VEGETATION ZONE
WIDTH (LEAST
BUFFERED SIDE)
SCORE: I tO
CATEGORY
OPTtUAl
Oceurrenca of riffles is
relatively frequent; the
distance between riffles
divided by ths width of
the stream equals 5 to 7;
variety of habitat.
20 19 18 17 16
Water reaches the base
of both banks and a
minimal area of channel
substrate Is exposed.
20 19(?§)17 16
Banks stable; no
evidence of erosion or
bank failure.
20 19 18 17 16
More than 90% of tho
stream bank surfaces
era covered by
vegetation.
20 19 18 17f5*D
Vegetative disruption,
through grazing or
mowing Is minimal or
not evident; almost all
plants are allowed to
grow naturally.
20 19 18 17 16
Width of riparian zone is
greater than 18m;
human activities (I.e.;
parking lots, roadbeds,
ctearcuts, lawns, or
crops) have not
Impacted this zone.
20 19 18 17 16
^IJR-OPTIMAI
Occurrence of riffles Is
Infrequent; distance between
riffles divided by the width of
tho stream equals 7 to 15.
15 14 /f3) 12 11
Water fills more than 75% of
the available channel; or less
than 25% of the channel
substrate Is exposed.
15 14 13 12 11
Banks moderately stable;
Infrequent, small areas of
erosion mostly healed over.
tfS)l4 13 12 11
70 to 90% of the stream bank
surfaces are covered by
vegetation.
15 14 13 12 11
Disruption Is evident but Is
not affecting full plant growth
potential to any great extent;
more than one-hall of the
potential plant stubble height
remaining.
15 14 13 12 (TD
Zone width Is between 12 and
18m; human activities have
only minimally impacted this
zone.
15 14 13 12 11
P/f^Rf^MAI
Occasional riffle or bend;
bottom contours provide
some habitat; distance
between riffles divided by
the width of the stream Is
between 15 to 25.
10 9 8 7 6
Water fill 25 to 75% of the
available channel; and/or
riffle substrates are mostly
exposed.
10 9 8 7 6
Moderately unstable; up to
60% of banks In reach have
areas of erosion.
10 9 8 7 6
50 to 70% of the stream
bank surfaces are covered
by vegetation.
10 9 8 7 6
Disruption Is obvious;
patches of bare soil or
closely cropped vegetation
are common; less than one-
half of the potential plant
stubble height remaining.
10 9 8 7 6
Zone width Is between 6 and
12 m; human activities have
Impacted the zone a great
deaL
(lo) 9876
Pnnp
Generally all flat water or
shallow riffles; poor habitat;
distance between riffles
divided by the width of the
stream Is greater than 25.
543210
Very little water In channel,
and mostly present as
standing pools.
543210
Unstable; many eroded
areas; "raw" areas frequent
along straight sections and
bends; on side slopes, 60 to
100% of bank has eroslonal
scars.
543210
Less than 50% of the stream
bank surfaces are covered
by vegetation.
543210
Disruption of stream bank
vegetation Is very high;
vegetation has been
removed to 2 Inches or less
In average stubble height
543210
Width of zone Is less than 6
m; little or no riparian
vegetation due to man-
Induced activities.
543210
Rev. oeo2A»7 (start*r.97) RAPID HABITAT ASSESSMENT FORM: RIFFLE/RUN - STREAMS - 2
Figure 14-2. Rapid Habitat Assessment Form for riffle/run prevalent streams (page 2).
200
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4. September 1998 Page 9 of 17
Reviewed by (initial):
RAPID HABITAT ASSESSMENT FORM: GLIDE/POOL PREVALENCE - STREAMS
SITE NAME: ty t I.L CKee< DATE: 7 / /f/ 97 VISIT: 01 D2
SITEID: M Al A97-_2.JL_2_ TEAMID(X): 01 D2 Q3 Q4 Q5 Q6 D7 D8
TOTAL | . ._
SCORE II I'f
HABITAT PARAMETER
1.- INSTREAM COVER
SCORE: § ff
2. EPIFAUNAL
SUBSTRATE
SCORE: \ $
3. POOL SUBSTRATE
CHARACTERIZATION
g>
4. POOL VARIABILITY
SCORE: | £
S. CHANNEL ALTERATION
SCORED j&
6. SEDIMENT
DEPOSITION
*7
OPTIMAL
Greater than 50% mix of
snags, submerged logs,
undercut banks, or other
stable habitat; rubble or
gravel may be present
20 19 18 17 16
Preferred benthlc
substrata (to be sampled)
Is abundant throughout
stream site and at a stags
to allow for full
colonization potential
(I.e.; logs and snags that
are not new fall and not
transient
20 19 18 17 16
Mixture of substrate
materials, with gravel and
firm sand prevalent; root
mats and submerged
vegetation are common.
20 19 18 17 16
Even mix of large-
shallow, large-deep,
small-shallow, and small-
deep pools are present
20 19 18 17 16
No channelization of
dredging present
20 19 18 17 ©
Less than 20% of the
bottom Is affected; minor
accumulation of fine and
oarse material at snags
and submerged
vegetation; little or no
enlargement of Islands or
lolnt bars.
20 19 18 17 16
CATEGORY
SlIB-OPTIM Al
30 to 50% mix of stable
habitat; adequate habitat for
maintenance of populations.
15 14 13 12 11
Substrate is common but is
not prevalent nor well-suited
for full colonization potential.
15 14 13 12 11
Mixture of soft sand, mud, or
clay; mud may be dominant;
some root mats and
submerged vegetation are
H&sent
15 14 13 12 11
The majority of pools are
large and deep; very few
shallow.
15 14 13 12 11
Some channelization Is
jresent, usually in areas of
>rldgo abutments; evidence
of past channelization. I.e.;
Iredglng (greater than past
20 yr) may be present, but
recent channelization Is not
)resent
15 14 13 12 11
20 to 50% affected; moderate
accumulation; substantial
edlmsnt movement only
during major storm events;
ome new Increase In bar
ormation.
15 14 13 .12 11
MARGINAL
1 0 to 30% mix of stable
habitat; habitat availability
Is less than desirable.
10 9 (S} 7 6
Substrate frequently
disturbed or removed.
10 9 (5} 7 6
All mud or clay or sand
bottom; little or no root
mat; no submerged
vegetation.
10 9 (tf) 7 6
Shallow pools much more
prevalent than deep pools.
10 9 (?) 7 6
lew embankments are
present on both banks;
channelization may be
extensive, usually In urban
areas or drainage areas of
agricultural lands; and
more than 80% of the
stream reach Is
channelized or disrupted.
10 9 8 7 6
50 to 80% affected; major
deposition; pools shallow
and heavily silted;
embankments may be
iresenton both banks;
frequent and substantial
sediment movement during
storm events.
10 9 8 ® 6
PfJPR
Less than 10% stable habitat;
lack of habitat is obvious.
543210
Substrate Is unstable or
lacking.
54321 0
-lard-pan clay or bedrock; no
root mat or vegetation.
543210
Majority of pools are small-
shallow or pools are absent
543210
ixtenstve channelization;
ranks shored with gabion or
cement; heavily urbanized
areas; Instream habitat
greatly altered or removed
entirely.
543210
Channelized; mud, silt
and/or sand In braided or
non-braided channels; pools
almost absent due to
deposition.
543210
Rev. 06WU97 (stxxrhgp.97) RAPID HABITAT ASSESSMENT FORM: GLIDE/POOL - STREAMS -1
Figure 14-3. Rapid Habitat Assessment Form for pool/glide prevalent streams (page 1).
201
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4. September 1998 Page 10 of 17
Reviewed by (initial):
RAPID HABITAT ASSESSMENT FORM: GLIDE/POOL- STREAMS (continued)
SITE NAME: MILL CREfK- DATE: 7 US' / 97 VISIT: ffl D2
SITE ID: M A 1 A 9 7 - ^ ? °( TEAM ID (X): H1 D2 D3 D4 D5 D6 D7 D8
H ABfTAT PAHAMb 1 EH
7. CHANNEL SINUOSITY
SCORE: i / 3
8. CHANNEL FLOW
STATUS
SCORE: | / 9
9. CONDITION OF BANKS
SCORE: 1 <^
10. BANK VEGETATIVE
PROTECTION
SCORE: I J
11. GRAZING OR OTHER
DISRUPTIVE
PRESSURE
SCORE: | §
12. RIPARIAN
VEGETATIOUN ZONE
WIDTH (LEAST
BUFFERED SIDE)
SCORE: 1 7
CATEGORY
OPTlUfll
The bends In the stream
Increase the stream
length 3 to 4 times
longer than If It was In a
straight line.
20 19 18 17 16
Water reaches the basa
of both lower banks and
a minimal amount of
channel substrate Is
exposed.
20 19@17 16
Banks stable; no
evidence of erosion or
bank failure.
20 19 18 17 16
Over 90% of the stream
bank surfaces Is covered
by vegetation.
20 19 18 17 16
Vegetative disruption
minimal or not evident;
almost all plants are
allowed to grow
naturally.
20 19 18 17 16
Width of riparian zone Is
greater than 18 meters;
human activities (I.e.;
parking lots, roadbeds,
dearcuts, lawns, or
crops) have not
Impacted this zone.
20 19 18 17 16
^IIR-OPTIMfll
The bends In the stream
Increase the stream length 2
to 3 times long or than If It was
In a straight line.
15 14 (13)12 11
Water fills more than 75% of
the available channel; or less
than 25% of the channel
substrate is exposed.
15 14 13 12 11
Banks moderately stable;
Infrequent, small areas of
erosion mostly healed over.
15 14 13 12 11
70 to 90% of the stream bank
surfaces Is covered by
vegetation.
15 14 13 12 11
Disruption is evident but Is
not affecting full plant growth
potential to any great extent;
more than one^ialf of the
potential plant stubble height
remaining.
15 14 13 12 11
Width of riparian zone Is
between 12 and 18 meters;
human activities have only
minimally Impacted this zone.
15 14 13 12 11
MAPGINAI
The bends In the stream
increase the stream length
between 1 and 2 times
longer than If It was In a
straight line.
10 9 8 7 6
Water fills 25 to 75% of the
available channel and/or
riffle substrates are mostly
exposed.
10 9 8 7 6
Moderately unstable; up to
60% of banks In reach have
areas of erosion.
10 (£) 876
50 to 70% of the stream
bank surfaces Is covered
by vegetation.
10 9 ^) 7 6
Disruption Is obvious;
patches of bare soli or
closely cropped vegetation
are common; less than one-
half of the potential plant
stubble height remaining.
10 9 (T) 7 6
Width of riparian zone is
between 6 and 12 meters;
human activities have
Impacted the zone a great
deal.
10 9 8 (?) 6
Pnnn
Channel is straight;
waterway has been
channelized for a long
distance.
543210
Very little water In channel,
and mostly present as
standing pools.
543210
Unstable; many eroded
areas; "raw" areas frequent
along straight sections and
bends; side slopes 60 to
100% of bank has eroslonal
scars.
543210
Less than 50% of the stream
bank surfaces are covered
by vegetation.
543210
Disruption of stream bank
vegetation Is very high;
vegetation has been
removed to 2 Inches or less
In average stubble height
543210
Width of riparian zone is less
than 6 meters; little or no
riparian vegetation due to
human activities.
543210
Rsv. 06/02/97 (stxxrhgp.97) RAPID HABITAT ASSESSMENT FORM: GLIDE/POOL - STREAMS - 2
Figure 14-4. Rapid Habitat Assessment Form for glide/pool prevalent streams (page 2).
202
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 11 of 17
observed disturbances, reach characteristics, waterbody character, general assessment,
and local anecdotal information. The procedure for conducting the visual assessment of the
sampling reach is presented in Table 14-3. Record data and observations for each compo-
nent of the assessment on the Assessment Form as shown in Figures 14-5 and 14-6.
Each watershed activity or disturbance is rated into one of four categories of abun-
dance or influence: not observed, low, medium, or high. Leave the line blank for any activity
or disturbance type not observed. The distinction between low, medium, and high will be
subjective. For example, if there are 2-3 houses on a stream, the rating for "Houses" would
be low. If the stream is in a suburban housing development, rate it as high. Similarly, a
small patch of clear cut logging on a hill overlooking the stream would be rated as low. Log-
ging activity right on the stream shore, however, would be rated as high.
When assessing reach characteristics, make your best estimate as to the percent of
the sampling reach (40 channel widths) that had each type of listed riparian zone land use
immediately adjacent to the stream. Also rate the water clarity, including whether you be-
lieve the clarity is influenced by recent storm events (see Section 4).
Water body character is defined as "the physical habitat integrity of the water body,
largely a function of riparian and littoral habitat structure, volume change, trash, turbidity,
slicks, scums, color, and odor." Water body character is assessed using two attributes, the
degree of human development, and aesthetics. Rate each of these attributes on a scale of
1 to 5. For development, give the stream a "5" rating if it is pristine, with no signs of any
human development. A rating of "1" indicates a stream which is totally developed (e.g., the
entire stream is lined with houses, or the riparian zone has been removed). For aesthetics,
base your decision on any factor about the stream that bothers you (e.g., trash, algal
growth, weed abundance, overcrowding).
The general assessment component includes any observations that will help in data
interpretation in the pertinent section. General assessment comments can include com-
ments on wildlife observed, diversity of terrestrial vegetation, age class of forest, or any
other observation. Comments from locals are often useful and should be recorded in the
"LOCAL ANECDOTAL INFORMATION" section. The back side of the form (Figure 14-6) is avail-
able for general comments.
203
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4. September 1998 Page 12 of 17
3.
4.
5.
TABLE 14-3. PROCEDURE FOR CONDUCTING THE FINAL VISUAL ASSESSMENT
OF A STREAM
After all other sampling and measurement activities are completed, fill out the header section
of an Assessment Form. Use your perceptions obtained during the course of the day, while at
the stream or driving/walking through the catchment to complete the remainder of the form.
WATERSHED ACTIVITIES AND DISTURBANCES OBSERVED: Rate each type of activity or disturbance
listed on the form as either "Not observed", "Low", "Medium", or "High", and record the rating
on the Assessment Form. Keep in mind that ratings will be somewhat subjective and that an
extensive effort to quantify the presence and intensity of each type of stressor is not required.
General categories of activities and types of disturbance are described below:
Residential: The presence of any of the listed disturbances adjacent to or near the
stream.
Recreational: The presence of organized public or private parks, campgrounds, beaches
or other recreation areas around the stream. If there are signs of informal areas of camp-
ing, swimming or boating around the stream (e.g., swimming hole), record them as "prim-
itive" parks, camping.
Agriculture: The presence of cropland, pasture, orchards, poultry, and/or livestock.
Industrial: Any industrial activity (e.g., canning, chemical, pulp), commercial activity
(stores, businesses) or logging/mining activities around the stream or in the catchment.
Describe in more detail in the comments section.
Management: Any evidence of liming activity, water treatment, dredging or
channelization, flow control structures, etc.
Any oddities, or further elaboration should be recorded in the Comments section.
REACH CHARACTERISTICS: For each type of riparian vegetation cover or land use category listed
on the Assessment Form, estimate the proportion of the sampling reach immediately adjacent
to the stream that is affected. Place and "X" in the appropriate extent class box (Rare [< 5%],
Sparse [5 to 25%], Moderate [25 to 75%], and Extensive [> 75%]) on the form.
Classify the overall water clarity within the sampling reach as clear, murky, or highly turbid.
Place an "X" in the appropriate box on the "WATER CLARITY" line of the Assessment Fornx If
you believe that water clarity has been influenced by a recent storm event, also place an "X" in
the "STORM INFLUENCED" box.
WATER BODY CHARACTER: Assign a rating of 1 (highly disturbed) to 5 (pristine) based on your
general impression of the intensity of impact from human disturbance. Place an "X" in the box
next to the assigned rating on the Assessment Form.
(continued)
204
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 13 of 17
TABLE 14-3 (Continued)
5. WATERBODY CHARACTER (CONT.): Assign a rating to the stream based on overall aesthetic
quality, based on your opinion of how suitable the stream water is for recreation and aesthetic
enjoyment today. Place and "X" in the box next to the assigned rating on the Assessment
Form.
5. Beautiful, could not be any nicer.
4. Very minor aesthetic problems; excellent for swimming, boating, enjoyment.
3. Enjoyment impaired.
2. Level of enjoyment substantially reduced.
1. Enjoyment nearly impossible.
Add any comments you feel might aid data interpretation in the Comments Section.
6. GENERAL ASSESSMENT: record comments on wildlife observed, perceived diversity of terrestrial
vegetation, and the estimated age class of forest (0 to 25 yr, 25 to 75 yr, or > 75 yr.) on the
Assessment Form.
7. LOCAL ANECDOTAL INFORMATION: Record any information regarding the past or present
characteristics or condition of the stream provided by local residents.
205
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 14 of 17
Reviewed by (initial):
ASSESSMENT FORM - STREAMS/RIVERS
SUE NAME: M//.L
DATE: 7 //TV 97 VISIT: 01 D2
: in- M A I A 9 7 - *?
TEAM ID (X): H1 O2 D3 D4 D5 D6 D7 D8
WATERSHED ACTIVmES AND DISTURBANCES OBSERVED (INTENSITY: BIAMK=NOT OBSERVED, L=Low, M^MOOERATE, H=HEAVY)
RESIDENTIAL
RECREATIONAL
AGRICULTURAL
UVEHTOCKUSE
INDUSTRIAL
l«"nn««LPL"ire
HMES/QIURRIES
OHAjtsWELlS
POWEBPLWT*
STREAM MANAGEMENT
DRMQNQ WATER TREATMENT
ANQLMQ PRESSURE
OREDOIHCi
HBK1ATION PUMPS
W»Tm Lewi. FtuerixTTOls
REACH CHARACTERISTICS (percent of reach)
FOREST
SHRUB
GRASS
WETLAND
BARE GROUND
MACROPHYTES
MoDBIATC(25TO75%) Q EXTP»«(>75%)
[]]R»1H(<5%)
E»rB««(>75%)
Q ExmUNt(>76%)
AGRICULTURE • Row CHOP
AGRICULTURE- GRAZING
LOGGING
DEVELOPMENT (RESIDENTIAL & URBAN)
g]RAM( 75%)
ExTEMM(>75%)
(57025%) QMODEIIATE(aST075%) Q EXTENMVI (> 75%)
gJRAM(<5%)
Q ExrEMVI (> 75%)
WATER CLARITY
| | HtOH.YTWIBK)
WATERBODY CHARACTER (XONE)
PRISTINE
D5
D4
D =
IS*
D1
HIGHLY DISTURBED
APPEALING
UNAPPEALING
GENERAL ASSESSMENT (wildlife, vegetation diversity, forest age class (0-25 yrs. 25-75 yrs, >75)
Lt
JLel* *t t+Hle.
LOCAL ANECDOTAL INFORMATION: I Lot*
•'* <*rf* } m
R»v.OS/D2/B7 (ttrvasso.97)
ASSESSMENT FORM - STREAMS/RIVERS -1
Figure 14-5. Assessment Form (page 1).
206
-------
EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 15 of 17
Reviewed by (initial):.
ASSESSMENT FORM • STREAMS/RIVERS (continued)
SITE NAME:
DATE: / /97 VISIT: D1 D2
SITE ID: MAIA97-.
TEAM ID 00: D1 CI2 D3 d4 I~l5 I~IS I~I7 Ha
Rev. 06/02/97 (strvasse.97)
Figure 14-6. Assessment Form (page 2).
ASSESSMENT FORM - STREAMS/RIVERS - 2
207
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EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4. September 1998 Page 16 of 17
14.3 EQUIPMENT AND SUPPLIES
Figure 14-7 is a checklist of the supplies required to complete the rapid habitat and visual
stream assessments. This checklist may differ from the checklists presented in Appendix
A, which are used at a base site to ensure that all equipment and supplies are brought to
and are available at the stream site. Field teams are required to use the checklist pre-
sented in this section to ensure that equipment and supplies are organized and available to
conduct the protocols efficiently.
14.4 LITERATURE CITED
Plafkin, J.L., M.T. Barbour, K.D. Porter, S.K. Gross, and R.M. Hughes. 1989. Rapid Bio-
assessment Protocols for Use in Streams and Rivers: Benthic Macroinvertebrates and
Fish. EPA/440/4-89/001. U.S. Environmental Protection Agency, Washington, D.C.
208
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EMAP-SW-Streams Field Operations Manual, Section 14 (Rapid Habitat and Visual Stream Assessments),
Rev. 4, September 1998 Page 17 of 17
EQUIPMENT AND SUPPLIES FOR RAPID HABITAT AND VISUAL STREAM ASSESSMENTS
QTY.
1
1
1
6
1
1 copy
1 set
Item
Rapid Habitat Assessment Form for Riffle/run prevalent streams
Rapid Habitat Assessment Form for Pool/glide prevalent streams
Assessment Form for visual stream assessment
Soft (#2) lead pencils
Covered clipboard or forms holder
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides for rapid
habitat and visual assessments
Figure 14-7. Checklist of equipment and supplies required for rapid habitat and visual stream
assessments.
209
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SECTION 15
FINAL SITE ACTIVITIES
by
James M. Lazorchak1
Before leaving a stream site, the team leader reviews all of the data forms and
sample labels for accuracy, completeness, and legibility. A second team member inspects
all sample containers and packages them in preparation for transport, storage, or shipment.
Refer to Section 3 for details on preparing and shipping samples.
When reviewing field data forms, ensure that all required data forms for the stream
have been completed. Confirm that the stream identification code, the year, the visit num-
ber, and the date of the visit are correct on all forms. On each form, verify that all informa-
tion has been recorded accurately, the recorded information is legible, and any flags are
explained in the comments section. Ensure that written comments are legible and use no
"shorthand" or abbreviations. 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.
When inspecting samples, ensure that each sample is labeled, all labels are com-
pletely filled in and legible, and each label is covered with clear plastic tape. Compare
sample label information with the information recorded on the corresponding field data
forms (e.g., the Sample Collection Form) to ensure accuracy.
The other team members should return all of the equipment and supplies to the
vehicle for transport and clean up the stream site. Pack all equipment and supplies in the
vehicle for transport. Keep them organized so they can be inventoried using the equipment
and supply checklists presented in Appendix A. Clean up and dispose of all waste material
at the stream site. Transport it out of the area if necessary.
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin Luther King
Dr., Cincinnati, OH 45268.
211
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APPENDIX A
EQUIPMENT AND SUPPLY CHECKLISTS
FIELD DATA FORMS AND SAMPLE LABELS . A-2
OFFICE SUPPLIES AND TOOLS A-3
PERSONAL EQUIPMENT AND SUPPLIES . : A-3
CHEMICALS A-4
PACKING AND SHIPPING SUPPLIES . . A-4
SITE VERIFICATION AND SAMPLING REACH LAYOUT . . .A-5
WATER CHEMISTRY . . .....' A-5
STREAM DISCHARGE A-6
PHYSICAL HABITAT A-6
PERIPHYTON , A-7
SEDIMENT METABOLISM AND SEDIMENT TOXICITY A-8
BENTHIC MACROINVERTEBRATES A-9
AQUATIC VERTEBRATES AND FISH TISSUE CONTAMINANTS A-10
A-1
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EMAP-SW-Streams Field Operations Manual. Appendix A, Rev. 0, September 1998 Page 2 of 10
FIELD DATA FORMS AND SAMPLE LABELS
Number
per site
Item
1
1
11+ extras
1
+ extras
extras
Verification Form
Sample Collection Form
Field Measurement Form
Channel/Riparian Cross-section and Thalweg Profile Forms
Slope and Bearing Form
Vertebrate Collection Form
Vertebrate Length Recording Form
Rapid Habitat Assessment Form for Riffle/run prevalent streams
Rapid Habitat Assessment Form for Pool/glide prevalent streams^
Assessment Form for visual stream assessment
Water chemistry labels (same ID number)
Periphyton labels (same ID number)
Sediment metabolism labels (different ID numbers)
Sediment toxicity labels
Composite Benthic sample labels, with preprinted ID numbers (barcodes)
Composite Benthic sample labels without preprinted ID numbers
Blank benthic sample labels on waterproof paper for inside of jars
2 sets
Sheet of pre-printed aquatic vertebrate jar labels (4) and voucher bag tags
(36), all with same preprinted sample ID number (barcode)
Fish tissue sample labels (2 labels per set; each set with a different sample ID
number [barcode])
2 copies
2 sets
Field operations and methods manual
Laminated sheets of procedure tables and/or quick reference guides
A-2
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EMAP-SW-Streams Field Operations Manual, Appendix A. Rev. 0, September 1998 Page 3 of 10
OFFICE SUPPLIES AND TOOLS
Number
per site
1
1
1
1
1
4
1
12
6
2
1pr
1
1
1
Item
Dossier of access information for scheduled stream site
Topographic map with "X-site" marked
Site information sheet with map coordinates and elevation of X-site
Sampling itinerary form or notebook
Safety log and/or personal safety information for each team member
Covered clipboards or forms holders
Field notebook (optional)
Soft (#2) lead pencils
Fine-tip indelible markers
Grease pencils
Scissors for cutting labels
Pocket knife or multipurpose tool
Battery charger (if needed for electrofishing unit)
Toolbox with basic tools needed to maintain/repair sampling gear
PERSONAL EQUIPMENT AND SUPPLIES
Number
per site
1 pair per
person
1 per person
3 pair
1
1 per person
1 or2
Item
Chest waders with felt-soled boots for safety and speed if waders are the
neoprene "stocking" type. Hip waders can be used in shallower streams
(except for electrofishing).
Life vests
Polarized sunglasses
First aid kit
Rain gear
Fisherman's vest for physical habitat characterization.
A-3
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EMAP-SW-Streams Field Operations Manual, Appendix A, Rev. 0, September 1998 Paqe 4 of 10
CHEMICALS
Number
1pr
2pr
1
1
2 gal
1
2 gal
Item
Safety glasses
Chemical-resistant gloves
Laboratory apron, resistant to ethanol and formalin
Cooler (with suitable absorbent material) for transporting ethanol and samples
95% ethanol
Cooler (with suitable absorbent material) for transporting
formaldehyde/formalin
10% (buffered) formalin solution OR 0.2 gal buffered formaldehyde solution
Gasoline for electrofishing unit in approved container
PACKING AND SHIPPING SUPPLIES f
Number
per site
1 box
1-box
1 box
1 roll
2pkg.
4 rolls
3
1
2
6
Item
Ice (also dry ice if it is used to ship frozen samples)
1-gal heavy-duty self-sealing (e.g., with a zipper-type closure) plastic bags
30-gal plastic garbage bags
Heavv-dutv clastic bags for sediment toxicity samples
Clear tape for sealing tissue sample bags
Clear tape strips for covering labels
Plastic electrical tape
Insulated shipping containers for samples
Portable freezer, cooler with dry ice, or cooler with bags of ice to store frozen
samples (special containers may be needed if dry ice is used)
Containers and absorbent material suitable to transport and/or ship samples
preserved in formalin or ethanol
Shipping airbills and adhesive plastic sleeves
A-4
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EMAP-SW-Streams Field Operations Manual, Appendix A, Rev. 0, September 1998 Page 5 of 10
SITE VERIFICATION AND SAMPLING REACH LAYOUT
Number
per site
1
1
1
2 rolls
1
Item
GPS receiver and operating manual
Extra batteries for GPS receiver
Surveyor's telescoping leveling rod (round profile, metric scale, 7.5 m
extended)
50-m fiberglass measuring tape with reel
Surveyor's flagging tape (2 colors)
Waterproof camera and film
WATER CHEMISTRY
Number
per site
1
1
1
1
1
1
1
1
1
2
1
2
Item
Dissolved oxygen/Temperature meter with probe and operating manual
DO repair kit containing additional membranes and probe filling solution
Conductivity meter with probe and operating manual
Extra batteries for dissolved oxygen and conductivity meters
500-mL plastic bottle of conductivity QCCS labeled "Rinse" (in plastic bag)
500-mL plastic bottle of conductivity QCCS labeled "Test" (in plastic bag)
500-mL plastic bottle of deionized water to store conductivity probe
Field thermometer
500 mL plastic beaker with handle (in clean plastic bag)
4-L cubitainer
60 mL plastic syringes
Plastic container with snap-on lid to hold filled syringes
Syringe valves
A-5
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EMAP-SW-Streams Field Operations Manual, Appendix A. Rev. 0, September 1998 Page 6 of 10
STREAM DISCHARGE
Number
per site
1
1
1
1
1
1
1
Item
Current velocity meter and probe, with operating manual (e.g.
Model 201 , Swoffer Model 21 00, or equivalent)
Top-set wading rod (metric scale) for use with current velocity
Marsh-McBirney
meter
Portable Weir with 60° "V" notch (optional)
Plastic sheeting to use with weir
Plastic bucket (or similar container) with volume graduations
Stopwatch
Neutrally buoyant object (e.g., orange, small rubber ball, stick)
PHYSICAL HABITAT
Number
jper site
1
1
1
1
2
1
1
1
Item
Fisherman's vest with lots of pockets and snap fittings.
Hip chain (metric) for measuring reach lengths (Optional)
Clinometer (or Abney level) with percent and degree scales.
Lightweight telescoping camera tripod, (necessary only if slope measurements
are being determined by only one person
Vt-inch diameter PVC pipe, 2-3 m long, each marked at the same height (for
use in slope determinations involving two persons)
Spherical convex canopy densiometer, modified with taped "V"
Bearing compass (Backpacking type)
Meter stick. Alternatively, a short (1-2 m) rod or pole (e.g., a ski pole) with cm
markings for thalweg measurements
A-6
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EMAP-SW-Streams Field Operations Manual, Appendix A, Rev. 0, September 1998 Page 7 of 10
PERIPHYTON
Number
per site
1
1
1
1
1
2
1
4
1 box
1 pair
1
1
1
2
2
1
Item
Large funnel (15-20 cm diameter)
12-cm2 area delimiter (3.8 cm diameter pipe, 3 cm tall)
Stiff-bristle Toothbrush with handle bent at 90° angle
1 -L wash bottle for stream water
1-L wash bottle containing deionized water
500-mL plastic bottles for composite index samples, labeled "EROSIONAL"
and "DEPOSITIONAL"
60 mL plastic syringe with a 3/8" hole bored into the end
50-mL screw-top centrifuge tubes (or similar sample vials)
Glass-fiber filters for chlorophyll sample
Forceps for filter handling.
25-mL or 50-mL graduated cylinder
Filtration unit, including filter funnel, cap, filter holder, and receiving chamber
Hand-operated vacuum pump with length of flexible plastic tubing
Pre-leached, pre-ashed, weighed glass-fiber filters in numbered containers for
biomass sample
Aluminum foil squares (3" x 6")
Small syringe or bulb pipette for dispensing formalin
A-7
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EMAP-SW-Streams Field Operations Manual, Appendix A, Rev. 0. September 1998 Page 8 of 10
SEDIMENT METABOLISM AND SEDIMENT TOXICITY
Number
per site
1
1
1
1 set
1
5
1
1
1
1
Item
Small scoop sampler for sediments
Wide-mouthed plastic jar labeled "COMPOSITE SEDIMENT SAMPLE". If
sediment is only being collected for metabolism samples, use a 250-mL jar is
sufficient, if metabolism and toxicity samples are being prepared, use a 1-
gallon jar
YSI Model 58 Dissolved Oxygen meter with Model 5730 Stirring BOD probe
and operating manual
Spare batteries for DO meter
Small plastic spoon or spatula to transfer sediment from the composite
sample container to respiration tubes
50-mL, screw-top, centrifuge tubes
50-ML screw-cap centrifuge tube labeled "BLANK"
Small cooler used as incubation chamber
1 000-mL plastic beaker to holding centrifuge tubes during incubation
Plastic container with snap-on lid to hold the sediment toxicity sample
A-8
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EMAP-SW-Streams Field Operations Manual, Appendix A, Rev. 0, September 1998 Page 9 of 10
BENTHIC MACROINVERTEBRATES
Number
per site
1
2
1
1
2pr.
1
1
4 to 6 ea.
1
1 pkg.
Item
Modified kick net ( closed bag with 595/600 urn mesh) and 4-ft handle (Wildco
#425-C50)
Spare net(s) for the kick net sampler or extra sampler
Buckets, plastic, 8- to 10-qt capacity, labeled "RIFFLE" and "POOL"
Sieve, U.S. Standard #30 mesh
Sieve bucket, 595-um mesh openings
Watchmakers' forceps
Small spatula, spoon, or scoop to transfer sample
Funnel, with large bore spout
Sample jars, plastic with screw caps, 1/4 and 1 gallon capacity, suitable for use
with ethanol
Field checklist sheet
Kim wipes in small self-sealing plastic bag
A-9
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EMAP-SW-Streams Field Operations Manual. Appendix A. Rev. 0, September 1998 Page 10 of 10
Number
1
3pr
2
1
2
2
10
1 set
1
1
1
15-20
1
2ea.
1
4
1
1
1 roll
2 gal
1
Item
Gasoline or battery-powered backpack electrofishing unit with electrode
wand
Extra battery
heavy-duty rubber gloves for electrofishing
Long-handled dip nets (0.6 cm mesh) with insulated handles
Minnow seine (2m * 1.25 m, 0.6 cm mesh) with brailles
Large seines (3 m x 2 m, 0.6 cm mesh) with brailles
larger sized seines for block nets (if necessary)
Collapsible buckets for holding and processing aquatic vertebrates
Taxonomic reference books and keys for fishes and amphibians of the
region
Fish measuring board
List of vertebrate species codes and common names
List of external anomaly codes
Small nylon mesh bags for holding voucher specimens (bags can also be
constructed from sections of nylon stockings or panty hose)
Small fillet knife or scalpel for preparing larger voucher specimens for
preservation
% or 1-gal screw-top plastic jars for voucher samples
Plastic bucket for anesthetization
carbon dioxide tablets (Alka-Seltzer® or equivalent)
Teflon beaker for weighing primary tissue sample
Portable scale, precision ±5g
Aluminum foil
10% (buffered) formalin solution OR 0.2 gal buffered formaldehyde solution
Container to hold preserved voucher sample jars
A-10
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APPENDIX B
QUICK REFERENCE GUIDES
The following pages are tabular summaries of different field activities and
procedures described in this manual. These were developed by the principal investigators
for each ecological indicator to provide a field team with a quick way to access information
about each procedure. They are intended to be laminated and taken to the stream site after
the crew has been formally trained in the detailed procedures as presented in the manual.
They are arranged here in the general sequence of their use in the field.
QUICK REFERENCE GUIDE FOR INITIAL SITE ACTIVITIES B-3
QUICK REFERENCE GUIDE FOR WATER CHEMISTRY B-5
QUICK REFERENCE GUIDE FOR PHYSICAL HABITAT CHARACTERIZATION B-7
QUICK REFERENCE GUIDE FOR PERIPHYTON B-13
QUICK REFERENCE GUIDE FOR SEDIMENT METABOLISM B-15
QUICK REFERENCE GUIDE FOR SEDIMENT TOXICITY B-17
QUICK REFERENCE GUIDE FOR BENTHIC MACRO IN VERTEBRATES , B-19
QUICK REFERENCE GUIDE FOR AQUATIC VERTEBRATES B-25
QUICK REFERENCE GUIDE FOR FISH TISSUE CONTAMINANTS B-29
B-1
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 3 of 31
QUICK REFERENCE GUIDE FOR INITIAL SITE ACTIVITIES
Find the stream location in the field corresponding to the "X" on 7.5" topo map (X-site). Crews
should use all available means to insure that they are at the correct site, as marked on the
map, including: 1:24,000 USGS map orienteering, topographic landmarks, county road maps,
and global positioning system (GPS) confirmation of site latitude and longitude.
Classify the site. AT THE X-SITE. as:
NON-TARGET
TARGET
INACCESSIBLE
No Stream Channel
Impounded Stream
Marsh/Wetland
Unwadeable Stream (> 50% of reach is unwadeable)
Regular Stream
Intermittent Stream
Dry Channel
Altered Channel (stream channel different form map representation)
Physical Barriers (Physically unable to reach the X-site)
No Permission
Record class on Site Verification form, do not sample Non-target or inaccessible sites. Take
samples from Target sites as discussed in field operations and methods manual.
3. Measure the stream width at five "typical" places within 10 m of the X-site. Average and round
the width to the nearest meter. Record width on the stream site verification form. Lay out a
sample reach with a length of 40 times the stream width. If the stream is less than 4 m wide,
use 150 m as the sample reach length.
4. Do a reconnaissance of the sample reach.
5. Proceed downstream half the required reach length; measure the distance with a tape
measure down the middle of the stream. Mark it as the reach start point (Transect "A").
6. Proceed upstream marking 10 more cross-section transects (Transects "B" through "K") at 1/10
intervals along the calculated reach length (every 4 channel widths or 1.5 meters in small
streams). At Transect "B", assign a sampling point (Left, Center, or Right as you face
downstream) for collecting periphyton and benthic macroinvertebrate samples by throwing a
die. Once the initial point has been determined, assign sampling points for Transects "C"
through "J" systematically using the order Left, Center, and Right.
NOTE: If there is a lake/pond or a stream order change (100,000 map-based) along the survey
reach, end the sample reach at the barrier. Make up for the loss of stream length by adding length
to the other end of the reach ("slide" the reach). Locations where the stream order changes will be
noted on the topo maps provided to the field teams. Do not "slide" the reach to avoid bridges, riprap,
small flow control structures, culverts and the like.
B-3
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4. September 1998 Page 5 of 31
QUICK REFERENCE GUIDE FOR WATER CHEMISTRY
I. EQUIPMENT TO CARRY IN FIELD FOR WATER CHEMISTRY
Rinse/Test bottles of QCCS in self-sealing plastic bag
D.O./Temperature/Conductivity Meter
Field Forms
One 500-mL plastic beaker with handle, in clean self-sealing plastic bag
One cubitainer in clean self-sealing plastic bag (barcode label attached)
Two 60-mL syringes in a plastic container (each one with a bar code label attached)
Two syringe valves in the plastic container
Opaque garbage bag
II. EXTRA EQUIPMENT TO CARRY IN VEHICLE
Cooler with 4 to 6 one-gallon self-sealing plastic bags filled with ice
Back-up labels, forms, syringes, and syringe valves
III. DAILY ACTIVITIES AFTER SAMPLING
1. Check that cubitainer lid is on tight and has a flush seal.
2. Prepare the sample for shipping (label and seal cooler, replace ice as close as possible to
shipping time).
3. Call Overnight shipping company to arrange pick-up of cooler.
4. Rinse the sampling beaker with deionized water three times.
5. Make sure field meters are clean and are stored with moist electrodes.
6. Label the next days sample containers (cubitainer and syringes), pack cubitainer and sample
beakers in clean self-sealing plastic bag, and pack two syringes and syringe valves in a plastic
container with a snap-on lid.
(continued)
B-5
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 6 of 31
QUICK REFERENCE GUIDE FOR WATER CHEMISTRY (Continued)
.SUMMARY OF SITE PROCEDURE FOR WATER CHEMISTRY
I. COLLECT WATER SAMPLE
A. Make sure cubitainers and syringes are labeled and have the same barcode ID.
B. Rinse the 500-mL sample beaker three times with streamwater from mainstream.
C. Rinse cubitainer three times with 25-50 mL of streamwater, using the sample beaker.
Rinse cubitainer lid with stream water.
D. Fill cubitainer with streamwater using the 500 mL sample beaker. Expel any trapped air
and cap the cubitainer. Make sure that the lid is seated correctly and that the seal is
tight.
DO NOT EXPAND CUBITAINER BY BLOWING IN IT.
E. Rinse each of the two, 60-mL syringes three times with 10-20 mL of streamwater.
F. Fill each of the syringes with streamwater from mid-stream by slowly pulling out the
plunger. If any air gets into the syringe, discard the sample and draw another.
G. Invert the syringe (tip up) and cap the syringe with a syringe valve. Open the valve, tap
the syringe to move any air bubbles to the tip, and expel any air and a few mL of water.
Make sure there is 50-60 mL of stream sample in the syringe. Close the valve and place
the syringes in their transport container.
H. Place the cubitainer and syringes in cooler/stream to keep cool (keep dark as well) while
the rest of the sampling is taking place. When you return to the vehicle, put the samples
in the cooler and surround with 4 to 6 one-gallon self-sealing plastic bags filled with ice.
II. IN SITU MEASUREMENTS
A. Conductivity
1., Turn on and check the zero and red line (if applicable) of the conductivity meter.
2. Measure and record the conductivity of the QCC solution. Rinse the probe in the
"Rinse" bottle of QCC solution before immersing in the "Test" bottle of QCC
solution.
3. Measure and record stream conductivity in mid-stream.
B. Dissolved Oxygen/Temperature
1. Calibrate the DO meter following meter instructions.
2. Measure the DO and temperature in mid-stream. If water velocity is slow, jiggle the
DO probe as you take the reading. _^====___ ———=======
B-6
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EMAP^SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 7 of 31
QUICK REFERENCE GUIDE FOR PHYSICAL HABITAT CHARACTERIZATION
FIELD SUMMARY: P-HAB LAYOUT AND WORKFLOW .
1. Habitat Sampling Layout:
Thalweq interval: 1.0 m for streams <2.5 m wide (from initial estimate).
1.5 m for streams 2.5 - 3.5 m wide
0.01 * (reach length) for streams >3.5 m wide
100 thalweg measurement intervals in each sample reach, except 150 in streams <2.5m wide
Channel/Riparian Cross Section Transect every 10th thalweg interval (every 15th for channels
<2.5m wide). Eleven of them, marked "A" thru "K".
Wetted Width at every cross-section transect and halfway in between transects (total of 21
measurements).
2. Work Flow:
At the downstream start point (Transect "A"), one person makes channel dimension,
substrate, bank, and canopy densiometer measurements. The second person records
those measurements while making visual estimates of riparian vegetation structure, fish
cover, and human disturbance. No bearing or slope at first cross section.
Proceed upstream between Transects "A" and "B", making measures at each thalweg
measurement station. One person in channel measures width (when required), thalweg
depth, and determines presence of soft/small sediment at thalweg. The other person
records those measurements, classifies channel habitat, and makes large woody debris
estimates.
When you complete 10 thalweg intervals and reach one of 11 pre-marked cross section
transect flags, stop and take out a new cross-section form for Transect "B". Repeat all
the Channel/Riparian measurements at this new location. In addition, do the slope &
bearing backsites together. Intermediate flagging (of a different color) may have to be
used if the stream is extremely brushy, sinuous, or steep to the point that you cannot site
for slope and bearing measures between the 11 points. (Note that you could tally woody
debris while doing the backsite, rather than during the thalweg profile measurements.)
Repeat the cycle of thalweg and cross section measurements until you reach transect 11
("K") at the upstream end. •
Discharge measurements made any time after choosing suitable location nearest to the
"X" site. Discharge measurements are done by the Chemistry/Macroinvert pair (rather
than the Habitat/Fish pair) just after chemical samples are taken.
(continued)
B-7
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 8 of 31
QUICK REFERENCE GUIDE FOR PHYSICAL HABITAT CHARACTERIZATION (Continued)
FIELD SUMMARY: COMPONENTS OF P-HAB PROTOCOL
Width, Depth Profile, Hab Classes, Woody Debris:
At 10 (15) equally spaced intervals between each of 11 channel cross-sections (100 or
150 along entire reach):
Measure max. depth ("Thalweg") at each increment and wetted width at the
required increments.
Classify habitat and pool-forming elements.
Determine presence of soft/small sediment at thalweg measurement points.
Between each of the channel cross sections, tally all Large Woody debris within and
above the bankfull channel according to size class. In the tally boxes provided on the
form, make separate tally for LWD wholly or partially within the bankfull channel and then
for LWD only bridging above the channel.
NOTE: If initial width estimate is <2.5 m, then 150 thalweg measurements are made at 1.0 m
intervals over a 150 m reach. If width is 2.5 to 3.5 m, then make 100 thalweg measurements
at 1.5 m intervals. In all other cases, 100 measurements are made at an interval 1/100th the
length of the sample reach.
Channel and Riparian Cross-Sections:
Measurements: Bankfull width, bankfull height, incision height, wetted width, bar width,
undercut, bank angle (with rod and clinometer); gradient (clinometer), sinuosity (compass
backsite), riparian canopy cover (densiometer).
Visual Estimates: Substrate size class and embeddedness; areal cover class and type of
riparian vegetation in-Canopy, Mid-Layer and Ground Cover; areal cover class offish
cover features, aquatic macrophytes, and filamentous algae; presence and proximity of
human disturbances
Discharge:
In medium and large streams measure water depth and velocity (at 0.6 depth from surface) at
15 to 20 equally spaced intervals across one carefully chosen channel cross-section. Let
meter equilibrate to average velocity for 20 seconds. In very small streams, measure
discharge by timing the passage of a neutrally-buoyant object 3 times or the filling of a bucket
5 times in succession.
(continued)
B-8
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EMAP-SW-Streams Field Operations Manual. Appendix B, Rev. 4, September 1998 Page 9 of 31
QUICK REFERENCE GUIDE FOR PHYSICAL HABITAT CHARACTERIZATION (Continued)
FIELD SUMMARY: RIP. VEG., HUMAN DISTURB., IN-CHANNEL COVER:
Observations upstream 5 meters and downstream 5 meters from each of the 11 cross-section
transects.
For riparian vegetation and human disturbances, include the visible area from the stream back
a distance of 10m (30 ft) shoreward from both the left and right banks. If the wetted channel is
split by a mid-channel bar, the bank and riparian measurements shall be for each side of the
channel, not the bar.
Three vegetation layers:
CANOPY LAYER (>5 m high)
UNDERSTORY (0.5 to 5 m high)
GROUND COVER layer (O.5 m high).
Canopy and Understory Vegetation Types:
(EJeciduous, Coniferous, Broadleaf Evergreen, Mixed, or Mane) in each of the two taller
layers (Canopy and Understory). "Mixed" if more than 10% of the areal coverage made
up of the alternate type.
Areal Cover Classes for Vegetation and In-Channel Cover:
0: (absent — zero cover)
1: (sparse— cover <10%) '
2: (moderate - cover 10-40%)
3: (heavy-cover40-75%)
4: (very heavy — cover >75%).
Tallying Human Disturbances:
B: The human activity or structure is ON THE STREAMBANK
C: CLOSE to the Bank (within 10m)
P: PRESENT, but farther than 10m from the bank
0: NOT PRESENT.
(continued)
B-9
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EMAP-SW-Streams Field Operations Manual, Appendix B. Rev. 4, September 1998 Page 10 of 31
QUICK REFERENCE GUIDE FOR PHYSICAL HABITAT CHARACTERIZATION (Continued)
FIELD SUMMARIES: SUBSTRATE AND WOODY DEBRIS SIZE CLASSES
Substrate size class and embeddedness are estimated, and depth is measured for 5 particles taken
@ 5 equally-spaced points on each cross-section. The cross-section is defined by laying the
surveyor's rod or tape to span the wetted channel.
SUBSTRATE SIZE CLASSES:
RS Bedrock (Smooth) >4000 mm
RR
BL
CB
GO
GF
SA
FN
HP
WD
Bedrock (Rough
Boulders
Cobbles
Gravel(Coarse)
Gravel (Fine)
Sand
Fines
Hardpan
Wood
OT Other
>250 to 4000 mm
64 to 250 mm
16 to 64 mm
2 to 16 mm
0.06 to 2 mm
<0.06 mm
>4000 mm
Regardless of
Size
Regardless of
Size
smooth surface rock or hardpan (bigger than a car)
(bigger than a car)
(basketball to car size)
(tennis ball to basketball size)
(marble to tennis ball size)
(ladybug to marble size)
(smaller than ladybug size, but visible as particles - gritty
between fingers).
Silt-Clay-Muck (not gritty between fingers)
(consists of firm, consolidated fines)
Wood or other organic material
Metal, Tires, Car bodies, asphalt, concrete, etc. (Describe
in comments if you enter "CT").
-ARGE WOODY DEBRIS SIZE CLASSES
LWD Definition:
Two Tallvs:
Diameter (small end) > 0.1 m (>4 in)
Length > 1.5 m (> 5 ft) -- count only part with diarn > 0.1 m.
(1) LWD at least partially within bankfull channel.
(2) LWD not within bankfull channel, but at least partially bridging above bankfull stage
(idea is that it will eventually fall into channel).
Size Categories for Tally (12 potential combinations):
Diameter
0.1 to <0.3 m
0.3 to <0.6 m
0.6 to <0.8 m
>0.8 m
(large end):
(4 to 12 inches)
(1 to 2 ft)
(2 to 2.6 ft)
(>2.6 ft)
Length:
1 .5 - <5 m
5-15m
>15 m
(5 -16 ft)
(16 -49 ft)
(>49 ft)
B-10
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 11 of 31
QUICK REFERENCE GUIDE FOR PHYSICAL HABITAT CHARACTERIZATION (Continued)
FIELD SUMMARY: HABITAT CLASSIFICATION AT CHANNEL UNIT SCALE
Channel Unit Habitat Classes9
Class (Code)
Pools: Still water, low velocity,
Plunge Pool (PP)
Trench Pool (PT)
Lateral Scour Pool (PL)
Backwater Pool (PB)
Impoundment Pool (PD)
Pool (P)
Glide (GL)
Riffle (Rl)
Rapid (RA)
Cascade (CA)
Falls (FA)
Dry Channel (DR)
Description
smooth, glassy surface, usually deep compared to other parts of the channel:
Pool at base of plunging cascade or falls.
Pool like trench in stream center
Pool scoured along one bank.
Pool separated from main flow off side of channel.
Pool formed by impoundment above dam or constriction.
Pool (unspecified type).
Water moving slowly, with smooth, unbroken surface. Low turbulence.
Water moving, with small ripples, waves and eddies — waves not breaking, surface
tension not broken. Sound: "babbling", "gurgling".
Water movement rapid and turbulent, surface with intermittent Whitewater with breaking
waves. Sound: continuous rushing, but not as loud as cascade.
Water movement rapid and very turbulent over steep channel bottom. Most of water
surface broken in short irregular plunges, mostly Whitewater. Sound: roaring.
Free falling water over vertical or near vertical drop into plunge, water turbulent and white
over high falls. Sound: from splash to roar.
No water in channel
Code
N
W
R
B
F
WR, RW, RBW
OT
Categories of Pool-forming Elements
Category
Not Applicable, Habitat Unit is not a pool
Large Woody Debris.
Rootwad
Boulder or Bedrock
Unknown cause (unseen fluvial processes)
Combinations
Other (describe in comments section of field form)
(continued)
B-11
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EMAP-SW-Streams Field Operations Manual. Appendix B, Rev. 4. September 1998 Page 12 of 31
QUICK REFERENCE GUIDE FOR PHYSICAL HABITAT CHARACTERIZATION (Continued)
FIELD SUMMARY: P-HAB PROBLEM AREAS
Mid-channel Bars: dry at baseflow, inundated at bankfull flow.
Measure wetted width across and over mid-channel bars, but record bar width in the column
provided on thalweg profile and cross-section form.
Islands: as high as the surrounding flood plain; dry even at bankfull flow.
Measure only the width of the main channel between island and shore; then if required,
measure the side channel width separately (record on another form). Handle the side
channels created by islands as follows:
* Visually estimate the percent of flow in the side channel.
* If <15% - Indicate presence of side channel on field data form.
* If 16-49% - Indicate presence of side channel, plus obtain and record detailed
channel & riparian cross-section measurements on the side channel. Designate
additional cross section transects as "XA", "XB", etc. corresponding to nearest main
channel Transect location.
Note continuous presence of side channels on the Thalweg Profile Form until channels
converge. In addition, note the points of side channel convergence and divergence in the
comment section on the thalweg profile form.
Drv and Intermittent Streams, where no water is in the channel:
Record zeros for depth and wetted width.
Record habitat type as dry channel ("DR").
Make all Channel Cross-section transect measures across the unvegetated portion of the
channel. For substrate, DiSTLB = the width of the unvegetated channel, and substrate
measurements are made along that transect.
B-12
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 13 of 31
QUICK REFERENCE GUIDE FOR PERIPHYTON
FIELD EQUIPMENT
1. Large funnel (15-20 cm diameter).
2. Scrape area delimiter (3.8 cm diameter pipe, 3 cm tall).
3. Stiff-bristle toothbrush with handle bent at 90° angle.
4. Wash bottle.
5. Collection bottle to catch removed periphyton.
6. 60 mL syringes with 3/8" hole bored into the end.
7. 50 mL centrifuge tubes or similar sample vials.
8. Formalin.
9. Glass-fiber filters (0.45 urn average pore size) for chlorophyll a.
10. Pre-leached, pre-ashed, weighed glass-fiber filters (0.45 urn average pore size) in
numbered pans for ash free dry mass (AFDM).
11. Forceps for filter handling.
12. Millipore®-type filtration apparatus with plastic or stainless steel filter base, and Nalgene®
funnel and suction flask.
13. Nalgene® hand-operated vacuum pump (need one additional pump as a backup).
14. Aluminum foil.
15. Ice chest.
FIELD PROTOCOLS
1. Periphyton samples will be collected using a random-systematic procedure. The location
(left, middle, or right 1/3 of the channel) of the first sample (Transect B) will be chosen
randomly. Subsequent samples (Transects C-J) will be collected sequentially from the
left, middle, then right 1/3 of the channel, resulting in three samples from each side and
middle.
Periphyton are collected, using the appropriate method, from flowing (riffles) and slack
water (pools) habitats.
Rock and wood samples which are small enough (< 15 cm diameter) and can be easily
removed from the stream are collected by placing the substrate in a funnel which drains
into a sample bottle. A defined area of substrate surface (12 cm2)is enclosed, and
attached periphyton is dislodged with 30 seconds of brushing with a stiff-bristled
toothbrush. Care must be taken to ensure that the upper surface of the rock is the
surface that is being scraped.
Loosened periphyton is then washed, using stream water from a wash bottle, from the
substrate into the 500-mL sample bottle.
Soft-sediments are collected by vacuuming the upper 1 cm of sediments confined within
the 12-cm2 sampling ring into a 60-mL syringe.
All samples, regardless of substrate type, are composited by habitat (riffle or pool) and
mixed thoroughly.
Record total volume of composited sample before proceeding to the next step!
(continued)
2.
3.
5.
6.
7.
B-13
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EMAP-SW-Streams Field Operations Manual. Appendix B, Rev. 4, September 1998 Page 14 of 31
QUICK REFERENCE GUIDE FOR PERIPHYTON (Continued)
8. Four subsamples will be taken from each composite sample. These are:
a. Identification/Enumeration
1) Withdraw 50 ml of mixed sample and place in a labeled sample vial (50-mL
centrifuge tubes work well). Cover label with clear tape.
2) Preserve sample with 2 mL of 10% formalin. Gloves should be worn.
3) Tightly cap tube and tape with electrical tape.
b. Chlorophyll a
1) Withdraw 25 mL of mixed sample and filter onto a glass-fiber filter (0.45 um
pore size) using a hand-operated vacuum pump. (Note: for soft-sediment
samples, allow grit to settle before withdrawing sample).
2) Fold filter so that the sample on the filter surface is folded together, wrap in
aluminum foil, and affix the tracking label to the outside, and seal with clear
tape.
3) Freeze filter as soon as possible by placing it in a freezer.
4) Store frozen for laboratory analysis.
c. Ash Free Dry Mass (AFDM)
1) Withdraw 25 mL of mixed sample and filter onto a pre-leached, pre-weighed
glass-fiber filter. (Note: for soft-sediment samples, allow grit to settle before
withdrawing sample).
2) Do not fold this filter. Return filter to it's numbered container, wrap in
aluminum foil, affix tracking label to outside, and seal with clear tape.
3) Freeze filter as soon as possible by placing it in a freezer.
4) Store frozen for laboratory for analysis.
d. Alkaline/Acid Phosphatase
1) Withdraw 50 mL of mixed sample and place in a labeled sample vial (50-mL
centrifuge tubes work well). Cover label with clear tape.
2) Tightly cap tube and tape with electrical tape.
3) Freeze sample as soon as possible by placing it on dry ice.
4) Store frozen for laboratory analysis.
B-14
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EMAP-SW-Streams Field Operations Manual. Appendix B, Rev. 4, September 1998 Page 15 of 31
QUICK REFERENCE GUIDE FOR SEDIMENT METABOLISM
FIELD EQUIPMENT
1. Ice chest for floating centrifuge tubes during incubation
2. 1000 ml_ Nalgene® beaker for holding centrifuge tubes during incubation.
3. Small scoop sampler for sediments.
4. 50-mL, screw-top, centrifuge tubes.
5. Digital dissolved oxygen meter (e.g. YSI 58) with a stirring probe (e.g., YSI 5730).
6. Spare batteries for D.O. meter.
7. Permanent markers for labeling tubes.
8. Sample labels and field data sheets.
9. Ice chest with dry ice for sample freezing.
FIELD PROTOCOLS
Dissolved Oxygen Meter Calibration (for YSI model 58, with YSI model 5730 stirring BOD probe)
1. Zero meter according to manufacturer's directions, and
2. Calibrate meter using the water-saturated atmosphere method described in the meter's
operations manual.
B-15
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 16 of 31
QUICK REFERENCE GUIDE FOR SEDIMENT METABOLISM (Continued)
Sediment Collection and Experimental Set-up
1. Collect and combine fine-grained, surface sediments (top 2 cm) from all depositional
areas along the stream reach (Transects B-J) established for the Physical Habitat
Characterization.
2. Fill ice chest 2/3 full with stream water and record temperature and dissolved oxygen
(D.O.).
3. Thoroughly mix composite sediment sample.
4. Place 10 mL of sediment in each of 5 labeled, 50 mL screw-top centrifuge tubes.
5. Fill each tube to the top (no head space) with stream water from the ice chest and seal.
6. Fill one additional tube with stream water only to serve as a blank.
7. Incubate tubes in closed ice chest for 2 hours.
8. Measure D.O. in each tube, including the blank.
9. Decant overlying water and save sediment.
10. Tightly seal tubes and freeze as soon as possible.
11. Store frozen for laboratory analysis.
12. If you are collecting samples for sediment toxicity tests, save 1 to 2 L of the remaining
sediment sample by placing it in a labeled plastic bag.
13. Store sediment toxicity sample chilled (but not frozen!) for laboratory analysis.
B-16
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EMAP-SW-Streams Field Operations Manual. Appendix B, Rev. 4, September 1998 Page 17 of 31
QUICK REFERENCE GUIDE FOR SEDIMENT TOXICITY
SAMPLE COLLECTION AND SHIPMENT FOR SEDIMENT TOXICITY SAMPLES
1. Use the sediment left over from the benthic (sediment) metabolism indicator in Section 9,
Benthic (Sediment) Metabolism: Field Methods.
2. Mix sediment well with a stainless steel or plastic mixing spoon or gloved hand.
3. Fill a 2-gallon polyethylene (4 mil) bag with at least 1 L of sediment.
4. Close bag, squeeze the air out and tie a knot in the remaining portion of the bag to seal.
5. Fill out ID label; place label on the outside of the bag. Place this bag inside a second 2-gallon
polyethylene bag and tie off the top to seal.
6. Place these bags inside a cooler with only sediment samples in them.
7. Hold sediment samples on ice (do not freeze!) for laboratory analysis.
8. Ship samples to the designated contact person or laboratory.
B-17
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-------
^ EMAP-SW-Streams Field Operations Manual, Appendix B. Rev. 4, September 1998 Page 19 of 31
QUICK REFERENCE GUIDE FOR BENTHIC MACROINVERTEBRATES
TABLE I. BASE PROTOCOLS FOR COLLECTING MACROINVERTEBRATES
1. Do the water chemistry.
2. Locate first sampling station (second flag) from downstream end of the study segment and roll
die to pick left (1), middle (2), or right side (3) of transect to sample. If stream is narrower than
three nets, pick left or right. If wide enough for only one net, then sample entire stream width.
After first transect, systematically sample remaining transects left, middle, or right so that three
samples are collected on left, middle, and right at the site.
3. If riffle or run use protocol in Table II. If pool use protocol in Table III or hand pick for 60
seconds if kick net cannot be used.
4. Go to next upstream station and repeat. Combine all riffle samples in one bucket and pool
samples in another. Check net after each sample for clinging organisms and transfer to
bucket.
5. After a sample is collected from each of nine interior transects and all samples are combined in
the proper bucket, obtain a composite sample as described in Table IV.
6. Assist with the fish collection.
7. Preserve and label each sample as described in Table V.
(continued)
B-19
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EMAP-SW-Streams Field Operations Manual, Appendix B. Rev. 4, September 1998 Page 20 of 31
QUICK REFERENCE GUIDE FOR BENTHIC MACROINVERTEBRATES (Continued)
TABLE II. PROCEDURES FOR RIFFLES AND RUNS USING KICK NET SAMPLER
1. Attach four foot pole to the sampler.
2. Position sampler quickly and securely on stream bottom with net opening upstream.
3. Hold the sampler in position on the substrate while checking for snails and clams in an area of
about 0.5 m2 in front of the net; kick the substrate vigorously for about 20 seconds in front of
the net.
4. Inspect and rub off with the hands any organisms clinging to the rocks, especially those
covered with algae or other debris.
5. Remove the net from the water with a quick upstream motion to wash the organisms to the
bottom of net.
6. Rinse net contents into the "riffle" bucket containing one or two gallons of water by inverting
the net in the water.
7. Inspect the net for clinging organisms. With forceps remove any organisms found and place
them into the bucket.
8. Large objects (rocks, sticks, leaves, etc.) in the bucket should be carefully inspected for
organisms before discarding.
9. Combine all riffle samples in the "riffle" bucket.
10. After all stations are sampled and all riffle samples combined in the "riffle" bucket, obtain a
composite sample as described in Table IV.
(continued)
B-20
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 21 of 31
QUICK REFERENCE GUIDE FOR BENTHIC MACROINVERTEBRATES (Continued)
TABLE III. PROCEDURES FOR POOLS USING THE MODIFIED KICK NET SAMPLER
1. Attach four-foot pole to the sampler.
2. Inspect about 1/2 square meter of bottom for any heavy organisms, such as mussels and
snails, which have to be hand picked and placed in the net.
3. While disturbing about 0.5 m2 of substrate by kicking, collect a 20-second sample by dragging
the net repeatedly through the area being disturbed. Keep moving the net all the time so that
the organisms trapped in the net will not escape.
4. After 20 'seconds remove the net from the water with a quick upstream motion to wash the
organisms to the bottom of the net.
5. Rinse net contents into a small bucket of water (about one or two gallons) by inverting the net
in the water.
6. Inspect the net for clinging organisms. With forceps remove any organisms found and place
them in the bucket.
7. Large objects in the bucket should be carefully inspected for organisms which are washed into
the bucket before discarding.
8. Combine this sample with the other pool samples in the "pool" bucket.
9. After all stations are sampled and all pool samples are combined together in the "pool" bucket,
obtain a composite sample as described in Table IV.
(continued)
B-21
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 22 of 31
QUICK REFERENCE GUIDE FOR BENTHIC MACROINVERTEBRATES (Continued)
TABLE IV. PROCEDURES FOR OBTAINING THE COMPOSITE SAMPLE
1. Pour the contents of the riffle bucket through a U.S. Standard 30 sieve. Examine the bucket
while rinsing it well to be sure all organisms are washed from the bucket onto the sieve.
2. Wash contents of the sieve to one side by gently agitating in water and wash into jar using as
little water from the squirt bottle as possible. Carefully examine the sieve for any remaining
organisms and place them in the jar.
3. Place properly filled out waterproof label in the jar and replace the cap.
TABLE V. SAMPLE PRESERVING AND LABELING
1. Fill in special pre-numbered barcoded label and place on jar. All additional jars used for a
sample must be labeled with same number. Enter this number which will be used for tracking
purposes in the computer.
2. Preserve samples in ethanol as follows:
a. If jar is more than 1/4 full of water, pour off enough to bring it to less than 1/4 full using
proper sieve to retain organisms.
b. Fill jar nearly full with 95% ethanol so that the concentration of ethanol is 70%. If there is
a small amount of water in the sample, it may not be necessary to fill the jar entirely full to
reach a 70% concentration.
c. Transfer any organisms on the sieve back into the jar with forceps.
3. Check to be sure waterproof label is in jar with the required information on it.
4. Check to be sure that the pre-numbered stick-on barcoded label is the on jar and agrees with
the inside label. Cover the entire label with clear, waterproof tape.
5. With a grease pencil write the site number, sample type (Riffle or Pool), and the number of
transects sampled for either Riffle or Pool on the cap.
6. Seal the caps with electrical tape.
7. Complete the check off sheet and place samples in cooler or other secure container for
transport.
8. Secure all equipment in the vehicle.
(continued)
B-22
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 23 of 31
QUICK REFERENCE GUIDE FOR BENTHIC MACROINVERTEBRATES (Continued)
TABLE VI. MACROINVERTEBRATE SAMPLING ACTIVITIES CHECKLIST
Date:
Time:
Site No.:
Stream Name and Location:
Crew ID: 123456
Collector:
1.
Initial observations, if any, on the Sample Collection Form - Streams.
2.
Composite riffle/run sample collected with label inside jar.
3.
Composite pool/glide sample collected with label inside jar.
4.
Correct barcode and label on all jars and sealed with clear, waterproof tape.
5.
All samples preserved.
6.
With a grease pencil write site number, sample type (Riffle or Pool), and number of
transects sampled for sample type on the cap. If two jars are used be sure to mark them
as such.
7.
Caps sealed with tape.
8.
Photos of site.
Sample jars in cooler or otherwise secured.
10. All equipment accounted for and secured in vehicle.
Signature:
Time sampling completed:
B-23
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EMAP-SW-Streams Field Operations Manual, Appendix B. Rev. 4, September 1998 Page 25 of 31
QUICK REFERENCE GUIDE FOR AQUATIC VERTEBRATES
FIELD PROTOCOLS FOR FISH COLLECTION
1. Site Selection
a. Determine channel width.
b. Survey sample reach.
c. Determine if reach requires block nets.
d. If conductivity is below 10 uS/cm or if flow, depth or turbidity make it unsafe to
electrofish, crew may elect to use seine only or not sample. THIS IS A SAFETY
DECISION.
e. In case of emergency, determine location of means of easy egress from stream.
2. Electrofishina
a. Set unit to 300VA and pulsed DC. Select initial voltage setting. Start generator, set
timer, and depress switch to begin fishing.
b. Fish in an upstream direction, parallel to the current. Adjust voltage and waveform
output according to sampling effectiveness and mortality fish specimens.
c. With switch depressed, sweep electrodes from side to side in the water. Sample
available cut-bank and snag habitat as well as riffles and pools.
d. Netters follow operator and net fish. Deposit fish in buckets. Block with seines in riffles,
pools and snags.
e.
Continue for 40 channel widths. Record total time spent collecting and shocking time on
data sheets.
f. Identify and release any threatened and endangered species.
g. Identify and measure (SL, TL) sport fish and very large specimens, record external
anomalies, and release unharmed.
h. Identify other specimens. Determine number of individuals in species, measure largest
and smallest individuals, and voucher as described in Voucher Protocol.
i. Retain a subsample of target species for Fish Tissue Contaminants analysis.
3. Seining will be used in conjunction with electrofishing and in sites where stream is too deep for
electrofishing to be conducted safely.
(continued)
B-25
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EMAP-SW-Streams Field Operations Manual. Appendix B. Rev. 4. September 1998 Page 26 of 31
QUICK REFERENCE GUIDE FOR AQUATIC VERTEBRATES (Continued)
ANOMALY CATEGORIES AND CODES
— — •• —
Absent
Blisters
Blackening*
Extensive Black spot disease
Cysts
Copepod
Deformities
Eroded fins
Eroded gills
Fungus
Fin anomalies
Grubs
Hemorrhaging
Ich
Lesions
Lice
Mucus
None
Other
Scale anomalies
Shortened operculum
Tumors
Leeches
Code
AB
BL
BK
BS
CY
CO
DE
EF
EG
FU
FA
GR
HM
1C
LE
LI
MU
NO
OT
SA
SO
TU
WR
EX
Definition 1
Absent eye, fin, tail.
n mouth, just under skin.
Tail or whole body with darkened pigmentation.
Small black cysts (dots) all over the fins and body.
Fluid-filled swellings; maybe small dots or large.
A parasitic infection characterized by a worm like copepod embedded in the flesh
of the fish; body extends out and leaves a sore/discoloration at base, may be in
mouth gills, fins, or anywhere on body.
Skeletal anomalies of the head, spine, and body shape; amphibians may have
extra tails, limbs, toes.
Appear as reductions or substantial fraying of fin surface area.
Gill filaments eroded from tip.
May appear as filamentous or "fuzzy" growth on the fins, eyes, or body.
Abnormal thickenings or irregularities of rays
White or yellow worms embedded in muscle or fins.
Red spots on mouth, body, fins, fin bases, eyes, and gills.
White spots on the fins, skin or gills.
Open sores or exposed tissue; raised, granular or warty outgrowths.
Scale-like, mobile arthropod.
Thick and excessive on skin or gill, as long cast from vent. 1
No anomalies present.
Anomalies or parasites not specified.
Missing patches, abnormal thickings, granular skin
Leaves a portion of the gill chamber uncovered
Areas of irregular cell growth which are firm and cannot be easily broken open 1
when pinched. (Masses caused by parasites can usually be opened easily.)
Annelid worms which have anterior and posterior suckers. They may attach
anywhere on the body.
(continued)
B-26
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 27 of 31
QUICK REFERENCE GUIDE FOR AQUATIC VERTEBRATES (Continued)
GUIDELINES AND PROCEDURES FOR PREPARING FISH VOUCHER SPECIMENS
Category 1. Large easily identified species OR adults may be difficult to identify OR the species is
uncommon in that region. Preserve 1-2 small (<150 mm total length) adult individuals per site plus
2-5 juveniles. If only large adults are collected, reserve smallest individual until voucher procedure
is complete and preserve ONLY if space is available. Photograph if considered too large for the jar.
Category 2. Small to moderate-sized fish OR difficult to identify species. Preserve 25 adults and
juveniles. If less than 25 individuals are collected, voucher all of them.
Category 3. Species of "special concern." These are state or federally listed species. Photograph
and release. If specimens have died, include in voucher collection, note on data sheet and notify
appropriate state official as soon as possible.
a.
b.
c.
After all individuals of a species have been processed, place the voucher sub sample in a kill
jar containing a strong (approximately 20%) formalin solution. Individuals > 160 mm should be
slit on the lower abdomen of the RIGHT side.
When specimens are dead, transfer to a small nylon bag containing a waterproof label with tag
#. Place in "Voucher" jar in 10% formalin. BE SURE THAT JAR IS LABELED INSIDE AND
OUT WITH A VOUCHER LABEL (site ID, barcode, and date).
Continue until all species are processed. Seal voucher jar with electrical or clear tape. Check
that the jar is correctly labeled. Enter BARCODE ID in appropriate place on field data sheet.
d. Transport to storage depot at end of week. Store in a cool, dark, ventilated space.
B-27
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 29 of 31
QUICK REFERENCE GUIDE FOR FISH TISSUE CONTAMINANTS
SELECTING FISH TISSUE SPECIMENS
If possible, obtain one sample each, containing the desired weight or number (see below) of
similarly sized individuals*, from the primary and secondary target species lists (2 composite
samples total):
I. PRIMARY TARGET SPECIES
Small adult fish
(in priority order)
1) Blacknose Dace
2) Another Dace species
3) Creek Chub or Fallfish
4) Slimy Sculpin/Mottled Sculpin
5) Stoneroller
6) A Darter species
7) A Shiner species
DESIRED
WEIGHT
50** - 400 g
50** - 400 g
50** - 400 g
50** - 400 g
50** - 400 g
50** - 400 g
50** - 400 g
A) Choose the highest priority target species from the above list, that has at least enough
individuals to attain the minimum weight (50 g). Get as much weight of fish as possible
within the desired weight range (50-400 g). Use scale provided to determine weight.
With clean hands, place the fish in fresh aluminum foil (dull side towards fish) before
placing fish in weighing container.
(B) If fewer than the desired number of individuals of any primary target species are
collected, send individuals of a small non-target species if 50 g or more are available.
* - Getting a sufficient sample amount is a higher priority than getting similar-sized individuals.
** - This weight represents the minimum amount needed for laboratory analysis. Crews
should not settle for the minimum weight if more fish are present. They should send as many
fish as possible up to 400 g weight goal.
B-29
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EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4. September 1998 Page 30 of 31—
QUICK REFERENCE GUIDE FOR FISH TISSUE CONTAMINANTS (Continued)
SELECTING FISH TISSUE SPECIMENS
II. SECONDARY TARGET SPECIES
Collect and save a sample of secondary target species if such a sample of desired number of
individuals of desired size is available. Collect similar sized individuals if enough are present.
A)
B)
C)
Larger adult fish DESIRED DESIRED
(in priority order) SIZE NUMBER
White sucker >120 mm 5
Hogsucker >120 mm 5
A Bass species >120 mm 5
A Trout species >120 mm 5
A Sunfish species >120 mm 5
Carp >120mm 5
D
2)
3)
4)
5)
6)
If fewer than the desired number of secondary target species individuals of desired size are
collected, add smaller individuals of the same species, if available, to achieve the desired
number (5).
If fewer than 5 fish of any size are available, you may send as few as 3 fish that are at or at
least near the minimum desired size (120 mm).
If an acceptable secondary target species sample (by the above criteria) is not available send
only the primary target species sample. If neither a primary nor secondary species sample
that meets these criteria is available, use your best judgement in sending some type offish
sample (may be mixed species).
(continued)
B-30
-------
EMAP-SW-Streams Field Operations Manual, Appendix B, Rev. 4, September 1998 Page 31 of 31
QUICK REFERENCE GUIDE FOR FISH TISSUE CONTAMINANTS (Continued)
PROCESSING TISSUE SPECIMENS
1. Keep hands, work surfaces, and wrapping materials clean and free of potential contaminants
(mud, fuel, formalin, sun screen, insect repellant, etc.)
2. Measure total weight of individuals for primary target species and count the total number of
individuals. Measure the total length (TL) of each secondary target species individual.
Record all of this information in the fish tissue section of the Sample Collection Form.
3. Write the bar-code number(s)*** on the collection form. Make sure that the form is filled out
completely.
4. Wrap fish in aluminum foil. Place the dull side of the aluminum foil in contact with the fish. The
primary target fish sample may be wrapped as a group. Secondary target fish should be
wrapped individually. Once wrapped, place each sample in a self-sealing plastic bag or a
garbage bag.
5. Expel excess air and seal the bag(s). Wrap clear tape around the bag(s) to seal and make a
surface for each sample label.
6. Complete bar-coded fish tissue label(s). Make sure the number(s) is/are the same one(s) on
the collection form. Apply it/them to the tape surface(s). Cover the label(s) with clear,
waterproof tape. As labels will sometimes fall off, there should always be a label on the inner
bag.
7. Place labeled bag(s) into a second plastic bag(s) and seal and label second bag(s). Repeat
previous two steps.
8. Place double-bagged sample(s) in cooler with dry ice until shipment.
9. Ship weekly on dry ice by Federal Express next day service. KEEP FROZEN UNTIL
SHIPMENT. If ice is used, double bag ice in self-sealing plastic bags and tape shut to avoid
contamination of samples if ice should melt.
*** - If both primary and secondary target species are collected, the two samples should be wrapped
and bagged separately, with separate bar codes and labels, but only one Sample Tracking Form.
B-31
-------
-------
APPENDIX C
FIELD DATA FORMS
Copies of field data forms are arranged according the general order of their use at
each stream site:
1. Verification Form
2. Sample Collection Form
3. Field Measurement Form
4. Channel/Riparian Cross-Section & Thalweg Profile Form
5. Slope and Bearing Form
6. Vertebrate Collection Form
7. Vertebrate Length Recording Form
8. Rapid Habitat Assessment Form (Riffle/Run Prevalent)
9. Rapid Habitat Assessment Form (Pool/Glide Prevalent)
10. Assessment Form
Electronic versions of the forms may be available through the EMAP-Surface Waters
Technical Director, U.S. EPA, 200 SW 35th St, Corvallis, OR 97333.
C-1
-------
-------
Reviewed by (initial)".
VERIFICATION J=ORM - STREAMS/RIVERS
SITE NAME:
DATE:
VISIT: D1 D2
SITE ID:
TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
STREAM/RIVER VERIFICATION INFORMATION
STREAM/RIVER VERIFIED BY (X all that apply):
Q OTHER (DESCRIBE HERE):
GPS I I LOCAL CONTACT fl SIGNS I I ROADS | | TOPO. MAP
Q NOT VERIFIED (EXPLAIN IN COMMENTS)
COORDINATES
LATITUDE (dd mm ss} North
LONGITUDE (ddd mm ss) West
TYPE OF GPS FIX
Are GPS ,
Coordinates
w/I10 Sec. of map?
MAf:
GPS:
|2D
I 3D
LJ YES
D NO
INDEX SITE STATUS - X ONE BOX FROM ONE SECTION ONLY
SAMPLEABLE
I I REGULAR-WADEABLE
I I REGULAR-NOTWADEABLE
I I INTERMITTENT - DRY SPOTS ALONG REACH
|~l DRY - No WATER ANYWHERE ALONG REACH
I I ALTERED - STREAM/RIVER PRESENT BUT NOT AS ON MAP
I I OTHER (EXPLAIN IN COMMENTS)
NON-SAMPLEABLE (No SAMPLE TAKEN)
I I No CHANNEL OR WATERBODY PRESENT
Q IMPOUNDED (UNDERNEATH LAKE/POND)
|~| WETLAND (NO DEFINABLE CHANNEL)
NO ACCESS
I I ACCESS PERMISSION DENIED
I | INACCESSIBLE (UNABLE TO REACH SITE)
DiREGtlONSTOSTREAJ¥i/R8VER SITE
GENERAL COMMENTS
RECORD INFORMATION USED TO DEFINE LENGTH OF REACH, AND SKETCH GENERAL FEATURES OF REACH ON REVERSE SIDE.
Rev. 06/02/97 (strweri.97) VERIFICATION FORM - STREAMS/RIVERS - 1
-------
Reviewed by (initial):.
ID: - TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
VERIFICATION FORM - STREAMS/RIVERS tcbntinued)
SITE NAME:
DATE:
VISIT: D1 D2
STREAM/RIVER REACH DETERMINATION
CHANNEL VWDTH USED TO DEFINE
REACH {M} (XX):
DISTANCE (M) FROM X-srre
UPSTREAM LENGTH DOWNSTREAM LENGTH^
COMMENT
Rev. 06/02/97 (strweri.97)
VERIFICATION FORM - STREAMS/RIVERS - 2
-------
Reviewed by (initial);
SAMPLE COLLECTION FORM - STREAMS
SITE NAME: DATE: / / VISIT: D1 D2
SITE ID: TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
COMPOSITE BENTHOS SAMPLES ' ,
SAMPLE ID
(BARCODE)
HABITAT No ~
^,°N?} OF FLAG " " ' CoMMru-n
R P JARS —
»
STATION A
RIFFLE PR POOL
• -0C ONE)"
LEFT, CENTER, OR .
RIGHT-
B C D E F G, H
DR DR DR DR DR DR DR
DP DP DP DP DP DP DP
DL DL DL DL DL DL DL
DC DC DC DC DC DC DC
DR DR DR DR DR DR DR
1 J K
DR DR
DP DP ,
DL DL
DC DC . '
DR DR
COMPOSITE PERIPHYTON SAMPLES HABITAT TYPE (X)- n RIFFLE DPOOL D OTHER
SAMPLE ID (BARCODE) -
ASSEMBLAGE ID
, (50-ML TUBE)
- SUB. SAMPLE VOL.
ML
COMPOSITE VOLUME- ML
CHLOROPHYLL BIOMASS
(GF/F FILTER) (TARED FILTER)
VOL. FILTERED FILTER No. " VOL. FILTERED
ML ML
APA SAMPLE
(50-ML TUBE)
SUB. SAMPLE VOL". *
ML
COMPOSITE PERIPHYTON SAMPLES • - '' HABITAT TYPE (X)- D RIFFLE d POOL D OTHER
; SAMPLE ID (BARCODE) -
ASSEMBLAGE ID
(50-ML TUBE)
SUB. SAMPLE VOL.
ML
COMPOSITE VOLUME- ML
CHLOROPHYLL „ BIOMASS
(GF/F FILTER) (TARED FILTER)
VOL, FILTERED FILTER No. VOL. FILTERED .
ML ' ML
'COMMENTS:,,
APA SAMPLE
(50-ML TUBE) '
, - SUB. SAMPLE VOL.
ML
Flag codes: K= Sample not collected; U= Suspect sample; F1, F2, etc.= misc. flag assigned by field crew. Explain all flags in Comment sections.
Rev. 06/02/97 (st_saco.97) SAMPLE COLLECTION FORM - STREAMS - 1
-------
Reviewed by (initial):.
SAMPLE COLLECTION FORM - STREAMS (continued}
SITE NAME:
DATE: / / VISIT: D1 D2
i " — TEAM.D(X): D1 D2 D3 D4 D5 D6 D7
M,CRQBIAL WATER SA^
ICHEMISTRY.
SAMPLE ID (BARCODE)
TRANSECT
FLAG
COMMENTS
SPPijyifeNTf OXlClTY SAMPLES
SAMPLE IP (BARCODE)
FLAG
COMMENTS
FISH TISSUE SAMPLES - PRIMARY SAMPLE (min. SOg total wgt)
available; 5 [individuals)
SAMPLE ID (BARCODE) -
Is COMPOS^ ^lUPlM COMPOSED OF INDIVIDUALS COLLECTED FRdP THROUGHOUT REACH? (X) ~
FISH TISSUE SAMPLES - SECONDARY SAMP' »=
SAMPLE »D (BARCODE)-
NUMBER OF
INDWIDUALS
FLAG
D YES D No
TOTAL LENGTH (MM)
FLAG
COMPOSITE SAMPLE COMPOSED OF INDIVIDUALS COLLECTED FROM THROUGHOUT REACH? (X)-»
D YES D No
IF No, EXPLAIN:
COMMENT OR FLAG EXPLANATION FOR FISH TISSUE
FlaB co<.e8: K* Samp.e not conactcd; U= Suspect sarnp.e; F1. F2. etc.= misc. flag assigned by fie.d crew. Exp.ain a,, flags in Comments sections.
SAMPLE COLLECTION FORM - STREAMS - 2
Rev. 06102137 (st_saco.97)
-------
Reviewed by (initial):
FIELD MEASUREMENT FORM - STREAMS/RIVERS
SITE NAME:
DATE:
VISIT: D1 D2
SITE ID: - . TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
WEATHER CONDITIONS i
CLOUD COVER
^PRECIPITATION
PREVIOUS PRECIPITATION (24 H)'
AIR TEMPERATURE XX
n
DNONE
DNONE
D 5-25%
D LIGHT
D LIGHT
D 25-50%
D MODERATE
D MODERATE
D 50-75%
D HEAVY
D HEAVY
D >75%
IN SITU MEASUREMENTS
STATION ID:
Assume X-site unless marked
FLAG
COMMENTS
QCCS CONP ^S/C
STREAM/RIVER COND /J.S/CM
STREAM/RIVER DO MG/L
STREAM/RIVER TEMP "C
STREAM/RIVER METABOLISM DETERMINATION
INITIAL O2
(MG/L)
INITIAL
INCUBATION
.TEMP.(°C)
INCUBATION TIME
(24-HR TIME)
START
(HH:HM)
FINISH
(HH:M»)
DURATION OF
INCUBATION
(HH:MM)
FLAG
COMMENTS
SAMPLE ID
(BARCODE}
FINAL O2
(MG/L?
FLAG
COMMENTS
- OXYGEN METER CALIBRATION INFORMATION
MEMBRANE CHECK
ELECTRONIC ZERO
CALIBRATION CHA? SBER TEMPERATURE:
SATURATED O2 @ TEMP.;
MG/L
STATJON ELEVATION (FROM TOPO. MAP OR ALTIMETER):
FT
ELSVATTON CORRECTION FACTOR:
Tha callbraHon value Is obtained by multiplying the saturated DO concentration times
an elevation correcsioa factor (o^salnad front ths tables on the back of iha YSI meter).
Adjust the meter reading to Die calibration value. ,
CALIBRATION VALUE:
COMMENTS:
Flag Codes: K = no measurement or observation made; U= suspect measurement or observation; Q = unacceptable QC check associated with
measurement; F1, F2, etc. = miscellaneous flags assigned by each field crew. Explain all flags in comments section.
Rev. 06/02/97 (strvfldm.97)
FIELD MEASUREMENT FORM - STREAMS/RIVERS -1
-------
Reviewed by (initial):,
FIELD "MEASUREMiNY FORM'TiflEAMSloopniiiiii)''
SITE NAME:
DATE:
/ / VISIT: D1 D2
SITE ID: _- TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
STREAM DISCHARGE"
VELOCITY AREA
Sl"
12
1S
DlST.FROW
BANK (CM)
VELOCITY
(FT/S)XXJC
DEPTH
(FEET) XX-X
FLAG
TIMED FILLING
REPEAT
VOL. (L) xx.x
TiME(S)
FLAG
Q NEUTRALLY BUOYANT OBJECT
MEASUREMENT
WIDTH (m)
DEPTH 1 (cm)
DEPTH
cm)
DCPTH 4 (cm)
i i i i,
V it Cross Section
cW'/-
THREE
17
:F1AG
5 (cm)
FLOAT
DISTANCE (m)
TIMEJS)
ssssssssss
COMMENTS
•laa Codos: K = no measurement or observation made; U = suspect measurement or observation; Q = unacceptable QC check associated
with measurement; F1, F2, etc. - miscellaneous flags assigned by each field crew. Explain all flags in comments section.
Rev. 06/02/97 (strvfldm.97)
FIELD MEASUREMENT FORM - STREAMS/RIVERS - 2
-------
Reviewed by (initiatt;
FIELD MEASUREMENT FORM - STREAMS (continued)
SITE NAME: DATE: / / V|S|T. D1 n2
SITE ID: TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
STREAM DISCHARGE , . ,
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
Iff
19
20
FL
LZI VELOCITY AREA
DlST, FROM
BANK (CM)
— j
AG
VELOCITY
(M/S)XX.X
m
.
•
m
.
DEPTH
(CM)XX.X
FLAG
n TIMED FILLING
REPEAT
1
2 .
3',
- 4 ~'J
-> - 5 - -
VOL. (L) xx.x
TlME(S)
FLAG
H] NEUTRALLY BUOYANT OBJECT
, MEASUREMENT
». f /•
WIDTH (m)
DEPTH 1 (cm)
DEPTH 2 (cm)
i- DEPTH 3 (cm)
DEPTH 4 (cm)
DEPTH 4 (cm)
>, FLOAT
DISTANCE (m)
FLOAT *
TIME(S)
•— "~~-*
COMMENTS
^ .'Cross Section
ONE
TWO
.~a
* THREE
•5B55iBSSS5BS55
Flag Codes: K = no measurement or observation made; U = suspect measurement or observation; Q = unacceptable QC check associated
with measurement; F1, F2, etc. = miscellaneous flags assigned by each field crew. Explain all flags in comments section.
Rev. 06/02/97 (strvfldm.97)
FIELD MEASUREMENT FORM - STREAMS/RIVERS - 3
-------
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-------
-------
Reviewed by (initial);
^PlPi^j^isiSESilVIENT FORM: RIFfLE/RUMPREVALENCE - STREAMS
SITE NAME: DATE: / / 97 VISIT: D1 O2
SITE ID: MAIA97- TEAM ID (X): Ol D2 D3 D4 D5 D6 D? D8
TOTAL SCORE ||
1. INSTREAM COVER
(FISH)
SCORE: ||
2. EPIFAUNAL
SUBSTRATE
SCOR&Jl
3. EMBEDDEDNESS
t.
~ SCORE:J|
4. VELOCITY/DEPTH
' REGIMES
1 ' SCORE: |
5. CHANNEL
ALTERATION
SCORE^j|
f-
6, SEDIMENT
DEPOSITION
n ' ,'
- SCORE: I)
CATEGORY
ODTIMSI
Greater than 50% mix of
boulder, cobble,
submerged logs,
undercut banks, or
other stable habitat.
20 19 18 17 1B
Well-developed riffle
and run; riffle is as wide
as stream and its length
extends two times the
width of stream;
abundance of cobble.
20 19 18 17 16
Gravel, cobble, and
boulder particles are
between 0 and 25%
surrounded by fine
sediment
i
20 19 18 17 16
All four velocity
regimes are present
(slow-deep, slow-
shallow, fast-deep, fast-
shallow),'
20 19 18 17 1®
No channelization of
dredging present., ,
! j - „ i
1 *
,/*
' '.',',-•
20 19 18 ,17 1&
Little or no enlargement
of islands or point bars
and less than 5% Of the
bottom Is affected by
sediment deposition.
f i
'.' r 'I -J *•
i ' j ' ' 1 1 1>
20 19 18 17 16
Si IR-OPTIM Al
30 to 50% mix of boulder,
cobble, or other stable
habitat; adequate habitat
A t
15 14 13 12 11
Riffle is as wide as stream,
but is less than two times
width; abundance of cobble;
boulders and gravel ,
common. *" , '' Vi% '
;, '> ••' r- ^
15 14 13 12 11
Gravel, cobble, and boulder
particles are between 25 and
50% surrounded by fine '
sediment „ "** f?
~ "' *rti ' D \t% > *E
'< IiiV ?-i t *!" v i '<
15 14 13 12 11
Only three of the four habitat
types are present (if fast-
shallow is missing, score
lower than if other regimes,
are missing). '
f - <* ' ' i >t
15 14 13 12 11
Some channelization is
present usually In areas of
bridge abutments; evidence
of past channelization, i.e.,
dredging (greater than past,
20 yr) may be present, but •
recent channelisation Is, not
jsresent. , i f
15 14 13-12 'ft ,
Some new Increase In bar '
formation, mostly from' •-
coarse gravel; 5 to 30% of
the bottom is affected; slight
deposition In pools. * ( .
i , ." ! ,B tfiif \ s.L
*~ ' * 4 * > 1 **• n * I^T M*^
„ • jK^^'^«(|ift(ri||
. ;/Avwi^l
15 14 is ia,,^,-
lUfAPfSIMAt
10 to 30% mix of boulder,
cobble, or other stable
habitat; habitat availability is
less than desirable. t
' ' , j; *•
10 9 8 7 6
Run area maybe lacking;
reduced riffle area that does
not extend across entire
cross section and is less
than two times the width;
gravel of large boulders and
bedrock prevalent; cobble
present '"
10 9 8 -7 6,,
Gravel, cobble, and boulder
particles are between 50 and
75% surrounded byflna
sediment .-
' .' , ' u 1. 1 l''i'i«i
. ..' '-, ', ?- *}*#!&
10 8 87 6 __,
Only two of the four habitat
types are present (if fast*
shallow or slow-shallow are
missing, score lowi. i ",,,,'
- :'.'"i;;\>,*'
10 9 '. 8 • 7 S
Mew embankments are :
present on both banks; and
40 to 80% of the stream '•
reach is channelized and
disrupted. ' " ! "gfi*
»* '.IL!-, '0€jft-
i .« i , " i "- '{!•
i i f '« ' *• . n'if wr
10 X 07 6, ,'r
Moderate deposition of new
jravsl or coarse sand on old
and new bars; 30 to 50% of
the bottom is affected; « H
sediment deposits at - > ,••
obstructions, constrictions,
and bends; moderate, , t,fc
deposition of pools ' ,",t*
prevalent * " T -f ,
10 ,9 8 7^6,;,
P/SAI?
Less than 10% of boulder,
cobble, or other stable
habitat; lack of habitat is
obvious. < ,' ,( -,
-.' t '( ,
5 4 3 2 1, 0
Riffles or run virtually non-
existent; gravel or large
boulders and bedrock
prevalent; cobble lacking.
" ' - '' ,- !, f, ' I, ,
' i ,' ?. (
' *• ««!** ,*T-*i J
5 4 3 2 1^,0
Gravel, cobble, and boulder
particles are over 75% .
surrounded by fine
sediment , 's * *,
1 ' 'i -r-' A ft
- » . -S ' *
5 4 3' 2, 1 ,0
Dominated by ona
velocity/depth regime <
(usually slow-deep).
^v'^^s'i^-i''"
\ • ' ,J ->• -
5 4 3 2' 1 0
Banks shored with gabion
or cement; over 80% of the
stream reach is Channelized
and disrupted. '
h", ,-"'(',;! TJf^i^^rffp
^^^'.'V-iifS^f
5 4. 3 2 1 0 „
Heavy deposits of fine ,
material; Increased bar >
development; more than
50% of the bottom Is > , .
changing frequently; pools
almost absent due to
substantial sediment, . „ "
deposition., tjl,'t^m
1 1 1 Ml * 1 , I , ,¥U V ,*(,
5 4,3 2 1, 0^
Rev. 06/02/97 (stxxrhrr.97)
RAPID HABITAT ASSESSMENT FORM: RIFFLE/RUN - STREAMS - 1
-------
Reviewed by (initial):
RAPID HABITAT ASSESSMENT FORM:
SITE NAME:
DATE:
VISIT: D1 D2
SITE ID: - TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
fHABttAY PARAMETER
7. FREQUENCY OF
, RIFFLES
SCORE:
:; =
8, CHANNEL FLOW
l""]| I i "i| "
lii SCORE; I
CATEGORY
9. CONWnONOF
BANKS
SCORE:
SCORE:
n x:,.:;),;i!iis;,; I,,::1:,,,,!:,;:!,:; :., ;; ;
aNCK^SrHER
SRUPTIVE
3RESSURE
i SCORE: I
Occurrence of riffles Is
relatively frequent; the
distance between riffles
divided by the width of
he stream equals 5 to
; variety of habitat
Jccurrence of riffles Is
nfrequent; distance between
iffles divided by the width of
he stream equals 7 to 15.
20 19 18 17 16
Water reaches the base
of both banks and a
minimal area of channel
substrate is exposed.
20 19 18 17 16
Banks stable; no
evidence of erosion or
bank failure.
20 19 18 17 16
Mora than 90% of the
stream bank surfaces
are covered by
vegetation.
20 19 18 17 16
__==================
Vegetative disruption,
through grazing or
mowlna is minima! or
through grazing or
"ng is minima! <
not evident; almost all
plants are allowed to
grow naturally.
Width of riparian zone Is
greater than 18m;
human activities fl.e.;
parking lots, roadbeds,
clearcuts, lawns, or
crops) have not
Impacted this zone.
Occasional riffle or bend; ,
bottom contours provldu
some habitat; distance
between riffles divided by,
he width of the stream Is
between 15 to 25.
15 14 13 12 11
Vater fills more than 75% of
he available channel; or less
than 25% of the channel
substrate is exposed.
15 14 13 12 11
Banks moderately stable;
Infrequent, small areas of
erosion mostly healed over.
15 14 13 12 11
70 to 90% of the stream bank
surfaces are covered by
vegetation., .
15 14 13 12 11
Disruption is evident but Is
not affecting full plant
growth potential to any great
extent; more than one-half of
the potential plant stubble
height remaining.
Zone width is between 12
and 18 m*, human activities
have only minimally
impacted this zone.
Senerally all flat water or
shallow riffles; poor habitat;
distance between riffles
divided by the width of the
stream is greater than 25.
10 9 8 7
Water fill 25 to 75% of ttie
available channel; and/or
riffle substrates are moirtty
exposed.
10 9 8 7 (5,
Moderately unstable; up to
60% of banks irt reach rwe
areas of erosion.' "> •><
10 9 8 7
50 to 70% of the stream •
bank surfaces are covered
by vegetation. >
10 9 8 7 B
Disruption is obvious;
patches of bare soil or
closely cropped vegetation
are common; less than
one-half of the potential
plant stubble height
remaining,
6
Zone width is between 6
and 12 m; human activ ties
have Impacted the zonii a
great deal: ,>( ,
543210
fery little water in channel,
ana mostly present as
standing pools.
543210
Jnstable; many eroded
areas; "raw" areas frequent
along straight sections and
bends; on aide slopes, 60 to
100% of bank has erosional
scars. <
543210
Less than 50% of the stream
bank surfaces are covered
by vegetation.
543210
Disruption of stream bank
vegetation is very high;
vegetation has been
removed to 2 inches or less
In average stubble height.
Width of zone Is less than 6
m; little or no riparian
vegetation due to man-
induced activities.
Rev. 06/02/97 (stxxrhrr.97)
RAPID HABITAT ASSESSMENT FORM: RIFFLE/RUN - STREAMS - 2
-------
Reviewed by (initia)):.
RAPID HABITAT ASSESSMENT FORM: GLIQE/POOL PREVALENCE - STREAMS
SITE NAME: DATE: / / VISIT: D1 Q2
SITE ID: _ . - : TEAM ID (X): ni D2 D3 D4 D5 D6 D7 D8
TOTAL ||
HABITAT PARAMETER
1. INSTREAM COVER
SCORE: ||
2. EPIFAUNAL.
SUBSTRATE
SCORE: ||
3. POOL SUBSTRATE
CHARACTERIZATION
SCORED
4. POOL VARIABILITY
SCORE: ||
5. CHANNEL ALTERATION
SCORE: ||
6. SEDIMENT
DEPOSITION
SCORE: |
CATI
DDTIMAI
Greater than 50% mix of
snags, submerged logs,
undercut banks, or other
stable habitat; rubble or
gravel may be present.
20 19 18 17 16
Preferred benthic
substrate (to be sampled)
is abundant throughout
stream site and at a
stage to allow for full
colonization potential
(i.e.; logs ana snags that
are not new fall and not ,
transient
20 19 18 17 16
Mixture of substrate
materials, with gravel
and firm sand prevalent;
root mats and
submerged vegetation
are common.
20 19 18 17 16
Even mix of large-
shallow, large-deep,
small-shallow, and small*
deep pools are present.
20 19 18 17 , 16
No channelization of
dredging present
20 19 18 17 16
Less than 20% of the
bottom is affected; minor
accumulation of fine and
coarse material at snags
and submerged
20 19 18 17 16
Sl IR-TlDTIMAI
30 to 50% mix of stable '
habitat; adequate habitat for
maintenance of populations.
15 14 13 12 11
Substrate is common but is
not prevalent nor well-suited
for full colonization , „, „
potential. * ,Vv
i ** ~ *' t
« s < i r;
1 " . r ' .- >," "r'Y,
: >. ' .' • > ' _,v
15 14 13 12 Tt
Mixture of soft sand, mud,
or clay; mud maybe
dominant; some root mats
and submerged vegetation
are present „ •|V
, <. i", -»<*' .,,,-'
15 14 13 12 11 ,
The majority of pools are
large and deep; very few
shallow. __*,_,
' f , " 1 ! . , "'
< 0 ~ ' f
15 14 13 12 11
Some channelization is
present, usually In areas of
bridge abutments; evidence
of past channelization, I.e.;
dredging (greater than past
« ., ',!>,'• I
15 14,13 12,11,
20 to 50% affected;
moderate accumulation;
substantial sediment
movement only during major
storm events; some new , /
< i "i i f i* M
15 14 13 12 ,11
zrsnisv
MADRIMAI
10 to 30% mix of stable
habitat; habitat availability
is less than desirable. t
" ,- * " ' •"
r (,
, 10 9 8 7 6
Substrate frequently
disturbed or removed.
; : i ' ""'
""%,',>
^ . ' M , , , A,. I
'.'*•.„• :*;ri>.y,
i '*j
• , • ' , ,. - - r o
10 9 ' 8 7 ' 6 ..
All mud or clay or sand
bottom; tittle or no roof ,
mat; no submerged
vegetation. ' --,
1 , f < .*.)* ,,'. ,\
•. * !. ( 1| .1.
10 9 8 7 6 ..
Shallow pools much more
prevalent than deep pools.
. T,' : , "-V1
"„ ",1"",;
t \ •„ ' i 'i*;1** *,
10 9 8 76,
New embankments are
present on both banks;
channelization maybe
extensive, usually in urban
areas or drainage areas of
*i ^- *< >, 5 i ^ >.
10 , 9 8 7 ± 6
50 to 80% affected; major
deposition; pools shallow
and heavily silted;
embankments may be
present on both banks;
r ' L W
10 9 8 7 6
Pnno
Less than 10% stable
habitat; lack of habitat is
obvious.
t i ^ *
1 V , , , , ^ •<
• • ' <- />,t"
543210
Substrate is unstable or
lacking. : - ,
I , -. i v. >\ -','
, > ,"-^, J^.^
'.'t ,i,,' „
- '"«;- -'V^tX
i h JH«, i '^l !«'<
' . I < 1 1'" M 'i "fciK <*
543210
Hard-pan clay or bedrock;
no root mat or vegetation. ,
i 0 f ** ,i*
r'-A1,.!',!;'-:
i * "'" ' 'i
« 5, 4 3 2, 1 ,0
Majority of pools are small-
shallow or pools are absent.
* ? ?» t(-c \^\ EJ4JM
( ,. ' I . -v^.j*. Kijt
543210,
Extensive channelization;
banks shored with gabion
or cement; heavily
urbanized areas; mstream
habitat greatly altered or ' *
^ H ' "
543210
Channelized; mud, silt,
and/or sand in braided or
non-braided channels;
pools almost absent due to
deposition. •. | , _
5 4 3 2 1 ,0
Rev. 06/02/97 (stxxrhgp.97)
RAPID HABITAT ASSESSMENT FORM: GLIDE/POOL - STREAMS -1
-------
Reviewed by (initial):
^j|t.!;:G^IPE/PQO^i^iWS;.(cionttnued)^
VISIT: D1 D2
rEAMID(X): D1 D2 D3 d4 D5 D6 D7 D8
Io'W|lriWiaW;if|;i¥;
ililStlllfiP
^iiiii:iiil|i|i:i
Veiy llttfe wat^tlrt channel
of both lovifM- banks and
20 "19 18 17 16
; no';.
evidence of sroslwi oft
t«ife|iHpijS^|>fth^stream
•" •'••'•" '-•J- --"ka-cdyered;.!1
raMHMMWjW
almost all plants are
allowed to grow
V>t lsra»Tr'*T^! "fw^.^TTTT^; ,:;:
halflht remaining.
543210
lass than 6 meters; little or
no riparian vnontation due
eatw than 18 meters;
wv*<«>r» ^MtulffcuB IF^L*'1:,;;.
human —^T,.™--, ,,,_._.-,
minimally impacted this
aBOIw:.'.":."'"!/! '..•vi'.^i,;'!,'::"1.
^'l.l;T1"w'~;
io'hunian^tnwiltiBfJif;;:1^. i'.:
ieiit-^yh^^li^fsiM:-!;;
Ji^'-H/i^J-^p^*:!':!!1^"*'-'-'
fitiilil•>!;'>..;.'-•|3ft:wl;.i!:^!I;:;-:i;:M >''. -:
R«v. 06/02/97 (stxxrhgp.97)
RAPID HABITAT ASSESSMENT FORM: GLIDE/POOL - STREAMS - 2
-------
Reviewed by (initial):
ASSESSMENT FORM - STREAMS/RIVERS
SITE NAME:
DATE:
VISIT: Dl D2
SITE ID: ' - TEAM ID (X): D1 D2 D3 D4 D5 D6 D7 D8
WATERSHED ACTIVITIES AND DISTURBANCES OBSERVED (INTENSITY: BLANK=NOT OBSERVED, L=Low, IMMODERATE, H=HEAW)
RESIDENTIAL
RECREATIONAL
AGRICULTURAL
INDUSTRIAL
STREAM MANAGEMENT
RESIDENCES
PARKS. CAMPGROUNDS
CROPLAND
INDUSTRIA
ING_
MAINTAINED LAWNS
PRIMITIVE PARKS. CAMPING
PASTURE
MINES/QUARRIES
DRINKING WATER TREATMENT
CONSTRUCTION
TRASH/LITTER
LIVESTOCK USE
OiLtaAS WELLS
ANGLING PRESSURE
PIPES. DRAINS
SURFACE FILMS. SCUMS. OR SLICKS
ORCHARDS
POWER PLANTS
DREDGING
DUMPING
POULTRY
LOGGING
CHANNELIZATION
IRRIGATION PUMPS
EVIDENCE OF FIRE
WATER LEVEL FLUCTUATIONS
BRIDGE/CULVERTS
ODORS
FISH STOCKING
BREACH CHARACTERISTiGS (peircent of reach)
FOREST
SHRUB
- 'GRASS
1 WETLAND
BARE GROUND
MACROPHYTES
]RARE( 75%)
EXTENSIVE (> 75%)
EXTENSIVE (> 75%)
EXTENSIVE (> 75%)
EXTENSIVE (> 75%)
EXTENSIVE (> 75%)
AGRICULTURE - Row CROP
AGRICULTURE- GRAZING
- : LOGGING
DEVELOPMENT (RESIDENTIAL & URBAN)
~|RARE(<5%)
SPARSE (5 TO 25%)
MODERATE (25 TO 75%)
EXTENSIVE (> 75%)
I RARE (< 5%)
JRARE(<5%)
SPARSE (5 TO 25%)
MODERATE (25 TO 75%)
EXTENSIVE (> 75%)
SPARSE (5 TO 25%)
MODERATE (25 TO 75%)
RARE (< 5%)
SPARSE (5 TO 25%) -
MODERATE (25 TO 75%)
EXTENSIVE (> 75%)
EXTENSIVE (> 75%)
WATER CLARJTY
HlftH! VTllRRln
"EP
-r-pr
PRISTINE
TF"
~TF
HIGHLY
APPEALING
TT2-
TTT
GENERAL ASSESSMENT (wildiife, vefietation diversity, forest age ciass (0-25 yrs. 25-75 yrs, >75),
LOCAL ANECDOTAL INFORMATION*
Rev. 06/02/97 (strvasse.97)
ASSESSMENT FORM - STREAMS/RIVERS - 1
-------
r
ASSESSMENT FORM - STREAMS/RIVERS (continued)
Rav. 06/02«7 (strvasse.97)
ASSESSMENT FORM - STREAMS/RIVERS - 2
-------
APPENDIX D
SPECIES CODES FOR AQUATIC VERTEBRATES:
MID-ATLANTIC REGION
The following table contains the unique 6-character species code, the scientific
name, and the common name assigned to each aquatic vertebrate species expected to be
collected by EMAP sampling protocols in the Mid-Atlantic region. Generally, the species
code is composed of the first four letters of the genus plus the first two letters of the species
name. Modifications to this coding scheme were made in cases where two species could
be assigned the same code. Species entries are arranged first by family (alphabetically),
then by the assigned species code.
Similar lists have been compiled for EMAP-related studies occurring in regions other
than the Mid-Atlantic. Information regarding the availability of species codes and associated
information may be obtained through the EMAP-Surface Waters Technical Director, c/o
U.S. EPA, 200 SW 35th St., Corvallis, OR 97333.
D-1
-------
EMAP-SW-Streams Field Operations Manual. Appendix D. Rev. 0. September 1998 Page 2 of 10
SPECIES LIST AND STATUS FOR MID-ATLANTIC REGION
ICHTBD
ICHTFO
ICHTGR
ICHTUN
ILAMPAE
ILAMPAP
JPETRMA
ILAMPZZ
ACIPBR
I
ACIPFU
SCAPPL
POLYSP
LEPIOC
LEPIOS
ILEPIPL
IAMIACA
IANGURO
IAPHRSA
I LABIS!
IMENIBE
CARPCA
CARPCY
CARPVE
CATOCA
ICATOCO
ICYCLEL
IERIMOB
IERIMSU
[HYPENI
[HYPERO
Latin Name
chthyomyzon bdellium
chthyomyzon fossor
chthyomyzon greeleyi
chthyomyzon unicuspis
ampetra aepyptera
ampetra appendix
Petromyzon marinus
Acipenser brevirostrum
Acipenser fulvescens
Scaphirhynchus platorynchus
Polyodon spatula
Lepisosteus oculatus
Lepisosteus osseus
Lepisosteus platostomus
Amia calva
Anguilla rostrata
Aphredoderus sayanus
Labidesthes sicculus
Menidia beryllina
Carpiodes carpio
Carpiodes cyprinus
Carpiodes velifer
Catostomus catostomus
Catostomus commersoni
Cycleptus elongatus
Erimyzon oblongus
Erimyzon sucetia
Hypentelium nigricans
Hypentelium roanokense
Dhio Lamprey |
orthern Brook Lamprey
Mountain Brook Lamprey
ilver Lamprey
east brook lamprey
American brook lamprey
Sea lamprey
Unknown lamprey
Atlantic sturgeon
Lake sturgeon
Shovelnose sturgeon
Paddlefish
Spotted gar
Longnose gar
Shortnose gar
Bowfin
American eel
Pirate perch
Brook silversides
Inland silverside
River carpsucker
Quillback
Highfin carpsucker |
Longnose sucker |
White sucker |
Blue sucker |
Creek chubsucker |
Lake chubsucker |
Northern hogsucker |
Roanoke hogsucker |
D-2
-------
EMAP-SW-Streams Field Operations Manual, Appendix D, Rev. 0. September 1998 Page 3 of 10
CODE Latin Name
ICTIBU
ICTICY
MINYME
MOXOAN
MOXOAR
MOXOCA
MOXOCE
MOXODU
MOXOER
MOXOHA
MOXOMA
MOXOPA
MOXORH
MOXORO
MOXOVA
CATOZZ
ACANPO
AMBLCA
AMBLRU
ARCHIN
CENTMA
ENNECH
ENNEGL
ENNEOB
LEPOAU
LEPOCY
LEPOGI
LEPOGU
LEPOMA
LEPOME
Ictiobus bubalus
Ictiobus cyprinellus
Minytrema melanops
Moxostoma anisurum
Moxostoma ariommum
Moxostoma carinatum
Moxostoma cervinum
Moxostoma duquesnei
Moxostoma erythrurum
Moxostoma hamiltoni
Moxostoma macrolepidotum
Moxostoma pappillosum
Moxostoma rhothoecum
Moxostoma robustum
Moxostoma valenciennesi
Ancartharcus pomotis
Ambloplites cavifrons
Ambloplites rupestris
Archoplites interruptus
Centrarchus macropterus
Enneacanthus chaetodon
Enneacanthus gloriosus
Enneacanthus obesus
Lepomis auritus
Lepomis cyanellus
Lepomis gibbosus
epomis gulosus "
epomis macrochirus
epomis megalotis
Common Name
Smallmouth buffalo
Bigmouth buffalo
Spotted sucker
Silver redhorse
Bigeye jumprock
River redhorse
Black jumprock
Black redhorse
Golden redhorse
Rustyside sucker
Shorthead redhorse
V-lip redhorse
Torrent sucker
Smallfin Sucker
Greater Redhorse
Unknown catostomid
Mud sunfish
Roanoke rockbass
tockbass
Sacramento perch
Flier
Blackbanded sunfish
Bluespotted sunfish
Banded sunfish
Redbreast sunfish
Green sunfish
Pumpkinseed
i/Varmouth
Bluegill
ongear sunfish
D-3
-------
EMAP-SW-Streams Field Operations Manual. Appendix D. Rev. 0. September 1998 Page 4 of 10
1 1
CODE 1
L=— =====
LEPOM1
I
JMICRDO
MICRPU
MICRSA
POMOAN
IPOMONI
CENTZZ
ALOSCH
ALOSPS
ALOSSA
IDOROCE
ICOTTBA
JCOTTBL
IJCOTTCA
ICOTTCF
ICOTTCO
ICOTTGI
ICOTTGU
ICOTTMA
ICOTTRI
JcOTTTE
IJcOTTZZ
ICAMPAN
JCLINEL
ICLINFU
JcOUEPL
ICYPRAN
ICYPRGA
ICYPRMO
UCYPRSP
Latin Name
===========================
epomis microlophus
Micropterus dolomieu
Micropterus punctulatus
Micropterus salmoides
Pomoxis annularis
Pomoxis nigromaculatus
Alosa chrysochloris
Alosa pseudoharengus
Alosa sapidissima
Dorosoma cepedianum
Cottus bairdi
Cottus baileyi
Cottus carolinae
Cottus confusus
Cottus cognatus
Cottus girardi
Cottus gulosus
Cottus marginatus
Cottus rice;
Cottus tenuis
Campostoma anomalum
Clinostomus elongatus
Clinostomus funduloides
Couesius plumbeus
Cyprinella analostana
Cyprinella galactura
Cyprinella monacha
Cyprinella spiloptera
Common Name
Redear sunfish
mallmouth bass
"potted bass
argemouth bass
White crappie
Black crappie
Unknown centrarcid
Skipjack herring
Alewife (landlocked)
American shad
Gizzard shad
Mottled sculpin
Black sculpin
Banded sculpin
Shorthead sculpin
Slimy sculpin
Potomac sculpin
Riffle sculpin
Margined sculpin
Spoonhead sculpin
Slender Sculpin II
Unknown cottid
Stoneroller
Redside dace
Rosyside dace
Lake chub
Satinfin shiner
Whitetail shiner
D-4
-------
EMAP-SW-Streams Field Operations Manual, Appendix D, Rev. 0. September 1998 Page 5 of 10
CYPRWH
ERIMCA
ERIMDI
ERIMIN
ERIMXP
IEXOGLA
EXOGMA
HYBOHA
HYBORE
LUXIAL
LUXICC
LUXICE
LUXICH
LUXICR
LYTHAR
LYTHLI
LYTHUM
MACRAE
MACRST
MARGMA
NOCOBI
INOCOLE
NOCOMI
NOCORA
NOTECR
NOTRAB
NOTRAL
NOTRAN
NOTRAQ
NOTRAR
Latin Name
Cyprinella whipplei
Erimystax cahni
Erimystax dissimilis
Erimystax insignis
Erimystax x-punctatus
Exoglossum laurae
Exoglossum maxillingua
Hybognathus hankinsoni
Hybognathus regius
Luxilus albeolus
Luxilus coccogenis
Luxilus cerasinus
Luxilus chrysocephalus
Luxilus cornutus
Lythurus ardens
Lythurus lirus
Lythurus umbratilus
Macrhybopsis aestivalis
Macrhybopsis storeriana
Margariscus margarita
Nocomis. biguttatus
Nocomis leptocephalus
Nocomis micropogon
Nocomis raneyi
Notemigonus crysoleucas
Notropis amblops
Notropis alborus
Votropis anogenus
otropis amoenis
otropis ariommus
Common Name
Steelcolor shiner
Slender chub
Streamline chub
Blotched chub
Gravel chub
Tonguetied minnow
Cutlips minnow
Brassy minnow
Eastern silvery minnow
White shiner
Warpaint shiner
Crescent shiner
Striped shiner
Common shiner
Rosefin shiner
Mountain shiner
Redfin shiner
Speckled chub
Silver chub
3earl dace
Horneyhead chub
Jluehead chub
River chub
ull chub
Solden shiner
igeye chub
/Vhitemouth shiner
ugnose shiner
omely shiner
opeye shiner
D-5
-------
EMAP-SW-Streams Field Operations Manual, Appendix D. Fte" n. September 1998 Page 6 of 10
CODE
Common Name
=====£
NOTRAT
NOTRBC
NOTRBH
JNOTRBI
NOTRBL
_
JNOTRCH
INOTRDO
NOTRHD
I —
(NOTRHL
NOTRHU
I
INOTRLE
IJNOTRPH
INOTRPR
NOTRRR
'
INOTRRU
INOTRSC
INOTRSE
NOTRSP
INOTRST
INOTRTE
INOTRVO
======== • —
Notropis atherinoides
Notropis buccatus
Notropis buchanani
Notropis bifrenatus
Notropis blennius
Notropis chalybaeus
Notropis dorsalis
Notropis heterodon
Notropis heterolepis
Notropis hudsonius
Notropis leuciodus
Notropis photogenis
Notropis procne
Notropis rubricroceus
Notropis rubellus
Notropis scabriceps
Notropis semperasper
Notropis spectrunculus
Notropis stramineus
Notropis telescopus
Notropis volucellus
Emerald shiner
Bilverjaw minnow
Shost shiner
Bridled shiner
River shiner
roncolor shiner
Bigmouth shiner
Blackchin shiner
Blacknose shiner
Spottail shiner
Tennessee shiner
Silver shiner
Swallowtail shiner
Saffron shiner
Rosyface Shiner
New River Shiner
Roughhead Shiner
Mirror Shiner
Sand Shiner
Telescope Shiner
Mimic Shiner
Piinnncp minnow
OPSOEM
[PHENCR
IPHENMI
IPHENTE
IPHENUR
IPHOXCU
IPHOXEO
IPHOXER
PHOXNE
Opsopoeodus emiliae
Phenacobius crassilabrum
Phenacobius mirabilis
Phenacobius teretulus
Phenacobius uranops
Phoxinus cumberlandensis
Phoxinus eos
Phoxinus erythrogaster
Phoxinus neogaeus
rugnose IIIIINIUW
Fatlips minnow
Suckermouth minr
Kanawha minnow
Stargazing minnov
Blackside dace
Northern redbelly i
Southern redbelly
Finescale dace
D-6
-------
EMAP-SW-Streams Field Operations Manual, Appendix D, Rev. 0. September 1998 Page 7 of 10
CODE
PHOXOR
PHOXTE
PIMENO
PIMEPR
PIMEVi
RHINAT
RHINBO
RHINCA
RHINOS
SCARER
SEMOAT
SEMOCO
CARAAU
CYPRCA
CTENID
LEUCID
TINCTI
CYPRZZ
FUNDCA
FUNDDI
FUNDRA
LUCAPA
ESOXAM
ESOXLM
ESOXLU
ESOXMA
ESOXNI
LOTALO
APELQU
CULEIN
Latin Name
Phoxinus oreas
Phoxinus tennesseensis
Pimephales notatus
Pimephales promelas
Pimephales vigilax
Rhinichthys atratulus
Rhinichthys bowersi
Rhinichthys cataractae
Rhinichthys osculus
Scardinius erythrophthal.
Semotilus atromaculatus
Semotilus corpora/is
Carassius auratus
Cyprinus carpio
Ctenopharyngodon idella
Leuciscus idus
Tinea tinea
Fundulus catenatus
Fundulus diaphahus
Fundulus rathbuni
Lucania parva
Esox americanus
Esox lucius x masq.
Esox lucius
Esox masquinongy
Esox niger
Lota lota
Apeltes quadracus
Culea inconstans
Common Name
Mountain redbelly dace
Tennessee dace
Bluntnose minnow
Fathead minnow
Bullhead minnow
Blacknose dace
Cheat minnow
Longnose dace
Speckled dace
Rudd
Creek chub
Fallfish
Goldfish
Common carp
Grass carp
Ide
Tench
Unknown cyprinid
Northern studfish
Banded killifish
Speckled killifish
Rainwater killifish
Redf in/grass pickerel
Tiger muskellunge
Northern pike
Muskellunge
Chain pickerel
Burbot
Fourspine stickleback
Brook stickleback
D-7
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EMAP-SW-Streams Field Operations Manual, Appendix D. Rev. 0. September 1998 Page 8 of 10
GASTAC
PUNGPU
HIODAL
HIODTE
AMEIBR
AMEICA
AMEIME
AMEINA
AMEINE
AMEIPL
ICTAPU
NOTUEL
NOTUEX
NOTUFI
NOTUFU
NOTUGI
NOTUGY
NOTUIN
NOTUMI
NOTUST
PYLOOL
MOROAM
MOROCH
MOROSA
AMMOAS
AMMOPE
ETHEAC
ETHEBL
ETHECE
ETHECI
Latin Name
Gasterosteus aculeatus
Pungftius pungitius
Hiodon alosoides
Hiodon tergisus
Ameiurus brunneus
Ameiurus catus
Ameiurus me/as
Ameiurus natalis
Ameiurus nebulosus
Ameiurus platycephalus
Ictalurus punctatus
Noturus eleutherus
Noturus exilis
Noturus flavipinnis
Noturus flavus
Noturus gilbert!
Noturus gyrinus
Noturus insignis
Noturus miurus
Noturus stigmosus
Pylodictis olivaris
\Aorone americana
Morone chrysops
Morone saxatilis
Ammocrypta asprella
Ammocrypta pellucida
Etheostoma acuticeps
Etheostoma blennioides
Etheostoma caeruleum
Etheostoma cinereum
Common Name
Threespine stickleback
Ninespine stickleback
Goldeye
Mooneye
Snail bullhead
White catfish
Black bullhead
Yellow bullhead
Brown bullhead
Flat bullhead
Channel catfish
Mountain madtom
Slender madtom
Yellowfin madtom
Stonecat
Orangefin madtom
Tadpole madtom
Margined madtom
Brindled madtom
Northern madtom
Flathead catfish
White perch
White bass
Striped bass
Crystal darter
Eastern sand darter
Sharphead darter
Greenside darter
Rainbow darter
Ashy darter
D-8
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EMAP-SW-Streams Field Operations Manual, Appendix D, Rev. 0, September 1998 Page 9 of 10
CODE
ETHECM
ETHEEX
ETHEFL
ETHEFU
ETHEJE
ETHEKA
ETHEKE
ETHELO
ETHEMA
ETHENI
ETHEOL
ETHEOS
ETHEPO
ETHERU
ETHESI
ETHEST
ETHESW
ETHETI
ETHEUN
ETHEVA
ETHEVU
ETHEZO
PERCAU
PERCBU
PERCCA
PERCCO
PERCEV
PERCFL
PERCGY
PERCMR
Latin Name
Etheostoma camurum
Etheostoma exile
Etheostoma flabellare
Etheostoma fusiforme
Etheostoma jessiae
Etheostoma kanawhae
Etheostoma kennecotti
Etheostoma longimanum
Etheostoma maculatum
Etheostoma nigrum
Etheostoma olmstedi
Etheostoma osburni
Etheostoma podostemone
Etheostoma rufilineatum
Etheostoma simoterum
Etheostoma stigmaeum
Etheostoma swannanoa
Etheostoma tippecanoe
Etheostoma unknownOI
Etheostoma variatum
Etheostoma vulneratum
Etheostoma zonale
Percina aurantiaca
Percina burtoni
Percina caprodes
Percina copelandi
Percina evides
Perca flavescens
Percina gymnocephala
Percina macrocephala
Common Name
Bluebreast darter
Iowa darter
Fantail darter
Swamp darter
Blueside darter
Kanawha darter
Stripetail darter
Longfin darter
Spotted darter
Johnny darter
Tesselated darter
Candy darter
Riverweed darter
Redline darter
Snubnose darter
Speckled darter
Swannanoa darter
Tippecanoe darter
Duskytail darter
Variegate darter
Wounded darter
Banded darter
Tangerine darter
Blotchside logperch
Logperch
Channel darter
Gilt darter
Yellow perch
Appalachia darter
.onghead darter
D-9
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EMAP-SW-Streams Field Operations Manual, Appendix D. Rev. 0. September 1998 Page 10 of 10
CODE
PERCMU
PERCNO
PERCOX
PERCPE
PERCRE
PERCRO
PERCSC
PERCSH
STIZCA
STIZVI
PERCOM
GAMBAF
GAMBHO
ONCOMY
SALMTR
SALVFO
SALVNA
SALMZZ
APLOGR
UMBRLI
Latin Name
Percina maculate
Percina notogramma
Percina oxyrhynchus
Percina peltata
Percina rex
Percina roanoka
^ercina sciera
Percina shumardi
Stizostedion canadense
Stizostedion vftreum
Percopsis omiscomaycus
Gambusia affinis
Gambusia holbrooki
Oncorhynchus mykiss
Salmo trutta
Salvelinus fontinalis
Salvelinus namaycush
Aplodinotus grunniens
Umbra limi
Common Name
Blackside darter
Stripeback darter
Sharpnose darter
Shield darter
Roanoke logperch
Roanoke darter
Dusky darter
River darter
Sauger
Walleye
Troutperch
Western mosquitoflsh
Eastern mosquitofish
Rainbow trout
Brown trout
Brook trout
Lake trout
Unknown salmonid
Freshwater drum
Central mudminnow
Eastern mudminnow
D-10
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APPENDIX E
MODIFIED PROTOCOL FOR COLLECTING BENTHIC MACROJNVERTEBRATES
by
Donald J. Klemm1 and David V. Peck2
Field procedures described here are modified from those developed by the Oregon
Department of Environmental Quality (Oregon Dept. of Environmental Quality, 1997), and
the Washington Department of Ecology (Washington Dept. of Ecology, 1997). These
procedures were implemented in an EMAP study of wadeable streams in Oregon in 1997,
and the EPA Region 10 R-EMAP study in 1996-1997. Modifications to the basic EMAP
protocol (Section 11 of the EMAP field operations manual for streams) were desired to
maximize the comparability of EMAP results with both the R-EMAP project results, and with
other data both State agencies routinely collect as part of their respective monitoring pro-
grams.
Within the defined sampling reach of 150 to 500 m, benthic invertebrate samples are
collected from two principal macrohabitat types, erosional (operationally termed "riffle") and
depositional (operationally termed "pool"). Riffle macrohabitats include low-gradient areas
that are generally more shallow than pools. Many riffles exhibit surface turbulence associ-
ated with increased velocity and shallow water depth over gravel or cobble beds. However,
the riffle classification also includes shallow areas without surface turbulence such as
glides. Pool macrohabitats include areas of slow, deep water with low gradient. They are
typically created by scour adjacent to obstructions or impoundments of water behind chan-
nel blockages and hydraulic controls such as logjams, bedforms, or beaver dams.
Individual kick net samples are collected from up to five points within each macro-
habitat type, spaced throughout the sampling reach. Individual kick net samples collected
from each macrohabitat type are processed and composited into a single sample for the
stream. Thus for each stream, there will be two composite samples, one for riffles and one
U.S. EPA, National Exposure Research Laboratory, Ecological Exposure Research Division, 26 W. Martin L. King Dr,
Cincinnati, OH 45268.
2 U.S. EPA, National Health and Environmental Effects Research Laboratory, Western Ecology Division, 200 SW 35th St.,
Corvallis, OR 97333.
E-1
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EMAP-SW-Streams Field Operations Manual, Appendix E. Rev. 0, September 1998 Page 2 of 9
for pools. Each composite sample is contained in a 500-mL or 1-L plastic screw-top jar, and
preserved with 95% ethanol to a final concentration of 70% ethanol.
The sampling protocols described here differ from those presented in Section 11 of
the EMAP streams field operations manual in that sampling points are allocated evenly
across the two major macrohabitat types, rather than by sampling at predefined points
located on the transects established for physical habitat characterization. Differences from
R-EMAP protocols used by Oregon DEQ (1997)and Washington Department of Ecology
(1997) include not determining the depth at each sampling point and not determining the
substrate particle distribution at each sampling points.
E.1 SAMPLE COLLECTION
E.1.1 Selection of Sampling Points
Table E-1 presents the procedure for selecting individual sampling points within the
two major macrohabitat types (riffle and pool). Note that in some stream reaches, one
macrohabitat type will predominate to the extent that the other type is not sampled. There
may also be stream reaches where two kick net samples are collected from a single macro-
habitat unit. It is also permissable to sample a short distance beyond the upstream end of
the sample reach in order to obtain the desired number (5) of macrohabitat units of each
type.
E.1.2 Collection of Kick Net Samples
The kick net is designed to obtain a qualitative and semi-quantitative sample of
benthic macroinvertebrates from a variety of substrates in streams. A modified USGS kick
net (Wildco # 425-J50-595) is used. This is the same net as is described in Section 10 of
the EMAP field operations manual for streams for EMAP. Modifications from the standard
configuration include the net mesh size (600 pm), the length of the net (61 cm or 24 in.) and
the type of bag (tapering closed bag). The frame dimensions of the net are 30.48 cm (12
in,) high and 50.8 cm (20 in.) wide. The style and dimensions of this net differ from that
used by Oregon DEQ and Washington Department of Ecology, who use a smaller net of a
D-frame configuration. However, mesh sizes are the same.
Procedures for collecting a point sample using the kick net from riffle and pool
macrohabitat units are presented in Tables E-2 and E-3, respectively. At each sampling
E-2
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EMAP-SW-Streams Field Operations Manual, Appendix E, Rev. 0, September 1998 Page 3 of 9
TABLE E-1. LOCATING SAMPLING POINTS FOR KICK NET SAMPLES:
WADEABLE STREAMS
1. Before sampling, survey the stream reach to visually estimate the number of pool and riffle
macrohabitat "units" contained in the defined stream reach. To be considered as a unit, the
length of a stream occupied by a particular macrohabitat type unit should be at least equal to
the stream's average wetted width estimate used to define the length of stream reach.
A. Do not sample poorly represented habitats. If the reach contains < 2 macrohabitat units
of a given type, then do not sample that macrohabitat type. If only one macrohabitat unit
occurs in the defined reach but, more are present within 100 meters upstream, sample
those as they were part of the reach.
B. If the reach contains 3 or 4 macrohabitat units of a given type, then randomly select
those macrohabitat unit(s) from which to collect a second kick net sample to bring the
total number of kick net samples for the macrohabitat type to five.
C. If the number of units is greater than five of either, skip one or more habitat units at
random as you work upstream.
2. Begin sampling at the most downstream unit, and sample units as they are encountered to
minimize instream disturbance. This will require separate containers for pool and riffle sam-
ples.
3. At each unit, exclude "margin" habitats by constraining the potential sampling area,- Margin
habitats are edges, along the channel margins or upstream or downstream edges of the
macrohabitat unit. Define a core area for each unit as the central portion, visually estimating a
"buffer" strip circumscribing the identified unit. In some cases, the macrohabitat unit may be
so small that it will not be feasible to define a core area and avoid an edge.
4. Visually lay out the core area of the unit sampled into 9 equal quadrats (i.e., a 3 * 3 grid). For
each macrohabitat type, select a quadrat for sampling as follows:
First unit: Lower right quadrat
' Second unit: Center quadrat
Third unit: Upper left quadrat
Fourth unit: Lower left quadrat
Fifth unit: Upper right quadrat.
5. Collect the kick sample in the center of the selected quadrat, following the protocol for the type
of macrohabitat unit.
6. If a second sample is required from a single macrohabitat unit, select a new quadrat.
E-3
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EMAP-SW-Streams Field Operations Manual. Appendix E. R*v n September 1998 Page 4 of 9
TABLE E-2. COLLECTING A KICK NET SAMPLE FROM WADEABLE STREAMS:
RIFFLE MACROHABITATS
1. Locate the sampling point within the macrohabitat unit as described in Table 1.
2 Position the kick net quickly and securely on the stream bottom so as to eliminate gaps be-
tween the frame and the stream bottom. If necessary, rotate the net so the narrower side is
against the bottom.
3 Hold the sampler firmly in position on the substrate. Define a quadrat immediately upstream
" from the mouth of the net having a width equal to the width of the net frame and a total area -
05m2 If the kick net is oriented normally, the length of the quadrat = 1 m (approx equal to 2
times the width of the net [0.5 m]). If the net is rotated so the short side is against the sub-
strate, the length of the quadrat = 1.67 m.
4. Lightly kick the substrate throughout the quadrat. Start at the upstream end and work toward
the net.
5 Hold the net in place with the knees and pick up any loose rocks in the quadrat and rub off
' organisms so that they are washed into the net. With a small brush dislodge organisms from
the rocks into the net. Scrub all rocks that are golf ball-sized or larger and which are over
hallway into the quadrat. Large rocks that are less than halfway into the sampling area are
pushed aside.
6. Keep holding the sampler securely in position and kick through the quadrat again, this time
vigorously, for 20 seconds.
7 Pull the net up out of the water. Immerse the net in the stream several times to remove fine
sediments and to concentrate organisms at the end of the net. Avoid having any water or
material enter the mouth of the net during this operation.
8 Invert the net into a plastic bucket marked "riffle" and transfer the sample Inspect the net for
any residual organisms clinging to the net and deposit them into the "riffle" bucket. Use watch-
makers' forceps if necessary to remove organisms from the net.
9. Thoroughly rinse the net before proceeding to the next macrohabitat unit.
10. Repeat steps 1-9 at subsequent riffle macrohabitat units until 5 kick samples have been col-
lected and placed into the "riffle" bucket.
E-4
-------
1.
2.
3.
4.
5.
6.
7.
8.
EMAP-SW-Streams Field Operations Manual, Appendix E, Rev. 0, September 1998 Page 5 of 9
TABLE E-3. COLLECTING A KICK NET SAMPLE FROM WADEABLE STREAMS:
POOL MACROHABITATS '
Locate the sampling point within the macrohabitat unit as described in table 1.
Define a sampling area as a quadrat having a width equal to the width of the net frame and a
total area = 0.5 m2. If the kick net is oriented normally, the length of the quadrat = 1 m (approx.
equal to 2 times the width of the net [0.5 m]). If the net is rotated so the short side is against
the substrate, the length of the quadrat = 1.67 m. •
Inspect the quadrat for heavy organisms such as mussels and snails. Hand pick any of these
large organisms and place them into the sieve bucket or plastic bucket marked "pool".
Kick vigorously with the feet within the quadrat for 10 seconds. Then drag the net repeatedly
through the disturbed area just above the bottom. Keep moving the net to prevent organisms
from escaping. Continue this for 1 minute.
Pull the net up out of the water. Immerse the net into 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.
Invert the net into the bucket marked "pool" and transfer the sample. Inspect the net for any
residual organisms clinging to the net and deposit them into the "pool" sieve bucket. Use
watchmakers' forceps if necessary to remove organisms from the net.
Thoroughly rinse the net before proceeding to the next macrohabitat unit to prevent cross-
contamination of riffle and pool samples.
Repeat steps 1-7 at subsequent pool macrohabitat units until 5 kick samples have been col-
lected and placed into the "pool" sieve bucket.
E-5
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EMAP-SW-Streams Field Operations Manual. Appendix E. Rev. 0, September 1998 Page 6 of 9
point, a quadrat having a total area of 0.5 m2 is sampled. The dimensions of the quadrat
will vary depending on how the kick net must be oriented against the substrate. In narrow
streams, the net may have to be rotated so that the narrow side is against the stream
bottom. Riffle and pool samples are kept in separate containers. Note that in pool units,
the substrate is first disturbed, and the net is dragged through the disturbed area just above
the substrate. Because units are sampled in the order they are encountered, it is very
important to rinse the kick net thoroughly between samples to avoid carryover and possible
cross-contamination of riffle and pool samples.
E.2 Sample Processing
The procedure for processing kick net samples is presented in Table E-4; the proce-
dure is identical for riffle and pool samples. Process one sample at a time to avoid mixing
riffle and pool samples in the same container. Reduce the amount of residue in each
composite sample as much as possible without losing organisms. However, if there is a
sizable quantity of material remaining, distribute the sample into additional containers to
ensure proper preservation. A sample jar should not be more than half-full of material.
Modified sample labels are shown in Figure E-1, and the modified Sample Collection Form
is presented in Figure E-2.
E.3. LITERATURE CITED
Oregon Department of Environmental Quality. 1997. Biological Assessment of Wadeable
Streams of the Upper Deschutes River Basin: Quality Assurance Project Plan.
Washington Department of Ecology. 1997. Biological Assessment of Wadeable Streams of
the Chehalis River Basin: Quality Assurance Project Plan.
E-6
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EMAP-SW-Streams Field Operations Manual, Appendix E, Rev. 0, September 1998 Page 7 of 9
TABLE E-4. PROCESSING KICK NET SAMPLES: WADEABLE STREAMS
1. Fill out a sample label for the riffle composite samples. Attach a label to a 1-gallon plastic bag
with a zipper-type closure. If the sample contains a large volume of material, complete a
sample label for additional containers and attach it to a second bag. Make sure the barcode
numbers on each label agree.
2. Hand pick large organisms from the bucket containing the composited riffle kick net samples
and place them into the appropriately labeled plastic bag.
3. Hand pick large rocks and sticks remaining in the bucket. Use a small brush to scrub debris
from them back into the bucket. Discard the rock or stick.
4. Empty the contents of the bucket into the labeled plastic bag. If necessary, distribute the
sample among two or more labeled bags. Rinse residue from the bucket into the plastic bag
using a wash bottle and a small volume of water.
5. Place each bag inside a second bag.
6. Add 95% ethanol to each labeled bag in a volume which is equal to the volume of the sample.
7. Rinse the bucket well to eliminate any residue.
8. Repeat Steps 1-7 for the pool composite sample.
9. Complete the Sample Collection Form. Record the barcode number of each composite sam-
ple (riffle and pool), and the habitat type from the sample label. If more than one container
was required for a sample, record the number of containers on the collection form. Also, note
any peculiarities associated with a particular sample by using a flag code and/or a written
comment on the collection form.
E-7
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EMAP-SW-Streams Field Operations Manual, Appendix E, Rev. 0, September 1998 Page 8 of 9
COMPOSITE BENTHOS
SITE ID: ORST97 -
DATE: / / 97
HABITAT: Riffle Pool
238200
Figure E-1. Modified sample labels.
E-8
-------
EMAP-SW-Streams Field Operations Manual, Appendix E, Rev. 0, September 1998 Page 9 of 9
Reviewed by (initial):.
SITE NAME: DATE: / / 97 VISIT: D1 D2
SITE ID: ORST97- TEAM ID (X): D1 O2 D3 D4 DS D6 D7 D8
- " J: 'jSdtePOSllE BENTHOS SfffflpW -/ '".., " ' * - 'V t
SAMPLE ID
* (BAHCODE)
s
V S S
HABITAT NCJ >
R R JAR " T ~ \
STATION A
RIFFLE on POOt-
(XONE)-
LEFT, CENTER, OR u
RK3MT- ' -
B ' C D B F 'Q' * H
DR DR DR DR DR DR DR
DP DP DP DP DP DP DP
DL DL DL DL DL DL DL
DC DC DC DC DC DC DC
DR DR DR DR DR DR DR
I J t K
DR DR
DP DP
DL DL
DC DC
DR DR
COttP9w1IJPp^PWfTdl* SAMPLES. _„ 2 *"* HABITAT TYPE (X)~ D RIFFLE DPOOL D OTHER
SAMPLE ID (BARCODE) -
ASSEMBLAGE ID
(50-HLTUBE)
SUB. SAMPLE VOL.
ML
« <%>t Tj *• T- "V^ ^
SAMPLE 10 (BARCOBE),-
ASSEMBLAGE ID
(50-HLTUBE)
SUB. SAMPLI VOL.
ML
COM^OSlTEyOLUME-. «L
CHLOROPHYLL BIOMASS
«5F/F FILTER) (TARRED RLTEH)
VOL. FILTERED FILTER No. VOL. FILTERED
ML ML
.-V - ^ ,!<., 1
APASAMPLE"
(SO-MLTUBE)
' SUB. SAMPLE You.
ML
»ittOWSAlPt^' j:"*-' HABITATTYPEPO- DR.FFLE D POOL D OTHER
COMPOSITE VOLUME - »L
' < \ 1 4
CHLOROPHYLL BlOMASS / : "
(GF/FRLTEB) '(TAHEOFiLTEB)
VOL. FILTERED FtTEBNo. VOL.FILTEHED
. . uL . ML
' APft'SAMPlE-
SUB.SMJPLEVOL.
ML
^^^lij|^|^5:f. ;i|
•
Flag codes: K= Sample not collected; U= Suspect sample; F1, F2, etc.= misc. flag assigned by field crew. Explain all flags In Comment sections.
Rev. 06/02/97 (sl_saco.97)
Figure E-2. Modified Sample Collection Form.
SAMPLE COLLECTION FORM - STREAMS - 1
l!rU.S. GOVERNMENT PRINTING OFFICE: 1999-750-101/00031
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