Revised Protocols for Sampling Algal, Invertebrate, 

and Fish Communities as Part of the National 

Water-Quality Assessment Program

Open-File Report 02-150

U.S. Department of the Interior

U.S. Geological Survey
Front Cover: 

A. Nymph of the stonefly Calineuria californica (Banks). (Photograph by James Carter)

B. A crewmember scraping algae from a cobble. (Photograph by Mitchell Harris)

C. The diatom Navicula lacustris Greg. (Photograph by Phycology Section, 

The Academy of Natural Sciences of Philadelphia)

D. Crewmembers sampling fishes with barge electrofishing gear. 

(Photograph by Rodney Knight)

E. The redfin darter, Etheostoma whipplei (Girard). (Photograph copyright Joseph R. 

Tomelleri)

F. Two crewmembers sampling invertebrates with a Slack sampler. (Photograph by 

Dennis Sun)

A

B

C

D

E

F
Revised Protocols for Sampling Algal, Invertebrate, 

and Fish Communities as Part of the National 

Water-Quality Assessment Program

By Stephen R. Moulton II, Jonathan G. Kennen, Robert M. Goldstein, and Julie A. Hambrook

Open-File Report 02-150

Reston, Virginia

2002
U.S. DEPARTMENT OF THE INTERIOR

GALE A. NORTON, Secretary

U.S. GEOLOGICAL SURVEY

Charles G. Groat, Director

The use of trade or product names in this report is for identification purposes 

only and does not constitute endorsement by the U.S. Government.

For additional information write to: 

U.S. Geological Survey

Chief, National Water-Quality Assessment Program

413 National Center

12201 Sunrise Valley Drive

Reston, VA 20192

Information regarding the National Water-Quality Assessment (NAWQA) Program is available on the Internet at 

http://water.usgs.gov/nawqa/. Copies of this report can be downloaded from this address.
The U.S. Geological Survey (USGS) is

committed to providing the Nation with accurate and

timely scientific information that helps enhance and

protect the overall quality of life and that facilitates

effective management of water, biological, energy, and

mineral resources. Information on the quality of the

Nation’s water resources is critical to assuring the longterm availability of water that is safe for drinking and

recreation and suitable for industry, irrigation, and

habitat for fish and wildlife. Population growth and

increasing demands for multiple water uses make water

availability, now measured in terms of quantity and

quality, even more essential to the long-term

sustainability of our communities and ecosystems.

The USGS implemented the National WaterQuality Assessment (NAWQA) Program in 1991 to

support national, regional, and local information needs

and decisions related to water-quality management and

policy. Shaped by and coordinated with ongoing efforts

of other Federal, State, and local agencies, the

NAWQA Program is designed to answer: What is the

condition of our Nation’s streams and ground water?

How are the conditions changing over time? How do

natural features and human activities affect the quality

of streams and ground water, and where are those

effects most pronounced? By combining information

on water chemistry, physical characteristics, stream

habitat, and aquatic life, the NAWQA Program aims to

provide science-based insights for current and

emerging water issues and priorities. NAWQA results

can contribute to informed decisions that result in

practical and effective water-resource management and

strategies that protect and restore water quality.

From 1991–2001, the NAWQA Program

completed interdisciplinary assessments in 51 of the

Nation’s major river basins and aquifer systems,

referred to as Study Units. Baseline conditions were

established for comparison to future assessments, and

long-term monitoring was initiated in every basin

studied. Financial constraints will reduce the number

of Study Units that NAWQA can assess in the future;

therefore, during the next decade, 42 of the 51 Study

Units will be reassessed so that 10 years of comparable

monitoring data will be available to determine trends at

many of the Nation’s streams and aquifers. The next 10

years of study also will fill in critical gaps in

characterizing water-quality conditions, enhance

understanding of factors that affect water quality, and

establish links between sources of contaminants, the

transport of those contaminants through the hydrologic

system, and the potential effects of contaminants on

humans and aquatic ecosystems.

The USGS aims to disseminate credible, timely,

and relevant science information so that the most

current knowledge about water resources can be

applied in management and policy decisions. We hope

this NAWQA publication will provide you with

insights and information to meet your needs, and will

foster increased citizen awareness and involvement in

the protection and restoration of our Nation’s waters.

The USGS recognizes that a national assessment

by a single program cannot address all water-resource

issues of interest. External coordination at all levels is

critical for a fully integrated understanding of

watersheds and for cost-effective management,

regulation, and conservation of our Nation’s water

resources. The NAWQA Program, therefore, depends

on advice and information from other agencies—

Federal, State, interstate, Tribal, and local—as well as

nongovernment organizations, industry, academia, and

other stakeholder groups. Your assistance and

suggestions are greatly appreciated.

Timothy L. Miller

Chief, NAWQA Program

FOREWORD

(signed)
Moulton and others—Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program—OFR 02-150
 Contents V

CONTENTS

Abstract ....................................................................................................................................................................................................... 1

Introduction ................................................................................................................................................................................................ 1

Overview of the NAWQA Program .............................................................................................................................................. 1

Purpose and scope ........................................................................................................................................................................ 1

Summary of revisions to original protocols ............................................................................................................................... 2

Acknowledgments ......................................................................................................................................................................... 2

Field protocols ............................................................................................................................................................................................ 3

1. Overview of the sampling approach ....................................................................................................................................... 3

1.1 Establishing the sampling reach ................................................................................................................................ 3

1.2 Types of samples collected ......................................................................................................................................... 3

1.3 Identifying instream-habitat types for sampling ...................................................................................................... 4

1.4 Factors determining when to sample ......................................................................................................................... 4

1.5 Coordination of field activities .................................................................................................................................... 4

2. Pre-field activities ...................................................................................................................................................................... 5

2.1 Preparing a sampling plan ........................................................................................................................................... 6

2.2 Collecting permits ......................................................................................................................................................... 6

2.3 Supplies and chemicals ............................................................................................................................................... 6

2.4 Sample preservatives ................................................................................................................................................... 8

2.5 Sample identification and tracking ............................................................................................................................ 8

2.6 Sampling gear maintenance ....................................................................................................................................... 10

3. Safety in the field ........................................................................................................................................................................ 12

3.1 Personal and environmental safety ........................................................................................................................... 12

3.2 Chemical safety ............................................................................................................................................................. 13

3.3 Vehicles, boats, and sampling gear safety ............................................................................................................... 14

4. Algal sampling protocols .......................................................................................................................................................... 14

4.1 Algal sample types collected ...................................................................................................................................... 15

4.2 Identifying instream habitats for sampling ............................................................................................................... 15

4.3 Quantitative targeted-habitat sampling methods .................................................................................................... 16

4.4 Qualitative multihabitat sampling methods .............................................................................................................. 22

4.5 Sample processing procedures ................................................................................................................................. 24

4.6 Quality-control recommendations ............................................................................................................................. 28

4.7 Field data sheets ........................................................................................................................................................... 28

5. Invertebrate sampling protocols ............................................................................................................................................. 36

5.1 Invertebrate sample types collected ......................................................................................................................... 36

5.2 Identifying instream habitats for sampling ............................................................................................................... 36

5.3 Semi-quantitative targeted-habitat methods for sampling wadeable and nonwadeable streams ................ 37

5.4 Qualitative multihabitat methods for sampling wadeable and nonwadeable streams ..................................... 41

5.5 Sample processing procedures ................................................................................................................................. 42

5.6 Field data sheets ........................................................................................................................................................... 48

6. Fish Sampling Protocols ............................................................................................................................................................ 55

6.1 Overview of sampling methods .................................................................................................................................. 55

6.2 Sampling methods for wadeable streams ................................................................................................................ 56

6.3 Sampling methods for nonwadeable streams ......................................................................................................... 59

6.4 Alternative sampling methods .................................................................................................................................... 60

6.5 Sample processing procedures ................................................................................................................................. 60

6.6 Field data sheet ............................................................................................................................................................. 66
VI Contents 

7. Post-Field Activities ................................................................................................................................................................... 69

7.1 Logging samples into the Biological Transactional Data Base ............................................................................. 69

7.2 Replacing field sample labels ..................................................................................................................................... 69

7.3 Preparing packing lists and Analytical Services Request Forms ......................................................................... 69

7.4 Shipping samples to analytical laboratories ............................................................................................................ 69

Summary ..................................................................................................................................................................................................... 70

References cited ....................................................................................................................................................................................... 73

Appendix ..................................................................................................................................................................................................... 75

FIGURES

1. Sample identification code used to distinguish algal (top), invertebrate (middle) and fish 

(bottom) samples in the National Water-Quality Assessment Program .................................................................... 9

2. Examples of field sample labels used to uniquely identify algae ................................................................................ 11

3. Methods used to collect microalgal periphyton samples ............................................................................................ 17

4. Examples of common macroalgae ................................................................................................................................... 23

5. Steps followed in the field to process algal samples .................................................................................................... 25

6. Major steps in processing microalgal sample components in the field .................................................................... 26

7. Priority levels for determining the habitat type that is selected as the richest targeted 

habitat for semi-quantitatively sampling invertebrates ................................................................................................ 38

8. Samplers used to collect invertebrate samples ............................................................................................................. 39

9. Steps followed in the field to process invertebrate samples ...................................................................................... 43

10. Key steps in the collection and processing of invertebrate samples ........................................................................ 44

11. Steps followed in the field to process fish collections ................................................................................................. 61

12. Total and standard length measurements of a fish ....................................................................................................... 63

13. Deformities, eroded fins, lesions, and tumors noted on fish ........................................................................................ 65

TABLES

1. Hierarchical framework for geomorphic descriptors used to determine habitats from 

which algal and invertebrate samples are collected .................................................................................................... 5

2. Definition of character fields used to create sample identification codes for 

algae, invertebrates, and fish samples ............................................................................................................................ 8

3. Algal sample types, sample components, and subsamples collected ....................................................................... 15

4. Explanation of field data-sheet sections used to record information 

about algal sampling activities .......................................................................................................................................... 29

5. Generalized matrix of the 51 instream habitats .............................................................................................................. 37

6. Explanation of field data-sheet sections used to record information 

about invertebrate sampling activities ............................................................................................................................ 48

7. Two-letter codes used to denote external anomalies in fish ....................................................................................... 64

8. Explanation of field data-sheet sections used to record information 

about fish sampling activities ............................................................................................................................................ 66
 Contents VII

Multiply By To obtain

Length

micrometer (μm) 0.00003937 inch

millimeter (mm) 0.03937 inch

centimeter (cm) 0.3937 inch

meter (m) 3.281 foot

kilometer (km) 0.6214 mile

Area

square centimeter (cm2) 0.155 square inch

square meter (m2) 10.76 square foot

square kilometer (km2) 0.3861 square mile

Volume

liter (L) 1.057 quart

liter (L) 0.2642 gallon

milliliter (mL) 0.0338 ounce, fluid

Flow

centimeter per second (cm/s) 0.0328 foot per second

Mass

gram (g) 0.03527 ounce, avoirdupois

Pressure

kilopascal (kPa) 0.1450 pound-force per square inch

CONVERSION FACTORS, VERTICAL DATUM, AND ABBREVIATED 

WATER-QUALITY UNITS

Temperature Conversion

Degrees Celsius (°C) may be converted to degrees Fahrenheit (°F) by using the following equation:

°F = 1.8 (°C) + 32

ABBREVIATIONS FREQUENTLY USED IN THIS REPORT

AFDM ash-free dry mass QA/QC quality assurance/quality control

ANSP Academy of Natural Sciences of Philadelphia QC quality control

ASR analytical services request QMH qualitative multihabitat

Bio-TDB Biological Transactional Data Base RTH richest-targeted habitat

CHL A chlorophyll a SMCOD sample identification code

DTH depositional targeted habitat USGS U.S. Geological Survey

ID identification > greater than

NAWQA National Water-Quality Assessment < less than

NWQL National Water Quality Laboratory ≥ greater than or equal to

PFD personal flotation device ≤ less than or equal to

PHY phytoplankton ± plus or minus

QA quality assurance
VIII Glossary 

Algae—Chlorophyll-bearing, nonvascular aquatic plants (for 

example, diatoms, green algae, and red algae) including 

the primitive photosynthetic bacteria (for example, bluegreen algae).

Anadromous—Refers to fish species (for example, some 

species of salmon) that live in the ocean as adults and 

return to freshwater to spawn.

Benthic—Refers to bottom; for example, benthic organisms 

that live on or burrow into an aquatic substrate.

Brails—The poles supporting each end of a seine.

Coarse-grained substrates—Stream substrates (> 2 mm 

widest dimension) composed primarily of gravels, cobbles, and boulders.

Component—See sample component.

Composite sample—See sample.

Decant volume (algae only)—The volume associated with 

decanting an algal sample or subsample in order to concentrate the material by removing excess liquid. There 

are two types of decant volume: (1) before decant volume 

(BDV)—the volume before excess liquid is removed; and 

(2) after decant volume (ADV)—the volume after excess 

liquid is removed.

Depositional-targeted habitat (DTH)—A habitat targeted 

for sampling in the reach where fine sediments (for 

example, sand and silt) are deposited. A composited sample from this habitat is referred to as a "DTH sample."

Discrete collection—The material collected from a single 

location within a reach. Discrete collections are composited either physically (sensu invertebrates) or logically 

(sensu algae) to create a sample. For example, a single 

invertebrate RTH sample is composed of five discrete 

collections that are physically combined. Likewise, an 

algae QMH sample is composed of discrete collections 

that are composited according to growth form (either 

micro- or macroalgae) that then collectively form a logical algal sample from the reach.

Discrete sample—See discrete collection.

Ecological studies—Studies of biological communities and 

habitat characteristics to evaluate the effects of physical 

and chemical characteristics of water and hydrologic 

conditions on aquatic biota and to determine how biological and habitat characteristics differ among environmental settings in NAWQA study areas.

Epidendric—Benthic habitat consisting of woody substrates 

on which organisms are attached or loosely associated.

Epilithic—Benthic habitat consisting of natural, coarsegrained substrates (for example, gravels, cobbles, or 

boulders) or bedrocks, or artificial hard substrates such as 

submerged concrete on which organisms are attached or 

loosely associated.

Epipelic—Benthic habitat consisting of silt-sized 

(<0.064 mm) streambed sediments on which organisms 

are loosely associated. This habitat is commonly found in 

areas of low velocities, such as pools and side-channel 

areas, where silt can deposit.

Epiphytic—Benthic habitat consisting of plants on which 

organisms are attached or loosely associated.

Epipsammic—Benthic habitat consisting of sand-sized 

(> 0.064 – 2 mm) particles on which organisms are 

attached or loosely associated.

External anomalies—Visible body surface or subcutaneous 

disorders in fish, such as deformities, eroded fins, 

lesions, and tumors.

Fine-grained substrates—Stream substrates (< 2 mm widest dimension) composed primarily of silt and sand.

Fixed site—Sites on streams at which streamflow is measured and samples are collected to assess the broad-scale 

spatial and temporal character and transport of inorganic 

constituents of streamwater in relation to hydrologic conditions and environmental settings.

Formaldehyde—As used for a preservative, the term 

"formaldehyde" is marketed as a saturated water solution 

of formaldehyde gas (HCHO), representing a 

concentrated stock solution of formalin (37-percent 

formaldehyde by weight). Also referred to as 

“concentrated formalin" or "100 percent formalin."

Formalin—A mixture of formaldehyde and water. Tenpercent formalin is prepared by mixing one part formaldehyde with 9 parts water; this mixture is typically used 

for preserving invertebrate and fish samples.

Galvanotaxis—A response by fish to swim toward the 

anode during application of direct current during electrofishing.

Geomorphic channel units—Fluvial geomorphic descriptors of channel shape and stream velocity. Pools, riffles, 

and runs are three types of geomorphic channel units 

considered for NAWQA Program ecological sampling.

Habitat—In general, aquatic habitat includes all nonliving 

(physical) aspects of the aquatic ecosystem (Orth, 1983), 

although living components like macrophytes and riparian vegetation also are usually included. Measurements 

of habitat are typically made over a wider geographic 

scale than measurements of species distribution.

Hyporheic zone—The subbenthic habitat of interstitial 

spaces between substrate particles in the streambed bordered by surface water above and true ground water 

below. This zone acts as a faunal reservoir and refugium 

for aquatic invertebrates during floods, droughts, and 

extreme temperatures.

Invertebrates—Animals that do not have backbones, such 

as worms, clams, crustaceans, and insects.

Meandering stream—Typically a low gradient stream with 

high channel sinuosity.

Motile—Refers to organisms that have the ability to move, 

often rapidly and spontaneously.

GLOSSARY
 Glossary IX

Periphyton—Algae attached to an aquatic substrate; also 

known as benthic algae.

Phytoplankton—Floating or weakly swimming microscopic algae; also known as algal seston in streams.

Pool—A part of the reach with little velocity, commonly 

with water deeper than surrounding areas.

Preservative volume (PV, algae only)—The volume of the 

preservative added to an algal subsample that is submitted for taxonomic identification analysis.

Qualitative multihabitat (QMH)—A series of different 

habitats identified in a reach from which discrete collections of algae or invertebrates are taken and later combined to form a composite sample. The composited 

sample is referred to as a QMH sample.

Reach—A length of stream (150–300 m for wadeable 

streams; 300–1,000 m for nonwadeable streams) that is 

chosen to represent a uniform set of physical, chemical, 

and biological conditions within a segment. It is the principal sampling unit for collecting physical, chemical, and 

biological data.

Relative abundance—The number of organisms of a particular taxon present in a sample relative to the total number 

of organisms of that sample.

Replicate sample—The 2nd to nth independently collected 

sample of the same type from the same reach on the same 

day. For example, a replicated invertebrate sample would 

be the second invertebrate RTH sample collected on a 

particular date from the same reach.

Repeated-sampling replicate–See Replicate sample.

Retrospective analysis—The review and analysis of existing water-quality and ecological data to provide a historical perspective of water quality and biological integrity, 

to assess strengths and weaknesses of available information, and to evaluate implications for water-quality management and NAWQA Program design.

Richest-targeted habitat (RTH)—A targeted habitat (usually a riffle or woody snag where the taxonomically richest algal or invertebrate community is theoretically 

located) identified in a reach from which discrete collections of algae or invertebrates are taken and later combined to form a composite sample. The composited 

sample is referred to as a "RTH sample."

Riffle—A shallow part of the stream where water flows 

swiftly over completely or partially submerged obstructions to produce surface agitation.

Riffle-pool stream—A stream composed of alternating riffles and pools.

Run—A shallow part of a stream with moderate velocity 

and little or no surface turbulence.

Sample—Operationally defined as all of the material and 

organisms collected during one application of the 

NAWQA Program sampling protocol for a particular 

sample type (for example, invertebrate RTH sample). A 

single sample may be subdivided during field processing 

to create sample components (sensu invertebrates and 

algae) or subsamples (sensu algae).

Sample component—(Algae) Discrete collections of algae 

that are composited according to growth form (for example, micro- or macroalgae) to represent a particular sample or subsample. (Invertebrates) A subset of an 

invertebrate sample that is produced after the sample is 

processed in the field. Field processing can produce up to 

four different sample components: large-rare, main-body, 

split, and elutriate.

Sample identification code (SMCOD)—A 16-character 

alphanumeric code that uniquely identifies each sample.

Sample medium—The type of biological community being 

sampled (algae, invertebrates, or fish).

Sample number—A four-digit number given to each sample type.

Sample type—A certain type of algal or invertebrate sample 

collected in a reach from either a single targeted habitat 

or multiple habitats.

Sample volume (SV, algae only)—The total volume of a 

composited algal sample before any preservative is added 

or subsamples taken.

Sensu—Latin, meaning "in the sense of."

Sinuosity—The ratio of the channel length between two 

points on a channel to the straight-line distance between 

the same two points; a measure of meandering.

Split (algae only)—The 2nd to nth subsample that will be 

analyzed for the same purpose as another subsample 

taken from the same sample. For example, the second 

chlorophyll a subsample taken from a particular RTH 

sample is a split.

Subsample (algae only)—A subset of the sample that is 

usually created during field processing. For example, a 

single periphyton RTH sample is typically subdivided 

into three subsamples, each one for a different laboratory 

analysis, such as identification/enumeration, chlorophyll 

a, and ash-free dry mass.

Subsample volume (SSV, algae only)—The volume of the 

material removed from a composite sample for a particular analysis. Subsample volume is measured before any 

preservative is added.

Voucher collection—An assortment of preserved specimens 

representing selected species and maintained for validating taxonomic identifications and documenting spatial 

and temporal distributions.

Wadeable—From a field-sampling perspective, a stream 

that can be safely waded, allowing effective deployment 

and performance of sampling gear (for example, stream 

depth does not exceed the height of the sampler opening 

or net). In situations where wadeable conditions do not 

exist, nonwadeable sampling methods are used.
 Glossary X

Woody snag—A submerged segment of wood (branch or 

log) having a minimum diameter of 1 cm and colonized 

by aquatic organisms. Organic (for example, leaf packs) 

and mineral debris also might be associated with woody 

snags; also known as snag or submerged woody debris.
Abstract 1

ABSTRACT

Algal, invertebrate, and fish communities 

are characterized as part of ecological studies in 

the U.S. Geological Survey’s National 

Water-Quality Assessment Program. Information 

from these ecological studies, together with 

chemical and physical data, provide an integrated 

assessment of water quality at local, regional, and 

national scales. Analysis and interpretation of 

water-quality data at these various geographic 

scales require accurate and consistent application 

of sampling protocols and sample-processing 

procedures. This report revises and unifies into a 

single document the algal, invertebrate, and fish 

community sampling protocols used in the 

National Water-Quality Assessment Program.

INTRODUCTION

Overview of the NAWQA Program

The U.S. Geological Survey’s (USGS) National 

Water-Quality Assessment (NAWQA) Program is a 

perennial program designed to provide a comprehensive, interdisciplinary water-quality assessment of the 

Nation’s flowing water resources (Hirsch and others, 

1988; Leahy and others, 1990). During the Program’s 

first decade of operation (1991–2001), ecological studies were conducted to assess the occurrence and distribution of algal, invertebrate, and fish communities in 

about 59 study units (Gilliom and others, 1995). These 

study units represented the dominant hydrologic systems nationwide and are staggered in time with respect 

to implementation, high-intensity, and low-intensity 

sampling periods (Gilliom and others, 1995). In the 

second decade of the Program (2001–2011), ecological 

studies will be conducted at selected fixed sites to provide long-term-trends monitoring. Ecological studies 

also are part of nationally guided studies addressing 

selected water-quality issues.

Accurate and consistent guidance and application of field-sampling and processing protocols are 

central to the capability of the NAWQA Program to 

analyze and interpret the physical, chemical, and biological attributes of aquatic systems at local, regional, 

and national scales. Protocols were developed or 

adapted for sampling algal, invertebrate, and fish communities during the first decade of the Program (see 

Cuffney and others, 1993; Meador and others, 1993; 

Porter and others, 1993). Although these protocols 

were tested during pilot studies (for example, Cuffney 

and others, 1997) and implemented by study units at 

fixed sites, they also were applied extensively in 

NAWQA synoptic studies (for example, Sorenson and 

others, 1999) and USGS District cooperative programs. 

Field experience acquired during the nearly 

decade-long application of these protocols has resulted 

in the need to revise the protocols to improve clarity 

and consistency in their use, and to reflect new program practices.

Purpose and scope

The purpose of this report is to provide revised 

protocols for sampling algal, invertebrate, and fish 

communities as part of the NAWQA Program. The 

revised protocols describe required methods for collecting and processing biological samples. When the 

application of required methods is not possible, appropriate alternative methods are used following discussion and approval by NAWQA regional and national 

biologists. Where appropriate, sampling and processing procedures are presented in a concise, stepwise 

format to promote consistent interpretation and application of methods by NAWQA sampling teams. Important procedural or safety information is italicized. Tip 

boxes are used to convey helpful information that was 

derived from experiences of NAWQA sampling teams. 

Revised Protocols for Sampling Algal, Invertebrate, and Fish 

Communities as Part of the National Water-Quality Assessment 

Program

By Stephen R. Moulton II, Jonathan G. Kennen, Robert M. Goldstein, and Julie A. Hambrook
2 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

Quality assurance/quality control (QA/QC) and safety 

guidance are provided where appropriate.

This report revises the original NAWQA protocols for algae (Porter and others, 1993), invertebrates 

(Cuffney and others, 1993), and fish (Meador and others, 1993). A revision of NAWQA habitat protocols 

was published by Fitzpatrick and others (1998). The 

revised protocols also may be adapted for use in studies 

by the USGS District cooperative program and other 

governmental agencies. Investigators evaluating the use 

of these revised NAWQA protocols also should consult 

additional sampling protocol discussion found in Britton and Greeson (1987), Lazorchak and others (1998), 

and Barbour and others (1999).

Summary of revisions to original protocols

Several additions, deletions, and modifications 

have been made to the original field protocols. The 

major changes are listed as follows:

Changes to the entire set of original protocols:

1. Added a glossary of terms frequently used in the 

protocols.

2. Added a checklist of sampling gear and commonly used equipment and supplies.

3. Added guidance to prevent the translocation of 

aquatic nuisance species (for example, zebra 

mussels) and parasites (for example, whirling disease).

4. Added guidance on the completion of field data 

sheets and post-field activities related to the Biological Transactional Data Base (Bio-TDB).

5. Revised the procedure and explanation for 

creating the alphanumeric sample identification 

code.

Changes to the algal protocols:

1. Added collection of light measurements in conjunction with algal sampling.

2. Added requirement to prepare microalgal subsamples for chlorophyll a and ash-free dry 

mass/carbon analysis from each richest-targeted 

habitat periphyton sample or phytoplankton sample collected.

3. Added photographs and flow chart to illustrate 

sample-collection and processing procedures.

4. Revised the algal sample identification code by 

adding a 17th character to identify the sample bottle/filter sequence and dropping the letter designation for a replicate microalgal sample.

5. Dropped the collection of mosses as a sample 

component of qualitative multihabitat samples.

Changes to the invertebrate protocols:

1. Added photographs and flow chart to illustrate 

sample-collection and processing procedures.

2. Dropped the collection of a depositional 

targeted-habitat sample type.

3. Changed sampling and field-processing mesh 

size to 500 μm for all samples.

4. Revised guidance on field processing of samples 

and the formation of sample components.

Changes to the fish protocols:

1. Added requirement for examination of deformities, eroded fins, lesions, and tumors as external 

anomalies of fish.

2. Emphasized use of fish quality-assurance (QA) 

report by Walsh and Meador (1998).

3. Established a standard seine mesh size of 6.4 mm 

(0.25 in) to provide a consistent minimum capture size.

4. Dropped requirement for measuring standard 

length.

Acknowledgments

We thank Thomas Cuffney (Raleigh, North 

Carolina), Martin Gurtz (Reston, Virginia), Michael 

Meador (Reston, Virginia), and Stephen Porter (Lakewood, Colorado) for their work to develop the original 

protocols for sampling algal, invertebrate, and fish 

communities; much of their original guidance has been 

used or modified in this report. The revision of these 

protocols would not have been possible without the 

contributions and experiences of these individuals and 

many other USGS personnel who applied the original 

protocols in a wide range of stream types across the 

Nation during the first decade of the Program. The fol-
Field Protocols 3

lowing USGS personnel (in alphabetical order) provided helpful information and comment during the 

preparation of this revision: Debbie Adolphson 

(Urbana, Illinois), Robert Anderson (Pittsburgh, Pennsylvania), Karen Beaulieu (East Hartford, Connecticut), Michael Bilger (Lemoyne, Pennsylvania), Larry 

Brown (Sacramento, California), Carmen Burton (San 

Diego, California), James Coles (Northborough, Massachusetts), Carol Couch (Reston, Virginia), Steven 

Goodbred (Sacramento, California), Evan Hornig 

(Austin, Texas), Ronald Kuzniar (Lakewood, Colorado), Terry Maret (Boise, Idaho), Karen Murray (Troy, 

New York), David Peterson (Cheyenne, Wyoming), 

Peter Ruhl (Reston, Virginia), Barbara Scudder (Middleton, Wisconsin), Clyde Sholar (Louisville, Kentucky), Terry Short (Menlo Park, California), Stephen 

Smith (Reston, Virginia), Ian Waite (Portland, Oregon), 

Holly Weyers (Dover, Delaware) and Humbert Zappia 

(Montgomery, Alabama).

Photographs used in this report were provided 

courtesy of Mark Ayers (USGS, West Trenton, New 

Jersey), Barry Biggs (National Institute of Water and 

Atmospheric Research, Christ Church, New Zealand), 

Carmen Burton, Richard Frehs (USGS, Columbus, 

Ohio), Julie Hambrook (USGS, Columbus, Ohio), 

Jonathan Kennen (USGS, West Trenton, New Jersey), 

USGS Leetown Science Center (Leetown, West Virginia); Michael Mac (USGS, Reston, Virginia), 

Michael Meador, Stephen Porter, Stephen Smith, Jason 

Sonneman (Monash University, Australia), Dennis Sun 

(USGS, West Trenton, New Jersey), and Wildlife Supply Company (Buffalo, NewYork).

This report was reviewed by Barbara Scudder 

(USGS, Middleton, Wisconsin) and Mitchell Harris 

(USGS, Urbana, Illinois). Editorial reviews were provided by Jon Raese (USGS, Lakewood, Colorado).

FIELD PROTOCOLS

1. OVERVIEW OF THE SAMPLING APPROACH

Ecological studies characterize benthic algae and 

invertebrate or fish communities in relation to chemical 

and physical characteristics at sites investigated for 

NAWQA ecological studies. Biological sampling and 

habitat assessments are conducted within a defined 

length of stream, or "reach," which provides a common 

spatial scale on which to relate biotic and abiotic factors that influence water quality (Cuffney and others, 

1993; Fitzpatrick and others, 1998)

1.1 Establishing the sampling reach

The sampling reach is established at a site by 

using a combination of geomorphic characteristics, 

such as stream width, stream depth (wadeable or nonwadeable), geomorphology (type and distribution of 

geomorphic channel units), and local habitat disturbance (Fitzpatrick and others, 1998). Although much 

of the sampling effort is concentrated in one primary 

reach, multiple sampling reaches (typically three) are 

established at certain sites to determine variability 

among reaches and to provide alternative sampling 

reaches should the primary reach become inaccessible. 

After reaches are established, their locations and identities (for example, letter designation A, B C, and so 

on) do not change. Sampling crewmembers (hereafter, 

"crewmembers") should consult Fitzpatrick and others 

(1998) for specific guidance on establishing sampling 

reaches. In general, reach length ranges from 150 to 

300 m for wadeable streams and from 500 to 1,000 m 

for nonwadeable streams (Fitzpatrick and others, 1998; 

Meador and others, 1993). Photographs should be 

taken at the reach to document instream and riparian 

physical characteristics, especially on the day of biological sampling.

1.2 Types of samples collected

Quantitative, semi-quantitative, and qualitative 

sample types are collected in the NAWQA Program to 

characterize algal and invertebrate communities within 

a sampling reach. A representative sample of the fish 

community is collected from the entire reach. Depending on study objectives, one or more different algal or 

invertebrate sample types might be collected in the 

reach.

Quantitative algae or semi-quantitative invertebrate samples are collected from a known area of substrate to provide estimates of abundance of algal or 

invertebrate taxa present in a targeted habitat (for 

example, riffle, woody snag, or pool). Two contrasting 

types of targeted habitats are defined in the NAWQA 

program: a richest-targeted habitat (RTH) and a 

depositional-targeted habitat (DTH). The RTH represents that habitat in the reach, usually erosional, where 

maximum taxa richness is likely to be observed. For 

example, invertebrate RTH samples typically are col-
4 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

lected from riffles in moderate to high gradient streams 

dominated by coarse-grained substrates; in low-gradient streams dominated by fine-grained substrates, 

invertebrate RTH samples often are collected from 

woody snags. Separate discrete collections of algae and 

invertebrates are taken in the RTH and then combined 

into a composited RTH algae or invertebrate sample. 

The DTH represents that habitat in the reach where 

sediments (for example, silt and sand) are deposited. 

Discrete collections of algae are taken in the DTH and 

then combined into a composited DTH sample. The 

DTH sample is not included as part of the invertebrate 

sampling protocols. Additional guidance and explanation of the targeted habitat samples is presented in the 

algal and invertebrate protocol sections of this report.

A seston sample [or phytoplankton (PHY), cf. 

Porter and others, 1993] represents a third type of 

quantitative algal sample that might be collected 

depending on study objectives. Seston sampling usually is conducted in conjunction with routine 

water-chemistry sampling because of the specialized 

sampling gear (for example, a D-96 sampler) used to 

collect whole-water samples.

Qualitative algal and invertebrate samples may 

be collected from a reach to document the presence of 

other algal or invertebrate taxa in addition to those taxa 

found in the targeted habitat samples (RTH and DTH). 

Separate discrete collections of algae and invertebrates 

are taken from each of the instream habitats present in 

the reach and then combined into a composited qualitative multihabitat (QMH) sample. Additional guidance 

and explanation of the QMH sample is presented in the 

algal and invertebrate protocol sections of this report.

1.3 Identifying instream-habitat types for sampling

Habitat types within each stream reach are categorized according to a hierarchical framework that 

includes three tiers of organization to promote sampling comparability among the diversity of streams 

across the Nation (table 1). These tiers are (1) major 

geomorphic channel units, (2) major channel boundaries, and (3) major channel features (Cuffney and others, 1993). This framework is used to guide where 

certain types of invertebrate and algae samples are collected in the reach of wadeable and nonwadeable 

streams. Tiers 1 and 2 are used to determine sampling 

locations for algae and invertebrates. However, there 

are differences in the terminology used to identify 

major channel features (Tier 3) based on common practices in the sampling disciplines for algae and invertebrates. For example, the habitat where an invertebrate 

collection is taken may be described as a "riffle-main 

channel-natural bed habitat." Similarly, the same habitat where an algal collection is taken may be described 

as a "riffle-main channel-epilithic habitat."

Except for qualitative sampling, areas downstream of obstructions (for example, bridge abutments 

and storm water outflow pipes) and along the edges of 

atypical habitat features (for example, island margins) 

are avoided because they might bias the quantitative 

sample. Specific guidance for determining the sampling habitats for algae, invertebrates, and fish are provided in the respective protocol section of this report.

1.4 Factors determining when to sample

Determining when to sample algal, invertebrate, 

and fish communities requires careful consideration of 

several factors, especially when sampling is performed 

annually. Some of these factors (listed as follows) are 

interrelated and might conflict with one another:

• historical sampling records 

• life history attributes of the community being sampled [for example, development of aquatic insects 

or spawning/migratory patterns of anadromous fish 

or threatened and endangered (T&E) species]

• current and historical hydrological conditions (for 

example, stage, flood recurrence interval)

• seasonal human recreational activities (for example, rafting)

• seasonal agricultural practices (for example, application of pesticides)

• access to sites (for example, road conditions, permission to cross or sample on private property)

• availability of crewmembers and technical specialists (for example, fish taxonomist)

Many of these factors are difficult or impossible 

to control or predict; therefore, the sampling period 

ultimately represents a compromise of the most important factors. In nearly all cases, historical sampling 

periods based on optimal hydrological conditions to 

permit safe and effective sampling and organism life 

cycles are the primary determinants of when to sample 

biological communities. Depending on the study objectives, all sites are sampled within a defined index 

period (for example, May–June) to ensure year-to-year 

data comparability (Cuffney and others, 1993).
Field Protocols 5

1.5 Coordination of field activities

Biological sampling, habitat assessment, and 

water-quality measurements (for example, 

dissolved-oxygen concentration, conductivity, and pH) 

are often performed during one or more visits to a sampling site. Consequently, coordination among crewmembers and operations is desirable when possible to 

minimize sampling interferences (for example, physical disturbance of the reach) and the time spent working at a site. However, close coordination of sampling 

operations may not always be achieved because of differing sampling requirements for various studies. 

Regardless of the approach taken to achieve coordination, disturbance to instream habitats of the reach by 

crewmembers must be avoided prior to sampling.

Because activities related to habitat assessment 

and fish sampling (for example, upstream progression 

of an electrofishing team or kick seining) can disturb 

substantially instream habitats of the reach, they are 

performed at a later date after algal and invertebrate 

sampling. Doing this also allows fish species to return 

to their preferred habitats in the reach. Before any samples are collected, crewmembers must identify the 

instream habitat types present in the reach, from which 

series of discrete samples are collected and composited 

according to sample type. Algal and invertebrate samples are collected within the reach beginning with the 

most downstream sampling location and progressing 

upstream. When sampling multiple reaches in a stream 

segment, sampling is completed within the 

downstream-most reach before proceeding to the next 

reach upstream. RTH and DTH samples are collected 

before QMH samples to minimize disturbance of the 

targeted habitats.

2. PRE-FIELD ACTIVITIES

The collection of high-quality biological samples 

and resulting data is dependent on careful planning and 

preparation prior to the actual visit to a study site. 

Tier 1

Major geomorphic

channel units Description

Riffle Turbulent flow; shallow, coarse-grained substrate

Run Laminar flow, less turbulent; variable depth and substrate

Pool Very low current velocity; relatively deep, depositional accumulations of fine sediment 

particles

Tier 2

Major channel

boundaries Description

Channel Flow paths associated with the main and secondary river channels

Channel margin Subhorizontal, fluvial areas associated with the streambanks

Island margin Subhorizontal, fluvial areas associated with the banks of islands

Tier 3

Major channel

feature (habitat) Description

Algae Invertebrates

Epilithic Natural bed Natural bed materials without extensive macrophyte beds

Manufactured bed Revetments, riprap, other manufactured or artificial bed materials

Epipsammic & Epipelic

Bar Shallow, gently sloping sand or gravel bars primarily associated with channel edges or 

major changes in water velocity

Slough Remnants of abandoned river channels that connect with the main channel even at normal low flows

Epidendric Woody snag Trees, branches, other woody debris of terrestrial origin that extend into the water column

Epiphytic Macrophyte bed Growths of emergent or submergent aquatic macrophytes

Table 1. Hierarchical framework for geomorphic descriptors used to determine habitats from which algal and invertebrate 

samples are collected (modified from Cuffney and others, 1993).
6 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

Pre-field activities include preparing a sampling plan, 

obtaining collecting permissions and field supplies, and 

ensuring that sampling gear is in proper working order. 

Sampling team crewmembers also should familiarize 

themselves with the sample-tracking functions of the 

NAWQA Biological Transactional Data Base 

(Bio-TDB).

2.1 Preparing a sampling plan

Prior to conducting any fieldwork, sampling 

teams are encouraged to prepare a sampling plan. A 

sampling plan is a tool that might be used to define the 

sampling objectives, describe the roles and responsibilities of crewmembers, list sampling methods and procedures, and define QA/QC activities appropriate for 

the study. The sampling plan should describe when 

fieldwork would be performed, including a process for 

determining alternative plans in the event of adverse 

weather or hydrological conditions. All members of a 

sampling team should be familiar with the standard 

protocols described in this report for collecting and 

processing algal, invertebrate, and fish samples. Training in the application of these protocols can be 

obtained by attending periodic nationally or regionally 

organized courses, or by mentoring opportunities 

among the sampling teams and the regional and 

national NAWQA biologists. Although these protocols 

are intended to maximize consistency among the algal, 

invertebrate, and fish community samples collected in 

the NAWQA Program, there might be occasions when 

these standard protocols cannot be effectively applied 

or study objectives warrant alternative sampling or processing protocols. Alternative protocols are reviewed in 

consultations with the Regional Biologist.

2.2 Collecting permits

The sampling team leader is responsible for 

obtaining collecting permits prior to sampling. Permission from a landowner is also required when sampling 

on or crossing private property. In some cases, multiple 

permits might be required to satisfy private, State, and 

Federal (for example, National Park Service or U.S. 

Fish and Wildlife Service) regulations. Walsh and Meador (1998) provide a comprehensive summary of state 

permitting agencies and their reporting conditions. 

Normally, permits are required only to collect fish and 

other aquatic vertebrates. However, some states also 

require a collecting permit or fishing license for invertebrates (for example, Arkansas and New York). Sampling team leaders also must inquire about and be 

familiar with applicable conditions regulating collection gear and methods, specimen-holding requirements, and number of specimens sacrificed and 

preserved for study; compliance with all permit conditions is required. A brief list of possible requirements 

that should be considered follows:

• Prepare a study plan in advance of sampling

• Document species targeted in sampling

• Document state and federally protected species 

having special concern, threatened and endangered 

(T&E) status that might be encountered (see Walsh 

and Meador, 1998)

• Obtain a sport-fishing license if necessary

• Notify permitting agency prior to conducting field 

work

• Submit a periodic (typically annual) summary 

report describing all sampling activities and listing 

the species and numbers collected. (The report format is usually specified and might consist of an 

electronic data form provided by the permitting 

agency.)

• Provide copies of research reports or publications 

acknowledging the permitting agency

2.3 Supplies and chemicals

The following supplies and chemicals are used to 

collect and field-process algal, invertebrate, or fish 

community samples in the NAWQA Program. Sampling teams should ensure that reserve quantities of 

supplies and replacement parts (for example, collection 

nets) are maintained during extended sampling trips. 

Although most supplies can be purchased from local 

retailers (for example, hardware and sporting goods 

stores), some supplies can be purchased only from 

retailers specializing in field or laboratory products 

(see Walsh and Meador, 1998, p. 23).
Field Protocols 7

Sampling Gear

Algae

❑ SG-92 with brushes (see section 4.3.1)

❑ Gravel sampler with large masonry trowel (see section 4.3.1)

Invertebrates

❑ Slack sampler with 500-μm mesh net (see section 5.3.1)

❑ Grab sampler (for example, petite Ponar or Ekman)

❑ D-frame kick net with 500-μm mesh net

Fish

❑ Backpack (with ammeter) or towed electrofishing unit

❑ Electrofishing boat

❑ 3 x 1.2-m seine (6.4-mm mesh)

❑ 7.6 to 9.6 x 1.2-m seine (6.4-mm mesh)

❑ 30 to 61 x 1.8-m seine (6.4-mm mesh)

Water-Quality Meters

❑ Current velocity meter

❑ Dissolved oxygen meter

❑ Thermometer

❑ Conductivity meter

❑ pH meter

❑ Light meter and quantum sensor

Basic Field Supplies

❑ Wire mesh sieves (20-cm and 30-cm diameter, 500-μm mesh)

❑ White buckets (19 L)

❑ Forceps, blunt and fine tipped

❑ Tongs

❑ Knives (pocket and putty), scalpels, scissors

❑ Spatula (without holes)

❑ Plastic petri dishes (47-mm diameter)

❑ OxfordTM Macro-Set hand pipettor (1 mL–5 mL w/tips)

❑ Heavy-duty aluminum foil

❑ Hand-held electric stirrer (periphyton homogenizer)

❑ Batteries (various sizes including 12 volt)

❑ WhatmanTM GF/F glass fiber filters (25-mm and 47-mm diameter)

❑ Hand-operated vacuum pump with pressure gauge

❑ Plastic Erlenmeyer flask (1 L)

❑ Filter funnel and base (for 25 mm and 47 mm filters)

❑ Hand saw or lopping shears

❑ White enamel or plastic trays

❑ Hand brush

❑ Dice

❑ Gas can with pre-mixed fuel

❑ Electrofishing battery charger

❑ Dip nets

❑ Portable aeration system

❑ Live cage for holding fish

❑ Small aquarium nets

❑ Small wooden club 

❑ Syringes with locking hypodermic needles

❑ Resealable plastic bags, various sizes

❑ Arm-length rubber gloves (per person)

❑ Surgical gloves

❑ Chemical splash goggles

❑ Respiratory mask with formaldehyde filter

❑ Heavy-duty nylon thread and sewing needles

❑ Hand rake

❑ Squirt bottles

❑ Wide-mouth NalgeneTM sample bottles 

(60, 125, 250, 500 mL, and 1 L)

❑ Plastic scintillation vials

❑ Alcohol/waterproof pens, black

❑ Pencils (lead, red wax)

❑ Waterproof paper

❑ Field data sheets printed on waterproof paper

❑ Measuring board

❑ Ruler and tape measure

❑ Large plastic insulated coolers

❑ Graduated cylinders (glass 10 mL, plastic 50–500 mL)

❑ Hanging scale

❑ Portable weighing balance

❑ First aid kit

❑ Insect repellant

❑ Sun screen

❑ Antibiotic liquid soap

❑ Polarized sunglasses

❑ Cellular phone

❑ Flashlight and lantern

❑ Safety plan (with emergency phone numbers)

❑ Collecting permits

❑ Hip boots (per person)

❑ Chest waders (per person)

❑ Wader repair kit

❑ Rain gear

❑ Personal flotation devices (per person)

❑ Digital camera (high-resolution)

❑ Geographic positioning system

❑ Field guides (plants, fish, insects)

❑ Folding table and chairs

❑ Braided, 1.9-cm nylon rope

❑ Trash can

❑ Glue gun

❑ Tool set

❑ Paper towels

Chemicals

❑ 37-percent buffered formaldehyde (buffered to pH 7)

❑ 10-percent buffered formalin

❑ 1-percent bleach solution

❑ Anesthetic (for example, CO2 tablets, clove oil)

❑ Dry ice

❑ Water

Basic Shipping Supplies

❑ Plastic tape (for example, 3M Scotch BrandTM 471)

❑ Resealable plastic bags, various sizes

❑ Heavy-duty trash bags

❑ Absorbent packing material

❑ Bubble packing sheets

❑ Ice chests

❑ Transparent packaging tape

❑ Shipping paperwork
8 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

2.4 Sample preservatives

Proper preservation of biological samples is paramount to obtaining high-quality taxonomic and ecological data about organisms. Samples are preserved in 

the field to prevent the breakdown of structural proteins, thereby maintaining the integrity of tissues and 

other morphological structures for taxonomic study. 

Many morphological structures and color patterns are 

destroyed when samples are poorly preserved, thereby 

resulting in undesirable levels of taxonomic resolution 

and quality of material for vouchering or referencing. 

Before working with sample preservatives, crewmembers must put on personal protective equipment (for 

example, safety glasses and gloves) and ensure that 

work is performed in a ventilated area (for example, an 

outdoor field processing area or laboratory fumehood).

Formaldehyde, whether concentrated 

(37-percent stock = 100-percent formalin) or diluted 

(for example, 10-percent formalin), is the required preservative for algal, invertebrate, and fish samples in the 

NAWQA Program. Algal samples are preserved by 

adding a sufficient volume of buffered formaldehyde to 

the sample to obtain a final concentration of 5-percent 

buffered formalin (see Algal Sampling Protocols). 

Invertebrate and fish samples are preserved with 

10-percent buffered formalin. Buffering formaldehyde 

or formalin to pH 7 with borax minimizes shrinkage 

and decalcification of tissues, especially mollusk 

shells. It is more economical to prepare 10-percent 

buffered formalin rather than purchase this concentration from a supply company.

Preparing a 1-L solution of 10-percent buffered 

formalin:

1. Add 100 mL of formaldehyde to 900 mL of water 

in a chemically resistant, nonbreakable bottle.

2. Add about 3 g of borax to 10 mL of water and 

mix.

3. Add dissolved borax solute to buffer formalin 

solution.

4. Tightly seal the bottle and mix by carefully 

inverting the bottle several times.

5. Label the outside of the bottle with "10-percent 

buffered formalin," the date of preparation, and 

related hazardous chemical stickers.

2.5 Sample identification and tracking

Algae, invertebrate, and fish community samples 

collected as part of NAWQA Program studies are 

tracked by using an alphanumeric sample identification 

code (SMCOD). The SMCOD is a smart code because 

it succinctly summarizes key information about a sample (table 2), and if created properly, it will uniquely 

identify each bottle or filter associated with a particular 

sample. Variables associated with each of the SMCOD 

character fields are explained in figure 1 for the different sample media. 

Sample numbers are assigned consecutively to 

all of the different samples in each sample medium collected each year. The following example shows how 

samples are numbered for a 2-year study at a site:

Sampling Year 1

Sample Medium Sample Number Sample Type

Algae 0001 RTH

Algae 0002 DTH

Algae 0003 QMH

Invertebrates 0001 RTH

Invertebrates 0002 QMH

Fish 0001 Not applicable

Sampling Year 2

Sample Medium Sample Number Sample Type

Algae 0001 RTH

Algae 0002 DTH

Algae 0003 QMH

Invertebrates 0001 RTH

Invertebrates 0002 QMH

Fish 0001 Not applicable

Sample identification code character fields

1–4 5–8 9 10 11 12–15 16 17

Algae

Study Unit 

abbreviation

Collection date 

(MMYY)

Sample 

medium

Sample 

type

Subsample

(analysis type) Sample 

number

Sample componentBottle/filter

sequence

Invertebrates Sample 

component

Bottle 

sequence –

Fish Sample medium – –

Table 2. Definition of character fields used to create sample identification codes for algae, invertebrates, and fish 

samples in the National Water Quality Assessment Program

[MMYY, month and year; –, not applicable]
Field Protocols 9

STUDY UNIT:

4–letter abbr.

DATE:

MMYY

SAMPLE NUMBER:

0001–9999

ALGAE SAMPLE COMPONENT:

 A = macroalgae

 B = microalgae

 S = split sample, microalgae

SAMPLE MEDIUM:

 A = algae

SAMPLE MEDIUM:

 I = invertebrate

ALGAL ANALYSIS TYPES:

 E = identification and enumeration

 C = chlorophyll a

 M = ash-free dry mass

ALGAL SAMPLE TYPES:

 R = RTH

 D = DTH

 Q = QMH

 P = phytoplankton

LINJ 0602 IRM 0001 A

LINJ 0602 FSH 0001

LINJ 0602 ARE 0001 B A ALGAE BOTTLE/FILTER SEQUENCE:

A–Z

INVERTEBRATE SAMPLE COMPONENTS:

 M = main-body

 L = large-rare

 S = split

INVERTEBRATE SAMPLE TYPES:

 R = RTH

 Q = QMH

INVERTEBRATE BOTTLE SEQUENCE:

A–Z

SAMPLE MEDIUM:

FSH–fish

Figure 1. Sample identification code used to distinguish algal (top), invertebrate (middle) and fish (bottom) samples in the National 

Water-Quality Assessment Program (modified from Cuffney and others, 1993 and Porter and others, 1993). abbr, abbreviation; MMYY, 

month/year; RTH, richest-targeted habitat; DTH, depositional targeted habitat; QMH, qualitative multihabitat; SMCOD, sample 

identification code.
10 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

2.5.1 Sample logs

Sampling teams are encouraged to maintain sample logs for each sample medium (algae, invertebrates, 

and fish) to provide an accounting of the samples collected in a study and to track the sample numbers that 

are used during a given year. The minimum information for each log should include:

• Study unit identification

• Log year

• Station identification code

• Reach designation

• Collection date (MM/DD/YYYY)

• Sample type (except for fish)

• Sample number

• Replicate flag (except for fish)

An algae sample log for the Long Island-New 

Jersey (LINJ) NAWQA study unit listing the samples 

collected at two study sites in one year might look like 

the following:

In this example, three algal sample types (RTH, 

DTH, and QMH) were collected in each of three sampling reaches (A, B, and C) at station 12345678 (total 

of nine algal samples). Note that the samples were 

numbered consecutively and that no replicate samples 

were collected. At station 87654321, four algal samples were collected from reach A only; the sample 

types collected include primary RTH, DTH, and QMH 

samples and a RTH repeated sampling replicate (sample number 0011).

2.5.2 Sample labels

Sampling teams are encouraged to prepare formatted sample labels (fig. 2) before going into the field 

so that sample information can be recorded neatly on 

the label immediately after a sample is collected and 

processed. Sheets of formatted sample labels should be 

printed or photocopied on waterproof paper (for example, "Write-in-the-Rain"TM paper). Information is 

recorded on the labels using pens (for example, RapidographTM and Pigma PenTM) containing water- and 

formalin-resistant ink. Field sample labels are placed 

on the outside of the sample bottles for each sample 

medium. An identical label is placed inside each bottle 

for invertebrate and fish samples. Do not place labels 

inside algae sample bottles because algal cells might 

adhere to the label and compromise the integrity of the 

sample.

2.6 Sampling gear maintenance

Sampling gear must always be maintained to 

ensure proper working order. The following maintenance checks should be performed before, during, and 

after each sampling event.

• Inspect metal sieve screening and nylon-collection 

nets for holes and tears. Use a black pen to mark 

the location of numerous small holes. Apply silicon adhesive or hot glue to both sides of a net or 

sieve to close small holes. Alternatively, a 

wood-burning tool can also be used to melt and 

close nylon netting surrounding small holes. Use 

heavy-duty needle and thread to close large tears. 

Replace the sieve or collection net if it is beyond 

repair. Store nets out of direct sunlight to dry when 

not in use; ultraviolet radiation from the sun and 

prolonged dampness promotes deterioration of 

nylon netting.

• Clean, inspect, and lubricate moving gear parts (for 

example, the jaw mechanism on grab samplers) 

and cables. Never lubricate sampling gear when the 

gear will also be used to collect samples for chemical analysis. 

• Rinse sand from threaded gear parts to ensure 

proper fitting and minimize leakage. Also rinse 

sieves and buckets to prevent jamming when they 

are stacked.

• Inspect and calibrate water-quality meters according to standard protocols described in Wilde and 

Radtke (1998). Document maintenance and calibration records on all sampling gear and meters for 

QA purposes.

LINJ Algae Sample Log — Year 2002

Station ID Reach Collection 

Date

Sample 

Type

Sample 

Number

Replicate 

Flag

12345678 A 06/15/2002 RTH 0001 No

12345678 A 06/15/2002 DTH 0002 No

12345678 A 06/15/2002 QMH 0003 No

12345678 B 06/16/2002 RTH 0004 No

12345678 B 06/16/2002 DTH 0005 No

12345678 B 06/16/2002 QMH 0006 No

12345678 C 06/17/2002 RTH 0007 No

12345678 C 06/17/2002 DTH 0008 No

12345678 C 06/17/2002 QMH 0009 No

87654321 A 06/20/2002 RTH 0010 No

87654321 A 06/20/2002 RTH 0011 Yes

87654321 A 06/20/2002 DTH 0012 No

87654321 A 06/20/2002 QMH 0013 No
Field Protocols 11

NAWQA Algae Sample

SU ID:__________Station name:_____________________________________

Station code:____________________________Reach:___________________

Collection date:____________________ Time:_____________

Collectors:________________________________________________________

Sample type (circle one): RTH / DTH / PHY / QMH

Sample component (circle one): Macroalgae / Microalgae

Subsample (circle one): ID / CHL A / AFDM / POC

Sample number:__________ Bottle/filter sequence identifier:______________

Total area of periphyton sample:__________________cm2

Sample volume:__________; Subsample volume:__________mL

Preservative: buffered formaldehyde; Preservative volume:__________mL

Field SMCOD (optional):____________________________________________

NAWQA Invertebrate Sample

SU ID:__________Station name:_____________________________________

Station code:______________________________Reach:__________________

Collection date:____________________Time:_____________

Collectors:__________________________________________________

Collection mesh: 500 μm; Preservative: 10% buffered formalin

Sample type (circle one): RTH / QMH Split ratio___________

Sample component (circle one): Large-rare / Main-body / Split

Sample number:_____________ Bottle sequence identifier:________________

Field SMCOD (optional):____________________________________________

NAWQA Fish Sample

SU ID:__________Station name:_____________________________________

Station code:______________________________Reach:__________________

Collection date:____________________Time:_____________

Collectors:________________________________________________________

Sampling method:_____________________ Preservative: 10% buffered formalin

Sample number:_____________

Field SMCOD (optional):____________________________________________

Species:__________________________________________________________

Taxonomic specialist:__________________________ID date:______________

State:_________________County:____________________________________

Latitude:________________________Longitude:________________________

USGS quad name:_________________________________________________

Figure 2. Examples of field sample labels used to uniquely identify algae (top), invertebrate 

(middle), and fish (bottom) samples in the National Water-Quality Assessment Program. SU 

ID, study unit abbreviation; RTH, richest targeted habitat; DTH, depositional targeted 

habitat; PHY; phytoplankton; QMH, qualitative multihabitat; SMCOD, sample identification 

code; NAWQA, National Water-Quality Assessment; USGS, U.S. Geological Survey; ID, 

identification; CHL A, chlorophyll a; AFDM, ash-free dry mass; mL, milliliter; μm, 

micrometer; %, percent.
12 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

There is potential for cross-contaminating samples or translocating organisms among sampling sites 

or drainages when sampling gear and field-processing 

equipment are not adequately cleaned between samples 

or sites. Many invertebrates have claws and body hooks 

that allow them to be easily retained on the fine mesh 

netting, seams or crevices of sampling gear or 

field-processing equipment. Algae are particularly susceptible for cross-contamination because of their 

microscopic size (for example, diatoms). Filamentous 

algae (for example, Cladophora) might promote 

cross-contamination because they represent either a 

microhabitat or an entrapment mechanism for invertebrates. Translocation of organisms relates to the introduction of nonindigenous or aquatic nuisance species 

into areas where they might cause substantial environmental, socioeconomic, or public health damage. Several nuisance species are well documented for plants 

(for example, Eurasian watermilfoil), animals (for 

example, zebra mussel), and disease-forming spores 

(for example, whirling disease parasite found in salmonids).

Cleaning sampling gear and equipment:

• After collecting and field processing each sample, 

inspect the sampling gear and net surface including 

all seams and crevices; remove debris and place 

attached organisms in the sample.

• Vigorously wash both sides of a net and inspect all 

seams and crevices; discard any remaining debris 

or organisms.

• Scrub and wash all sieves, dishpans, buckets, and 

brushes used to process samples. Closely inspect 

sieves for trapped organisms (especially tubificid 

worms and midges).

• After sampling at each site, wash mud from all 

sampling gear (including boats and electrofishing 

gear), field-processing equipment and waders, and 

spray with a 1-percent bleach solution.

Preparing a 1-L solution of 1-percent bleach:

1. Add 1 mL of bleach to 99 mL of water in a plastic 

spray bottle.

2. Label the outside of the bottle with "1-percent 

bleach" and the date of preparation.

3. SAFETY IN THE FIELD

Safety issues related to any aspect of scientific 

study deserve serious attention by all members of a 

sampling team. In particular, some field-sampling 

activities carry a real potential for personal injury or 

death. Sampling teams are encouraged to review field 

safety guidance presented by Yobbi and others (1996) 

and Lane and Fay (1997). All field-sampling activities 

must comply with official USGS safety guidelines 

(U.S. Geological Survey, 2001a) and memoranda. At a 

minimum, the team leader should work with the District safety officer to document that training sessions to 

promote safety awareness are conducted as appropriate 

for the sampling frequency. Approved training and dedicated application of safe operating procedures and 

maintenance for vehicles, trailers, watercraft, and sampling gear will minimize the potential for personal 

injury. Property damage, accidents, and injuries can be 

avoided by (1) wearing appropriate safety equipment, 

(2) handling chemicals safely, and (3) coping with 

common environmental hazards (for example, heat, 

thunderstorms, floods, poisonous plants, and venomous 

animals).

Sampling teams always must consist of at least 

two crewmembers; no one should sample alone. At 

least two crewmembers of each sampling team must be 

trained and currently certified in basic first aid, cardiopulmonary resuscitation techniques, and hazard 

communications. Each sampling team must carry a first 

aid kit, a cellular telephone, and an emergency contact 

list (including medical facilities closest to each sampling site) inside each field vehicle as part of the Job 

Hazard Analysis (JHA) for a particular study or sampling site. The JHA is prepared by the study team and 

reviewed by District management for the purpose of 

assigning and understanding responsibilities for safety 

in field and laboratory studies.

3.1 Personal and environmental safety

Crewmembers are responsible for wearing their 

personal protective equipment when it is appropriate to 

do so. Regular inspection of this equipment is necessary to ensure that it is in proper working condition. 

Waders, gloves, safety glasses or goggles, and facemasks are worn when dangerous objects (for example, 

scrap metal and broken glass), water-borne pathogens, 

toxic substances (including sample preservatives) are 

encountered at a sampling site. Crewmembers should 

ensure that vaccinations against water and soil-borne 

diseases are up-to-date. Hand sanitizing creams and 

bactericidal soap must be supplied to sampling teams.
Field Protocols 13

Personal-flotation devices (PFDs) must always 

be worn when working in or near the water regardless 

of hydrologic conditions except when a JHA documents approved exemptions. Sampling teams working 

at established USGS gaging stations must follow the 

guidelines established by the site-specific JHA for that 

station. Chest-wader suspenders can be worn on the 

outside of all clothing, including noninflating PFDs. 

However, inflatable suspender style PFDs must always 

be the outermost layer to allow for complete expansion 

of the PFD air bladder without injury to the wearer or 

damage to the bladder. Non-U.S. Coast Guard (USCG) 

flotation devices are allowed for wading, cableway and 

bridgework. Belts are sometimes worn to cinch 

loose-fitting waders. Wearing a belt traps air below the 

waist, slowing the onset of hypothermia in cold water, 

and also prevents waders from filling quickly if a crewmember falls in the stream.

Field sampling safety risks also include potential 

contact with dangerous plants and animals or exposure 

to adverse weather conditions. Crewmembers working 

at sites where dangerous wild animals (for example, 

bears, mountain lions, alligators, and poisonous 

snakes) might be encountered should receive animal 

awareness and avoidance training. This training should 

include (1) identification of the dangerous animals, (2) 

behavioral traits of the animals, (3) recognition of signs 

of animal activity, (4) how to avoid attracting or provoking the animals, and (5) how to react during 

encounters or if attacked. Training should be provided 

before work at such sites begins and at least every 3 

years thereafter. Crewmembers should be instructed on 

how to identify and avoid poisonous plants (for example, poison ivy and poison oak), and venomous arthropods (for example, wasps, spiders, and ticks) including 

first aid procedures. Prolonged exposure to the sun, 

wind, air, and water temperature extremes can result in 

varying degrees of skin irritation, discomfort, and 

injury (for example, frostbite), including hypo- and 

hyperthermia. Crewmembers should know the signs of 

heat stress, wear appropriate clothing and eyewear to 

protect and maintain comfort, and treat exposed skin 

with sun block and lip balm. Crewmembers periodically should stay out of direct sunlight or otherwise 

cool their bodies and stay hydrated by drinking plenty 

of water and electrolyte replacement drinks. Field sampling should be delayed or rescheduled when any form 

of severe weather is forecasted. Environmental safety 

training also should include (where applicable) procedures for hazards, such as floods, icy conditions, earthquakes, and fires.

3.2 Chemical safety

Crewmembers must receive appropriate occupational safety training before being assigned to hazardous duties. Unique or uncommon hazardous activities 

should be addressed separately as they are encountered. 

Crewmembers handling or using hazardous materials 

must receive initial Hazard Communication Training 

with refresher training every 3 years. Personal protective equipment appropriate for working with a particular chemical (for example, safety glasses or ventilation 

mask) and any emergency procedures must be part of 

Hazard Communication Training. A "Material Safety 

Data Sheet" (MSDS) for each chemical used in the 

field must be maintained in a field safety notebook containing a Chemical Hygiene Plan. This field safety 

notebook is kept in vehicles and mobile labs, and all 

crewmembers must be familiar with its contents. All 

chemical containers must be appropriately labeled with 

chemical name, concentration, date, and manufacturer 

hazard warnings. All secondary containers (for example, beakers, flasks, squirt bottles, and safety cans) 

must be labeled with the same information if the chemical is kept in the container past the immediate work 

shift or if the container is transported away from the 

immediate work area. Chemical formulas or nonspecific names, such as "Mixture A," are not acceptable 

identifiers. Even innocuous substances (for example, 

buffers and water) must be labeled and identified.

Crewmembers should practice good personal 

hygiene around hazardous materials, such as washing 

hands after handling chemicals, using required personal protective equipment, and not consuming or storing food and drink in areas where hazardous materials 

are used or stored. Coolers or other containers used for 

shipping samples must never be used for storing food 

and drink. Chemicals should be transported in their 

original U.S. Department of Transportation-approved 

shipping containers, according to State and Federal 

regulations. Ideally, chemicals should be transported in 

trailers or carriers attached to the vehicle and not inside 

the passenger area. Transporting chemicals in this manner minimizes the risk of exposure if a chemical bottle 

breaks or leaks.

Preservation of algal, invertebrate, and fish samples involves the use of aqueous buffered formaldehyde 

or 10-percent buffered formalin. Studies indicate that 
14 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

formaldehyde is a potential human carcinogen. The 

severity of irritation increases as formalin concentration increases. Airborne concentrations > 0.1 mg/L can 

cause irritation of the eyes, nose, and throat; at 

> 50 mg/L, formalin is acutely hazardous to life and 

might result in death. Dermal contact with formalin 

causes various skin reactions, including sensitization, 

which might force crewmembers so sensitized to seek 

other job duties. Specific hazard labeling is required for 

formaldehyde and formalin. Hazard labeling, including 

a warning that formaldehyde presents a potential cancer hazard, is required where formaldehyde levels, 

under reasonably foreseeable conditions of use, might 

potentially exceed 0.5 mg/L. These chemicals should 

be handled only in well-ventilated areas while wearing 

chemical-resistant gloves and approved eye protection. 

The use of formaldehyde in poorly ventilated areas 

might require the use of respirators and medical monitoring. Vehicles or mobile labs transporting and using 

formaldehyde or formalin must have eyewash solutions 

readily available.

3.3 Vehicles, boats, and sampling gear safety

All vehicle operators must be at least 18 years of 

age and have a valid driver’s license. Successful completion of USGS-approved driver education courses is 

required every 3 years. Operators of specialized vehicles (for example, all-terrain vehicles) should participate in training programs for these vehicles. Boat 

operators must complete the motorboat operator certification course training sponsored by the U.S. Department of Interior and be re-certified every 5 years. In 

addition, operators must be familiar with the type of 

boat and its equipment (for example, trailer, outboard 

motors, safety devices, winches, and USCG-required 

emergency equipment). All occupants on board a boat 

must wear a USCG-approved PFD at all times when 

working on watercraft less than 8 m long. Sampling 

teams also should review safety and communications 

procedures for towing and launching. Boat operators 

are responsible for the safety and security of occupants 

and equipment at all times.

The sampling team leader is responsible for 

instructing other crewmembers on the safe handling, 

operation, and storage of sampling gear. Several grab 

samplers (for example, Ponar and Ekman grabs) that 

are recommended for use in the NAWQA Program 

have spring-loaded jaws or are of sufficient mass that 

could inflict serious injury to fingers or other body 

parts.

Although electrofishing is a highly effective 

method to sample fish communities, it can result in 

serious personal injury or death if performed improperly. At least one crewmember of each electrofishing 

team must be certified by successful completion of the 

U.S. Fish and Wildlife Service training program on the 

safe operation of electrofishing gear (U.S. Geological 

Survey, 2001a). Certified individuals should act as the 

sampling team leaders and will be responsible for communication of safety protocols and individual responsibilities. At least two crewmembers must be trained and 

certified in basic first aid and cardiopulmonary resuscitation techniques (U.S. Geological Survey, 2001a). 

Only commercially manufactured electrofishing gear in 

good working condition is used. All equipment must 

receive periodic maintenance and inspection to ensure 

proper working condition and should never be altered 

for personal or operational conveniences. Dip nets must 

be made of nonconducting material (polyvinyl chloride 

tubing, or nylon); aluminum or wooden dip nets must 

never be used. Everyone in the sampling team must 

wear appropriately rated rubber gloves and waders that 

have been inspected for leaks before entering the water. 

Rubber gloves must cover the forearm for maximum 

protection. Chest waders with slip-resistant boot soles 

(for example, heavy-duty felt) must be worn in wadeable habitats; hip boots are preferable when boat electrofishing in nonwadeable habitats. Waders 

manufactured with a "breathable" fabric must not be 

used during electrofishing because of a potential for 

shock resulting from inadequate insulating properties 

of the fabric.

4. ALGAL SAMPLING PROTOCOLS

Stream algal communities are sampled in the 

NAWQA Program following nationally consistent protocols. Algae grow in streams as periphyton (attached 

to a substrate) or as phytoplankton (unattached, carried 

in current). Periphyton are further grouped into growth 

forms, either as microalgae (morphology is microscopic, appearing as pigmented accumulations or films 

attached to submerged surfaces, typically single-celled 

algae) or macroalgae (morphology is visible without 

magnification, typically filamentous). Algal communities are characterized by using richness and abundance 

data derived from quantitative and qualitative samples 
Field Protocols 15

of these communities. Regardless of which sample 

types are collected in a particular study, all samples are 

collected from instream habitats that are present in the 

reach. The instream habitats are identified according to 

a hierarchical framework of geomorphic channel units, 

channel boundaries, and channel features. The 

algal-sampling protocols described herein present 

guidelines for identifying instream habitats, methods 

for quantitatively and qualitatively sampling these habitats, and methods for processing collected samples in 

the field. These revised algal-sampling protocols are 

based on original guidance presented by Porter and others (1993).

4.1 Algal sample types collected

The algal-sampling protocols present methods 

for collecting three different types of quantitative algal 

sample (RTH, DTH, and PHY) and a qualitative algal 

sample (QMH) (table 3). These samples are formed by 

compositing discrete collections taken in the targeted 

sampling habitats (RTH and DTH) or multiple habitats 

throughout the reach (QMH). Each quantitative sample 

represents a composited sample of microalgae that is 

further split (except for DTH) into as many as three 

different subsamples, which correspond to a particular 

laboratory analysis (table 3). Two of these subsamples 

are processed by the National Water Quality 

Laboratory (NWQL) to measure chlorophyll a (CHL 

A) and ash-free dry mass (AFDM) except PHY 

samples, which are analyzed instead for CHL A and 

particulate organic carbon (POC). [Note: The NWQL 

determines POC by subtracting particulate inorganic 

carbon from total particulate carbon; this analysis 

requires a minimum of two filters.] A third subsample 

is processed for taxonomic identification (ID) by the 

Academy of Natural Sciences of Philadelphia (ANSP), 

Phycology Section to provide estimates of taxa 

richness, cell density, relative abundance, and 

biovolume of algal species. The QMH sample consists 

of both micro- and macroalgal sample components, 

each of which is only processed in a taxonomic 

laboratory to determine algal taxa that are present in 

the sample (table 3).

4.2 Identifying instream habitats for sampling

In general, the identification of instream habitats 

follows the same approach that is applied for sampling 

invertebrate communities in the reach. However, the 

terminology applied in referring to potential sampling 

habitats follows a convention among algal ecologists to 

classify these habitats according to the following 

groups: (1) epilithic, (2) epidendric, (3) epiphytic, (4) 

epipsammic, and (5) epipelic. Characteristics of each 

of these habitats are as follows:

• Epilithic—benthic habitat consisting of natural, 

coarse-grained substrates (for example, gravels, 

cobbles, or boulders) or bedrocks, or artificial hard 

substrates such as submerged concrete on which 

organisms are attached or loosely associated.

• Epidendric—benthic habitat consisting of woody 

substrates (for example, woody snags) on which 

organisms are attached or loosely associated.

• Epiphytic—benthic habitat consisting of plants on 

which organisms are attached or loosely associated.

• Epipsammic—benthic habitat consisting of 

sand-sized (> 0.064–2 mm) particles on which 

organisms are attached or loosely associated.

• Epipelic—benthic habitat consisting of silt-sized 

(< 0.064 mm) streambed sediments on which 

Quantitative algal samples Qualitative algal sample

Sample types RTH DTH PHY QMH

Sample components processed Micro Micro Micro Micro Macro

Subsamples (submitted for 

laboratory analysis)

ID

CHL A

AFDM

ID

ID

CHL A

POC

ID ID

Table 3. Algal sample types, sample components, and subsamples collected and prepared in the National 

Water-Quality Assessment Program

[RTH, richest targeted habitat; DTH, depositional targeted habitat; QMH, qualitative multihabitat; PHY, phytoplankton; Micro, 

microalgae; Macro, macroalgae; ID, Identification/enumeration; CHL A, chlorophyll a; AFDM, ash-free dry mass; POC, 

particulate organic carbon]
16 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

organisms are loosely associated. This habitat is 

commonly found in areas of low velocities, such as 

pools and side-channel areas, where silt can 

deposit.

4.3 Quantitative targeted-habitat sampling methods

The selection of the appropriate RTH and DTH 

to sample is based on information derived from the 

study objectives, reconnaissance sampling, and consultation with technical specialists. Selection of the habitat 

where the RTH microalgal sample components are collected is based on the following priority: (1) riffles in 

shallow streams with coarse-grained substrates 

(eplithic habitat); (2) woody snags in streams with 

fine-grained substrates (epidendric habitat); and (3) 

macrophyte beds (epiphytic habitat) in streams where 

riffles and woody snags are absent. In contrast, selection of the habitat where the DTH microalgal sample 

component is collected is based on the presence of 

organically rich or sandy depositional areas along 

stream margins (episammic and epipelic habitats). A 

DTH microalgal sample is only collected if a DTH is 

present in the reach.

Measurements of water depth and velocity are 

collected for discrete collections of periphyton and 

recorded on the appropriate field data sheet. Light measurements are taken at three locations in the reach to 

determine light extinction coefficients. Rinse water 

should be prepared before sampling by filtering native 

stream water. The volumes of water used to acquire or 

rinse discrete collections must be carefully monitored 

to ensure that the total sample volume after compositing does not exceed 475 mL. Doing this will allow 

enough empty volume to add formaldehyde and not 

exceed a 500-mL sample-volume limitation.

4.3.1 Sampling methods for epilithic habitats 

Epilithic habitats are sampled by using one of 

three methods depending on the types of rock substrate 

being sampled. The SG-92 (fig. 3a) is a modified 

syringe-sampling device and performs best on smooth 

cobble surfaces with moderate-to-dense coverage of 

microalgal periphyton. The "top-rock scrape" method 

(fig. 3b) is used for sampling irregular cobble surfaces 

or when cobble surfaces have sparse periphyton coverage. The gravel sampler (fig. 3c) is used when sampling microalgal periphyton attached to gravel 

substrates.

Using the SG-92 to sample epilithic habitats:

1. Assemble the SG-92 sampling device and periphyton brushes (fig 3a). This step should be completed before going into the field; also, prepare 

several SG-92 samplers and periphyton brushes.

a. Remove the end of a 30-mL syringe barrel 

opposite the syringe flanges. Sand the cut end 

of the barrel smooth.

b. Flatten one side of a rubber O-ring (inside 

diameter, 2.06 cm; outside diameter, 2.70 cm) 

using emory cloth.

c. Using cyanoacrylate adhesive, cement the flattened side of the O-ring to the flanged end of 

the syringe barrel.

d. Construct periphyton brushes by affixing 

(with epoxy adhesive) small, circular bristles 

from a stiff-bristled toothbrush to the ends of 

0.64-cm diameter plastic rods.

e. Trim the bristles to a length of about 4 mm. 

[Note: the periphyton brushes are discarded 

when the bristle lengths are worn to about 

1 mm (Porter and others, 1993)].

2. Collect five cobbles from each of five locations in 

riffles distributed throughout the reach (a total of 

25 cobbles per reach). Place cobbles in a plastic 

dishpan and transport them to an on-site processing station to collect periphyton from each cobble.

3. Place the SG-92 barrel on a smooth part of the 

cobble. Press down on the O-ring and rotate 

slightly to create a tight seal. Maintain this seal 

while the collection is made.

4. Using a pipettor, squirt about 5 mL of filtered 

stream water into the SG-92 barrel on the cobble. 

If the water leaks from the barrel, select another 

place on the cobble and try again. If the water 

does not leak, insert the periphyton brush into the 

barrel and scrub the enclosed area on the cobble 

to remove the periphyton.

5. Remove the periphyton and water mixture with 

the pipette and dispense it in a 100-mL graduated 

cylinder. [Note: dispensing into a graduated cylinder instead of a 500-mL sample bottle is recommended in case the SG-92 seal fails while 
Field Protocols 17

a. SG-92 sampler: preparing to scrape a cobble with 

microalgal periphyton brush (arrow points to O-ring seal 

on bottom of SG-92).

b. Top-rock scrape method: scraped area of a cobble 

covered with foil to determine the area sampled.

d. Inverted petri dish method: a collection being 

taken in an epipsammic habitat (arrow points to 

spatula placed beneath the petri dish to keep the 

collection intact). c. Gravel sampler: beveled edge (arrow) 

on bottom of sampler improves coring into 

gravel substrates.

collecting the sample, thereby causing the collector to start over. If the seal fails, then only the 

contents of the graduated cylinder are discarded.] Repeat this process several times until 

all of the visible periphyton is removed. Pour the 

contents of the graduated cylinder into a 500-mL 

sample bottle.

6. Repeat the sampling procedure for a single area 

on each of the cobbles selected (the composited 

sample is composed of 25 discrete collections 

taken from 25 cobbles). Ensure that the sample 

volume does not exceed 475 mL. Place the bottle 

on ice inside a cooler and keep in the dark until 

the sample is processed.

7. Measure the diameter of the area sampled by the 

SG-92 at the beginning and end of sampling. 

Record these diameters on the Quantitative 

Targeted-Habitat Periphyton Field Data Sheet to 

establish an average diameter from which the 

sampling area can be calculated.

8. Calculate the total sampling area by using the following formula:

Total sampling area (cm2)

where,

n = number of discrete SG-92 collections

π = 3.1416, and

d = average diameter of the sampled areas, in

centimeters.

2 = (n)(π )(d 2)

Figure 3. Methods used to collect microalgal periphyton samples (Photographs a, b, d by R. Frehs; c by S. Porter).
18 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

[Note: if using the inside diameter of a 30-mL 

syringe barrel, then the total surface area sampled for 25 cobbles will be about 75 cm2.]

9. Process the periphyton sample following the 

steps described in "Sample Processing Procedures" (Section 4.5).

Using the top-rock scrape method to sample epilithic 

habitats:

1. Collect five cobbles from each of five locations in 

riffles distributed throughout the reach (a total of 

25 cobbles per reach). Place cobbles in a plastic 

dishpan and transport them to an on-site processing station to collect periphyton from each cobble.

2. Identify the area on each cobble where periphyton are attached by using a red wax pencil to draw 

a line around the middle (side) of the cobble. The 

area above this line represents the sampling area 

to be scraped.

3. Using a small brush or pocket knife, scrape the 

periphyton from the sampling area on each cobble (typically the top and sides) down to the red 

line.

4. Rinse the periphyton from each cobble into the 

dishpan using a poultry baster and filtered stream 

water. Ensure that the sample volume does not 

exceed 475 mL.

5. Pour the contents of the dishpan through a funnel 

into a 500-mL sample bottle. Place the bottle on 

ice inside a cooler and keep in the dark until the 

sample is processed.

6. Wrap aluminum foil around the surface of each 

cobble, covering the area that was scraped down 

to the red line. Mold the foil tightly (fig. 3b) and 

trim the excess foil from the bottom edge of the 

scraped area.

7. Remove the formed foil from each cobble and 

flatten by making a series of radial cuts.

8. Place the foil templates in a labeled resealable 

plastic bag and determine the area of each template in the field office. The areas for all rocks 

sampled in the reach are summed and the total 

area recorded on the Quantitative 

Targeted-Habitat Periphyton Field Data Sheet 

and sample labels.

9. Process the periphyton sample following the 

steps described in "Sample Processing Procedures" (Section 4.5).
Field Protocols 19

Using the gravel sampler to sample epilithic habitats:

1. Assemble the gravel sampler from a plumbing 

"clean-out" (7.6-cm diameter) (fig. 3c). Attach 

the threaded cap; bevel the bottom edge of the 

clean-out by using a grinding wheel to improve 

the coring capability of the sampler. Obtain a 

large masonry trowel wide enough to enclose 

completely the bottom of the sampler.

2. Select 5 to 10 sampling locations throughout the 

reach.

3. Press the beveled end of the sampler into the 

gravel substrate. After the sampler is in place, 

carefully remove the gravel surrounding the outside of the sampler and insert the masonry trowel.

4. Slide the sampler onto the trowel and carefully 

lift it out of the water.

5. Quickly invert the sampler to contain the gravel 

and water in the sampler cap.

6. Pour each discrete collection into a dishpan and 

rinse the sampler before taking another discrete 

collection.

7. Repeat these steps to complete 5 to 10 discrete 

collections, which form the composited sample.

8. Extract macroalgal filaments (if present) from the 

gravel with forceps and then cut them into fine 

pieces.

9. Brush and rinse (with dishpan water) the gravel. 

Recycle rinse water to keep the sample volume 

< 475 mL.

10. Pour the sample from the dishpan through a funnel into a 500-mL sample bottle. Place the bottle 

on ice inside a cooler and keep in the dark if the 

sample is not processed immediately.

11. Calculate the total sampling area by using the formula presented for the SG-92 sampling method.

12. Process the periphyton sample following the steps 

described in "Sample Processing Procedures" 

(Section 4.5).

4.3.2 Sampling method for epidendric habitats

Collecting quantitative microalgal periphyton 

samples from epidendric habitats (or woody snags) presents a challenge because they generally have an irregular surface and are difficult to remove without loss of 

algal biomass (Porter and others, 1993). If the woody 

snag has a smooth surface, it can be sampled in a similar manner to epilithic habitats by using the SG-92. 

Otherwise, periphyton is collected from woody snags 

by using the cylinder scrape method.

Using the cylinder scrape method to sample epidendric 

habitats:

1. Select one woody snag in each of five locations 

throughout the reach.

2. Identify the part of the woody snag that will be 

sampled for periphyton. Carefully remove a 10- 

to 20-cm-long section with pruning shears or by 

sawing and place in a plastic dishpan. 

3. Scrub the entire surface of each woody snag section in the dishpan with a stiff brush. Rinse the 

brush and each section in the dishpan. Recycle 

rinse water to keep the sample volume less than 

475 mL.

4. Pour the sample from the dishpan through a funnel into a 500-mL sampling bottle.

5. Measure the length and diameter of each cleaned 

woody snag section and calculate the total sampling area by using the following formula 

(assumes a cylinder):

Total Sampling Area (cm2)

where,

n = number of discrete collections,

π = 3.1416,

di = diameter of each woody snag section, in 

centimeters, and

li = length of each woody snag section, in 

centimeters.

Alternatively, a foil template can be used (see 

top-rock scrape method) for irregularly shaped 

woody snag sections.

6. Process the periphyton sample following the 

steps described in "Sample Processing Procedures" (Section 4.5).

=

=

n

i

di li

1

(π )( )( )
20 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

4.3.3 Sampling method for epiphytic habitats

Epiphytic samples from macrophytes with small 

or finely dissected leaves (for example, Elodea or Ceratophyllum; Terrell and Perfetti, 1996) are difficult to 

quantify because the surface area of periphyton colonization cannot be reliably determined in the field. However, quantitative samples should be collected from 

these macrophytes if the epiphytic periphyton microhabitat represents the dominant RTH in the sampling 

reach.

Using the square frame method to sample epiphytic 

habitats:

1. Select five locations in the reach from which 

macrophytes can be sampled.

2. Carefully place a 50- x 50-cm square frame (for 

example, Slack sampler with area template) over 

one of the macrophyte beds. Do not disturb the 

macrophyte leaves; epiphytes are often loosely 

attached.

3. Cut the plants at their bases within the frame and 

place them in a plastic bag. Alternatively, macrophytes can be placed carefully in the Slack sampler net.

4. Rinse the macrophytes with water in the plastic 

bag; additional agitation or brushing might be 

necessary to remove epiphytic periphyton. Set the 

rinsed macrophytes aside to determine their surface area.

5. Repeat this collection procedure in four additional macrophyte beds in the reach.

6. Combine the discrete collections contained in the 

plastic bags. Ensure that the sample volume does 

not exceed 475 mL.

7. Pour the sample through a funnel into a 500-mL 

sample bottle. Place the sample on ice inside a 

cooler and keep in the dark until the sample is 

processed.

8. Determine the surface area of the sampled macrophytes. If necessary, save examples and record 

the number of each different macrophyte by 

pressing and drying for identification in the field 

office.

9. Process the periphyton samples following the 

steps described in "Sample Processing Procedures" (Section 4.5).

4.3.4 Sampling method for epipsammic/epipelic habitats

Quantitative microalgal periphyton samples are 

collected from the upper 5- to 7-mm layer of epipsammic (sand) or epipelic (silt) habitat in depositional 

areas of the reach.

Using the inverted petri-dish method to sample epipsammic or epipelic habitats:

1. Select five locations in the reach that have a depositional zone consisting of either sand or silt substrates. [Note: all five locations must be either 

sand or silt.]

2. At each location, hold the lid of a small plastic 

petri dish (about 47-mm diameter) upside down 

in the water; rub the inside of the lid to remove air 

bubbles.

3. Turn the inside of the lid toward the substrate that 

will be sampled without disturbing the sediment.

4. Carefully and slowly press (in cookie cutter fashion) the lid into the sediment.

5. Slide the lid onto a spatula (fig. 3d) to enclose the 

discrete collection. Holding the petri dish tight 

against the spatula, carefully wash extraneous 

sediment from the spatula, and then lift out of the 

water.

6. Invert the lid and remove the spatula.

7. Rinse the sediment from the lid with filtered 

stream water into a 500-mL sample bottle.

8. Repeat this collection procedure at each additional sampling location in the reach.

9. Combine the five discrete collections in a 

500-mL sample bottle (the total area sample will 

be about 85 cm2).

10. Process the sample following the steps described 

in "Sample Processing Procedures" (Section 4.5). 

[Note: subsamples for CHL A and AFDM analyses typically are not prepared from DTH samples; 

consult your study team leader or Regional Biologist if these analyses are required for this sample type.]
Field Protocols 21

4.3.5 Sampling method for phytoplankton

Quantitative samples of phytoplankton may be 

collected as part of routine water-quality sampling. The 

water sample is poured into a churn or cone splitter 

from which the PHY and water-quality samples are 

collected (Ward and Harr, 1990).

Collecting a quantitative phytoplankton sample:

1. Using a subsurface grab or depth/widthintegrating sampler (for example, a D-96 sampler), obtain a representative whole-water sample 

of sufficient volume to ensure adequate phytoplankton biomass for analysis. A 1-L sample is 

sufficient for productive, nutrient-enriched 

streams (as indicated by a noticeable color to the 

water); larger volumes up to about 5 L may be 

necessary for phytoplankton samples collected 

from clear, ground-water-fed streams or unproductive, low-nutrient rivers (as indicated by water 

transparency).

2. Withdraw subsamples for analysis by using a 

cone or churn splitter. Adequate subsample volumes range from 50 mL to more than 500 mL for 

clear, ground-water-fed streams.

3. Process the sample following the steps described 

in "Sample Processing Procedures" (Section 4.5).

4.3.6 Determining light availability

Light availability is a major factor affecting plant 

productivity in aquatic systems. The amount of sunlight that reaches the stream surface is influenced by 

local climate, canopy, and stream size. The area of the 

stream exposed to light (open canopy) and the amount 

of riparian shading are measured as part of the habitat 

characterization (CANSHADE and spherical densiometer estimates; see Fitzpatrick and others, 1998). 

Water clarity is an important factor once the light 

reaches the water surface because particles in water 

(sediment and plankton) absorb, reflect, and scatter 

wavelengths of light. The amount of light available for 

algal growth influences the structure of algal, invertebrate, and fish communities. Therefore, quantitative 

measurements of water clarity are important for evaluating the biological community structure of streams in 

relation to water quality.

Light meters and underwater sensors (for example, LI-CORTM 250 meter with LI-CORTM 192SA 

underwater quantum sensor) are used in the NAWQA 

program to obtain a direct measure of the solar energy 

available for algal growth. Measurements of the photosynthetically active radiation (PAR) (400–700 nm 

wavelength) available to the algal community are taken 

so that reach-specific factors can be distinguished from 

the effects of large-scale human and natural factors 

influencing water quality.

Water clarity can be determined using several 

methods. Descriptive terms (for example, clear, slightly 

turbid, turbid, and very turbid) are frequently used but 

are subjective and difficult to apply consistently. Secchi 

discs, although widely used in lakes and reservoirs, are 

not practical in most streams because pools are typically not deep enough to take a reading. Moreover, Secchi disc readings are considered subjective as a result 

of operator differences associated with viewing 

through the reflective surface of the water. Turbidity 

tubes are less subjective, but require careful reading 

and interpretation, because particles in the tube tend to 

settle out. Instruments (for example, HachTM Model 

2100P Turbidimeter) that measure water clarity in standard nephelometric turbidity units (NTUs), or light 

meters that measure decreases in PAR are now widely 

used for measuring water clarity.

Collecting light measurements:

1. Assemble a light meter and underwater light sensor to read and record the PAR, in units of photon 

flux density [micromoles (of photons) s-1 m-2].

a. Attach the light sensor to a 1.5-m piece of 

1.3-cm polyvinyl chloride (PVC) pipe by 

feeding the sensor cord through the pipe.

b. Secure the sensor (pointing up) to the bottom 

of the pipe with duct tape.

c. Mark the pipe at 5-cm intervals from the top 

of where the sensor is attached.

d. If necessary, modify the PVC pipe to ensure 

that it remains plumb and steady while light 

readings are taken. Helpful modifications 

include adding weight to the bottom of the 

pipe or a bubble level to the top of the pipe.

2. Locate a pool in the reach from which light readings can be taken. Ideally, the pools should be 

shaded if complete cloud cover is not available on 

the day of sampling. If pools are not present in 

the reach, locate a pool outside the reach (for 

example, a bridge scour) that can be assigned to 
22 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

the reach where community sampling was conducted. Otherwise, locate the deepest portion of 

the reach that can be safely waded. [Note: if the 

stream is not well mixed, then light readings 

should be taken at a second or third location in 

the reach.]

3. Take light readings quickly because the amount 

of available light at the surface is sensitive to 

changes in the wind, cloud cover, and disturbance 

at the water surface. [Note: it may be necessary to 

take up to three series of depth readings per location because of these factors; calculate an average of multiple readings for each depth at each 

pool location.]

4. Lower the sensor into the water and take the first 

light reading about 1 cm below the air/water 

interface.

5. Continue lowering the sensor and take a reading 

at every 5- or 10-cm interval until the sensor 

reaches the stream bottom or the meter reads 1 

percent of the first (subsurface) light reading, 

whichever comes first. The objective is to maximize the number of light readings at each pool 

location. 

6. Record the beginning time of the light readings, 

the depth (in centimeters as indicated by the 

depth interval on the PVC pipe), and the light 

reading for all measurements taken at each pool 

location on the Quantitative Targeted Habitat Periphyton Sample Field Data Sheet.

4.4 Qualitative multihabitat sampling methods

Periphyton are sampled qualitatively from multiple instream habitats for the purpose of documenting 

the presence (excluding abundance) of algal taxa 

throughout the reach. Discrete collections of microand macroalgae are taken by scraping, brushing, and 

siphoning periphyton from submerged substrates in 

five different habitats (epilithic, epidendric, epiphytic, 

epipsammic, and epipelic). The QMH sample is 

formed by combining the discrete collections for each 

of the micro- and macroalgal sample components.

4.4.1 Microalgae

Although examples of the five habitats might be 

present in the reach, these habitats will not be equally 

abundant. Therefore, to ensure that periphyton taxa 

associated with a particular habitat are adequately represented in the microalgal sample component, separate 

discrete collections are taken for each habitat. The discrete habitat collections then are processed with the 

equally weighted composite (EWC) method to create a 

single composited QMH microalgal sample component 

(see "Sample Processing Procedures," Section 4.5).

Collecting the qualitative microalgal sample component:

1. Identify the epilithic, epidendric, epiphytic, epipsammic, and epipelic habitats present in the 

reach.

2. Obtain five 125-mL bottles and label by habitat 

as (1) epilithic, (2) epidendric, (3) epiphytic, 

(4) episammic, and (5) epipelic. These bottles can 

be cleaned and reused again at other sampling 

sites.

3. Scrape, brush, or siphon periphyton from each of 

these habitats (if present) and place in the appropriate habitat bottle.

4. Process the discrete habitat collections following 

the equal EWC method described in "Sample 

Processing Procedures" (Section 4.5).

4.4.2 Macroalgae

Common examples of macroalgae (fig. 4) 

include filamentous growth forms (for example, Cladophora), plantlike algae with leaf-like structures, 

cushion and mat-forming filaments (for example, 

Vaucheria), round or flattened colonies (for example, 

Nostoc and Rivularia), short turf-forming filaments 

(for example, Audouinella), gelatinous masses, and 

short, tubular strands. Additional information and illustration necessary for identifying macroalgae can be 

found in Biggs and Kilroy (2000), Entwistle and others 

(1997), and Terrell and Perfretti (1996).

Collect macroalgae from each of the habitats 

present throughout the reach and place in an appropriately sized sample bottle (typically 125 or 250 mL). 

Preserve with buffered formaldehyde and place a label 

on the bottle. Record notes regarding the color and 

growth form (for example, mat-forming, upright filaments, leaf-like structures, and gelatinous masses) of 

algae found on the Qualitative Multihabitat Periphyton 

Sample Field Data Sheet.
Field Protocols 23

a. Cladophora, a common green alga with branched 

filaments.

b. Zygnema, like Spirogyra, is green alga with unbranched 

filaments that grows in slow-flowing backwaters 

in open environments.

c. Audouinella, short, upright, pink or blue-green filaments 

that cover rocks, like velvet, in swift flows. d. Vaucheria, a cushion to mat-forming yellow-green alga.

e. Nostoc, round, flattened blue-green algal colonies.

f. Rivularia, a colonial blue-green alga.

Figure 4. Examples of common macroalgae (photographs a, c, e, and f by J. Sonneman; b and d by B. Biggs).
24 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

4.5 Sample processing procedures

The goal of processing a composited algal 

sample in the field is to prepare subsamples for various 

laboratory analyses (table 3). Successful execution of 

the processing procedures described here to produce 

high-quality subsamples for analysis is dependent on 

measuring and tracking various volumes as the sample 

is processed. At least two subsamples are taken from 

each RTH and PHY sample for the purpose of 

determining CHL A and AFDM in the laboratory. 

However, AFDM frequently is undetectable in PHY 

samples unless the sample is collected from a highly 

eutrophic stream. Instead of measuring AFDM for 

PHY samples, a measure of carbon is obtained in the 

laboratory by analyzing the subsample for POC. POC 

is determined by subtracting particulate inorganic 

carbon from total particulate carbon. The remaining 

volume of RTH microalgal sample component (now 

the ID subsample) is preserved for taxonomic analysis. 

DTH and QMH samples are only preserved for 

taxonomic analysis. The major tasks associated with 

processing certain algal samples (fig. 5) in the field are 

as follows:

1. Accurately measure sample, subsample, preservative, and decant volumes as required during 

sample processing. [Note: this is important to 

ensure proper laboratory processing.]

2. Homogenize the sample to create a uniform suspension of algae from which CHL A and AFDM 

subsamples are taken. [Note: this step is not necessary for PHY samples.]

3. Filter the CHL A and AFDM or POC subsamples 

onto glass fiber filters.

4. Preserve the ID subsample with buffered formaldehyde for taxonomic analysis.

5. Prepare processed samples for shipping to the 

laboratory.

6. Record site and sample information for each sample collected on a field data sheet.

4.5.1 Quantitative microalgae

Each RTH sample is processed in the field to prepare subsamples that will be analyzed for CHL A and 

AFDM, and taxonomic identification. DTH samples 

are processed only for taxonomic identification. Phytoplankton samples are processed to prepare subsamples that will be analyzed for CHL A, POC, and 

taxonomic identification. After determining the original sample volume, the subsamples are prepared by following the procedures described below for 

homogenizing, filtering, preserving, and packaging.

Processing the quantitative microalgal sample component:

[Note: if processing a DTH sample, skip to step 

number 10.]

1. Measure the sample volume to the nearest 

milliliter.

2. Calibrate the pipette. [Note: the calibration is 

important, especially if the tip has been trimmed 

to enlarge the opening for extracting dense periphyton material.]

3. Assemble the filtration apparatus by attaching the 

filter base with rubber stopper to the filtering 

flask. Join the flask and a hand-operated vacuum 

pump (with gage) using a section of tubing 

(fig. 6a). [Note: if POC filters are prepared as 

part of dissolved organic carbon sample processing (not discussed in this report), then a Teflon 

filter apparatus must be used.]

4. Place a 47-mm glass fiber filter (for example, 

WhatmanTM GF/F) on the filter base and wet with 

deionized water. Use a 25-mm baked glass fiber 

filter for POC. [Note: wetting the filter will help 

keep it in place in windy weather. Attach the filter 

funnel to the base.]

5. Homogenize the microalgal sample component. 

[Note: a battery-powered stirrer (fig. 6b) works 

well for this purpose.] Cut algal filaments, if 

present, into pieces about 2 mm in length. 

Homogenization is not necessary for PHY samples because subsamples are taken directly from a 

cone or churn splitter.

6. Shake the sample component vigorously for 

about 30 seconds to ensure that it is well mixed 

before extracting subsamples.

7. Extract two 5-mL aliquots of homogenized 

microalgal sample using the pipette and dispense 

onto the wetted glass-fiber filter (fig. 6c). [Note: 

for the PHY CHL A subsample it will be necessary to filter a 50- to 500-mL subsample (depend-
Field Protocols 25

Homogenize sample

(not necessary for PHY)

Remove 10 mL aliquots each

for CHL A, AFDM, or POC

subsample analysis

Filter each 10 mL aliquot onto

separate filters

Prepare filters for shipping,

place on dry ice

CHL A subsample

filter

AFDM

(POC if PHY)

subsample filter(s)

Label samples and prepare

bottles for shipping

Log samples into Bio-TDB and submit to

ANSP for analysis

ID

subsample bottle

Log samples into Bio-TDB and submit to

NWQL for analysis within 3 weeks

What is the

sample type?

QMH

Remaining

sample

(ID SSV)

Measure SV

RTH, DTH, PHY

Measure SV

EWC Method

Microalgae Macroalgae

Preserve with formaldehyde

(measure PV)

Is volume

> 500 mL?

YES NO

Decant to

reduce SSV to

< 500 mL

Measure ADV

Measure SV RTH, PHY DTH

Figure 5. Steps followed in the field to process algal samples in the National Water-Quality Assessment Program. Steps in dashed box 

are typically performed in the field office. <, less than; >, greater than; mL, milliliter; RTH, richest targeted habitat; DTH, depositional 

targeted habitat; PHY, phytoplankton; QMH, qualitative multihabitat; SV, sample volume; SSV, subsample volume; ADV, after decant 

volume; PV, preservative volume; EWC, equal weight composite, CHL A, chlorophyll a; POC, particulate organic carbon; AFDM, 

ash-free dry-mass; ID, identification; Bio-TDB, Biological Transactional Data Base; NWQL, National Water Quality Laboratory; ANSP, 

Academy of Natural Sciences of Philadelphia.
26 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

a. Apparatus used to filter microalgal sample 

components: hand-operated pump, Erlenmeyer 

flask, tubing, filter funnel and base. 

b. Homogenizing the microalgal sample component 

with a battery-powered stirrer.

c. CHL A or AFDM subsample filter 

removed from filter base with

forceps.

d. Subsample filter folded in quarters and wrapped

in aluminum foil.

e. Label placed on petri dish containing 

subsample filter and then frozen in a 

plastic bag. 

f. Performing the EWC method to prepare 

the QMH microalgal sample component.

Figure 6. Major steps in processing microalgal sample components in the field (photographs by R. Frehs). 

CHL A, chlorophyll a; AFDM, ash-free dry mass; EWC, equal weight composite; QMH, qualitative multihabitat.
Field Protocols 27

ing on PHY density as indicated by the color of 

the filtered sample) taken from a cone or churn 

splitter. For the PHY POC subsample, at least 

two 25-mm filters must be prepared by filtering a 

30- to 100-mL subsample on each filter. The 

reduced subsample volume is necessary because 

of the smaller filter size. The filtered subsample 

should be a tan to light brown color on the filter.]

8. Filter the aliquots by using 10 psi (69 kPa).

a. Examine the filter. An adequate amount of 

microalgal biomass for analysis is indicated 

by the green or brown color of material 

retained on the filter (fig. 6c). Extract additional 5.0-mL aliquots and filter until the 

desired level of biomass is obtained.

b. Determine the number of 5.0-mL aliquots filtered and record the subsample volume on the 

field data sheet (For example, 2 aliquots x 

5.0 mL/aliquot = 10 mL subsample volume).

c. Rinse the funnel sides with deionized water; 

allow the water to be vacuumed completely 

before releasing the vacuum from the filtering 

apparatus.

d. Remove the filter from the funnel base with 

forceps.

e. Repeat the filtering steps for each subsample 

collected. [Note: additional subsamples might 

be collected and filtered as QC replicates for 

CHL A and AFDM analysis if required by the 

study.]

9. Prepare the filtered CHL A and AFDM or POC 

subsamples for shipping to the NWQL.

a. Fold each filter into quarters with filtered biomass inside. Wrap each filter in a small piece 

of aluminum foil and place in separate 47-mm 

plastic petri dishes (fig. 6d, e).

b. Seal the edge of the petri dish with plastic 

tape. Label the dish with the following 

required information: site, collection date, 

total sample area, sample volume, subsample 

volume, and sample identification code.

c. Place the labeled petri dishes in resealable 

plastic bags and pack in a cooler containing 

dry ice. About 4.5 kg (10 pounds) of dry ice is 

needed for subsamples packed in a small 

cooler (< 2 gal). Insulate the cooler with 

newspaper to minimize sublimation of the dry 

ice.

d. Complete an Analytical Services Request 

(ASR) form for each sample that indicates 

which laboratory analyses are to be performed. Use NWQL Schedule 1632 [analysis 

for periphyton using U.S. Environmental Protection Agency (USEPA) Fluorometric 

Method No. 445 for CHL A and AFDM determination] for RTH samples. Use NWQL 

Schedule 1637 (analysis of CHL A in phytoplankton) and NWQL Schedule 2631 (analysis of carbon) for PHY samples.

e. Contact NWQL personnel to make them 

aware of plans to ship (via overnight shipping 

service) coolers containing dry ice and frozen 

CHL A and AFDM or POC subsample filters. 

Coolers should be shipped within a day or two 

after the subsamples are prepared because of a 

25-day holding-time limit on the subsamples. 

Subsamples can be temporarily stored in 

freezers (at -20°C) at the field office over 

weekends.

10. Measure the volume of the remaining microalgal 

sample component (this represents the subsample 

volume of the ID subsample).

11. Preserve the ID subsample with a sufficient volume of buffered formaldehyde according to the 

following table to obtain a final concentration of 

3 to 5 percent buffered formalin. Record the preservative volume on the field data sheet.

12. Place a completed sample label (fig. 2) on the 

sample bottle.

13. If the volume of the preserved ID subsample 

(subsample volume + preservative volume) is 

> 500 mL, then the ID subsample must be 

decanted until volume is < 500 mL. [Note: this 

Volume (mL)

Sample 25 50 100 125 200 250 300 350 400 450 475

Buffered

formaldehyde 1 2 4 6 8 11 14 16 18 22 23
28 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

step is typically performed in the field office after 

sampling is completed.]

a. Record the "before decant volume" on the 

field data sheet. [Note: BDV = subsample volume + preservative volume.]

b. Allow all of the algal biomass to settle to the 

bottom of the sample bottle. Four to seven 

days might be necessary for settling.

c. Attach one end of tubing to a Pasteur pipette 

and the other end to the filtering flask, which 

is attached to the vacuum pump.

d. Create a vacuum and insert the pipette tip into 

the sample bottle. Carefully siphon the excess 

liquid into the flask, making sure not to disturb the biomass settled on the bottom. Continue siphoning until the sample volume is 

< 500 mL.

e. Measure the "after decant volume" of the ID 

subsample and record on the field data sheet.

4.5.2 Qualitative microalgae and macroalgae

The QMH micro- and macroalgal sample components are preserved and submitted to the laboratory 

only for taxonomic analysis. The microalgal sample 

component is prepared by using the EWC method to 

combine equal aliquots of the discrete collections taken 

from epilithic, epidendric, epiphytic, epipsammic, and 

epipelic habitats. The resulting composited microalgal 

sample component would contain about 20 percent of 

the periphyton from each habitat if all five habitats 

were sampled.

Using the EWC method to process the QMH microalgal sample component:

1. Place the five habitat bottles on a flat surface and 

homogenize each with a battery-powered tissue 

homogenizer (fig. 6f). Allow the contents in each 

bottle to settle for several minutes.

2. Order the habitat bottles from least to greatest 

volume of biomass. 

3. Beginning with the habitat bottle having the least 

volume (typically the epiphytic habitat bottle), 

remove 5-mL aliquots from the bottom of each 

habitat bottle and dispense into a 125- or 250-mL 

sample bottle.

4. Continue removing series of 5-mL aliquots until 

the habitat bottle with the least volume is < 5 mL.

5. After discarding the remaining discrete collections, rinse the labeled habitat bottles with filtered stream water and reuse later. Measure the 

volume of the composited microalgal sample 

component and record it on the Qualitative Multihabitat Periphyton Sample Field Data Sheet 

(Algae Tip 1).

6. Preserve the sample with buffered formaldehyde 

and place a completed sample label (fig. 2) on the 

sample bottle.

4.6 Quality-control recommendations

Additional (10–20 percent) subsample splits can 

be prepared for CHL A, AFDM or POC, and ID subsamples to evaluate precision of subsample preparation 

and laboratory analysis. Replicated sampling within the 

reach can be used to evaluate the variance in algal community structure. These QC samples are collected or 

prepared depending on the objectives of the study.

4.7 Field data sheets

Three different field data sheets are used to 

record information about the sampling site, the algal 

samples collected, and supporting sample data (for 

example, light measurements). The field data sheets are 

presented in the following order at the end of this section:

• Quantitative Targeted-Habitat Periphyton Sample 

Field Data Sheet (QTHP; 2 pages)

• Quantitative Phytoplankton Sample Field Data 

Sheet (QPS; 2 pages)

• Qualitative Multihabitat Periphyton Sample Field 

Data Sheet (QMH; 2 pages)

Algae Tip 1—Determining sample volumes: 

measurement by proxy

Measurement by proxy is a good technique for 

indirectly determining the volume of a sample. 

Applying this technique minimizes handling of the 

sample and reduces the chance for sample spillage. 

Obtain an empty bottle identical to the one containing the algal sample. Fill the bottle with water until 

the volume level is equal to that in the sample bottle. Pour the water into a graduated cylinder and 

determine volume in milliliters.
Field Protocols 29

Information that is recorded on each field data 

sheet is organized into sections (table 4) that correspond to data-entry screens in Bio-TDB. Except for 

these section labels (for example, "Sample Header"), 

data fields or subsections that are shown in bold face 

are required for entry into Bio-TDB. Consult the 

Bio-TDB User Manual (Ruhl and Scudder, accessed 

April 26, 2002) for additional information on entering 

field data into the data base.

Before leaving the sampling site, the team leader 

should review the field data sheets for completeness 

and accuracy by comparing information on sample bottle/filter labels and field data sheets, especially in the 

"Sampling Information" section. All missing or incorrect information should be completed or corrected. 

Field data sheets (blank and completed) should be 

printed on waterproof paper and stored in resealable 

bags for additional protection from water.

Section This section is used to 

document

Completed on field data 

sheet?

QTHP QPS QMH

Sample header Where and when a sample 

was collected Yes Yes Yes

Related sampling activitiesOther sampling activities Yes Yes Yes

Physical site 

conditions

Physical and chemical conditions at the time of sampling Yes Yes Yes

Sampling informationHow a sample is collected 

and processed in the field Yes Yes Yes

Supporting 

information

Ancillary sampling information taken in conjunction 

with discrete collections

Yes Yes No

Table 4. Explanation of field data-sheet sections used to record 

information about algal sampling activities

[QTHP, quantitative targeted-habitat periphyton sample; QPS, quantitative 

phytoplankton sample; QMH, qualitative multihabitat sample]
30 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

Quantitative Targeted-Habitat Periphyton Sample

Field Data Sheet

SAMPLE HEADER

Station: Date (MM/DD/YYYY):

/ /

Waterbody:

Station description:

Station ID: Reach: Start time: h

Collector (leader): End time: h

Sample and photographic notes:

RELATED SAMPLING ACTIVITIES 

(circle all that apply)

Water chemistry Discharge Habitat survey Tissues

Invertebrate survey Fish survey Bed sediment Other (specify):

PHYSICAL SITE CONDITIONS

Clouds: % Wind (circle): Calm Light Moderate Gusty

Precipitation (circle): None Rain Sleet Snow Precipitation intensity (circle): N/A Light Moderate Heavy

Other weather:

Beginning water measurements Ending water measurements

Time: Time:

Water temperature

(°C):

Water temperature

(°C):

pH: pH:

Dissolved oxygen

(mg/L):

Dissolved oxygen

(mg/L):

Specific conductance

[(μS/cm) @ 25 °C]:

Specific conductance

[(μS/cm) @ 25 °C]:

Turbidity

(NTUs)

Turbidity

(NTUs)

Discharge (ft3/s): Stream stage: ft @ time h

Riparian shading (circle): Shaded Partial Full sun

Water clarity (circle): Clear Slightly turbid Turbid Very turbid

Water color (circle): Black Brown Clear Dark green Light green Yellow

Physical site condition comments:

Page # Cumulative Page # .

Date field data reviewed: / / by (initials)

Date entered into Bio-TDB: / / by (initials)

Bio-TDB Sample ID
Field Protocols 31

SAMPLING INFORMATION

Primary sample or Repeated sampling replicate? (circle one) Sample Number:

Sample type (circle one): RTH DTH Sample component: Microalgae

Periphyton habitat sampled (circle): Epilithic Epidendric Epiphytic Epipsammic Epipelic 

Sampling method or device (circle): SG-92 Top-rock scrape Cylinder scrape Gravel sampler Petri dish Other:

Sampling area—Beginning diameter: cm; Ending diameter: cm; Average diameter: cm

Number of discrete collections constituting composite sample: ; Total area of periphyton sample: cm2

Sample volume: mL

Subsample bottles/filters (complete all)

SSV=subsample volume; PV=preservative volume; BDV=before decant volume; ADV=after decant volume

Preservative: buffered formaldehyde; NA (not applicable)

Sample 

analysis

SSV

(mL)

PV

(mL)

Split?

(Y or N)

Decant?

(Y or N)

BDV

(mL)

ADV

(mL)

SMCOD 

component 

code

Bottle/

filter 

sequence

Field SMCOD

ID B

ID S

CHL A NA B

CHL A NA S

CHL A NA S

AFDM NA B

AFDM NA S

AFDM NA S

Subsample comments:

Algae QTHP Sample Page # Cumulative Page # .

SUPPORTING INFORMATION

Sample 

location 

number

Water depth 

(m)

Secchi depth 

(m)

Velocity

(ft/s)

Riparian 

shading 

S=shaded

P=partial 

F=full sun

Type and color of macroalgae, if present

1

2

3

4

5

LIGHT MEASUREMENTS (complete all)

WOODY SNAG MEASUREMENTS Type of light meter used (circle): LI-250 Other:

Type of light sensor used (circle): LI-192SA Other:

Light

Reading

No.

Depth (cm) of 

reading below 

surface of 

water 

Light intensity reading Sample

location

number

Woody

snag number

Snag length

(cm)

Snag diameter

Location 1 Location 2 Location 3 (cm)

Time: Time: Time:

1 1 1

2 2

3 3

4 4

5 5

6 6

7 7

8 8

9 9

10 10

11
32 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

Page # Cumulative Page # .

Date field data reviewed: / / by (initials)

Date entered into Bio-TDB: / / by (initials)

Bio-TDB Sample ID

Quantitative Phytoplankton Sample

Field Data Sheet

SAMPLE HEADER

Station: Date (MM/DD/YYYY):

/ /

Waterbody:

Station description:

Station ID: Reach: Start time: h

Collector (leader): End time: h

Sample and photographic notes:

RELATED SAMPLING ACTIVITIES 

(circle all that apply)

Water chemistry Discharge Habitat survey Tissues

Invertebrate survey Fish survey Bed sediment Other (specify):

PHYSICAL SITE CONDITIONS

Clouds: % Wind (circle): Calm Light Moderate Gusty

Precipitation (circle): None Rain Sleet Snow Precipitation intensity (circle): N/A Light Moderate Heavy

Other weather:

Beginning water measurements Ending water measurements

Time: Time:

Water temperature

(°C):

Water temperature

(°C):

pH: pH:

Dissolved oxygen

(mg/L):

Dissolved oxygen

(mg/L):

Specific conductance

[(μS/cm) @ 25 °C]:

Specific conductance

[(μS/cm) @ 25 °C]:

Turbidity

(NTUs)

Turbidity

(NTUs)

Discharge (ft3/s): Stream stage: ft @ time h

Riparian shading (circle): Shaded Partial Full sun

Water clarity (circle): Clear Slightly turbid Turbid Very turbid

Water color (circle): Black Brown Clear Dark green Light green Yellow

Physical site condition comments:
Field Protocols 33

SAMPLING INFORMATION

Primary sample or Repeated sampling replicate? (circle one) Sample Number:

Sample type (circle one): PHY Sample component: Microalgae

Sampling method (circle): EDI/EWI Subsurface grab Other: ; if grab or other, collection depth is: cm

Sample volume: mL

Subsample bottles/filters (complete all)

SSV=subsample volume; PV=preservative volume; BDV=before decant volume; ADV=after decant volume

Preservative: buffered formaldehyde; NA (not applicable)

Sample 

analysis

SSV

(mL)

PV

(mL)

Split?

(Y or N)

Decant?

(Y or N)

BDV

(mL)

ADV

(mL)

SMCOD 

component 

code

Bottle/

filter 

sequence

Field SMCOD

ID B

ID S

CHL A NA B

CHL A NA S

CHL A NA S

POC NA B

POC NA S

POC NA S

Subsample comments:

Algae QPS Sample Page # Cumulative Page # .

SUPPORTING INFORMATION

Location 

number

Water depth 

(m)

Velocity

(ft/s)

Secchi depth 

(m)

1

2

3

4

5

6

7

8

9

10
34 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

Page # Cumulative Page # .

Date field data reviewed: / / by (initials)

Date entered into Bio-TDB: / / by (initials)

Bio-TDB Sample ID

Qualitative Multihabitat Periphyton Sample

Field Data Sheet

SAMPLE HEADER

Station: Date (MM/DD/YYYY):

/ /

Waterbody:

Station description:

Station ID: Reach: Start time: h

Collector (leader): End time: h

Sample and photographic notes:

RELATED SAMPLING ACTIVITIES 

(circle all that apply)

Water chemistry Discharge Habitat survey Tissues

Invertebrate survey Fish survey Bed sediment Other (specify):

PHYSICAL SITE CONDITIONS

Clouds: % Wind (circle): Calm Light Moderate Gusty

Precipitation (circle): None Rain Sleet Snow Precipitation Intensity (circle): N/A Light Moderate Heavy

Other weather:

Beginning water measurements Ending water measurements

Time: Time:

Water temperature

(°C):

Water temperature

(°C):

pH: pH:

Dissolved oxygen

(mg/L):

Dissolved oxygen

(mg/L):

Specific conductance

[(μS/cm) @ 25 °C]:

Specific conductance

[(μS/cm) @ 25 °C]:

Turbidity

(NTUs)

Turbidity

(NTUs)

Discharge (ft3/s): Stream stage: ft @ time h

Riparian shading (circle): Shaded Partial Full sun

Water clarity (circle): Clear Slightly turbid Turbid Very turbid

Water color (circle): Black Brown Clear Dark green Light green Yellow

Physical site condition comments:
Field Protocols 35

SAMPLING INFORMATION

Primary sample or Repeated sampling replicate? (circle one) Sample Number:

Sample type (circle one): QMH Sample component: Microalgae (code A) and Microalgae (code B)

Recognizable periphyton habitats

Epilithic: % Epidendric %

Epiphytic: % Epipsammic %

Epipelic: % Other %

Periphyton abundance (circle): Dense Moderate Sparse None Not rated

Recognizable algal taxa (circle): None Filamentous algae Cladophora Spirogyra Nostoc Others (record each):

Sample volume: mL

Subsample bottles/filters (complete all)

SSV=subsample volume; PV=preservative volume

Preservative: buffered formaldehyde

Sample 

analysis

SSV

(mL)

PV

(mL) SMCOD component 

code

Bottle

sequence

Field SMCOD

ID A

ID B

Subsample comments:

Algae QMH Sample Page # Cumulative Page # .
36 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

5. INVERTEBRATE SAMPLING PROTOCOLS

Stream invertebrate communities are sampled in 

the NAWQA Program following nationally consistent 

protocols. Invertebrate communities are characterized 

by using richness and abundance data derived from 

semi-quantitative and qualitative samples of these communities. These samples are collected from instream 

habitats that are present in the reach. The instream habitats are identified according to a hierarchical framework of geomorphic channel units, channel boundaries, 

and channel features. The invertebrate sampling protocols described herein present guidelines for identifying 

instream habitats, methods for semi-quantitatively and 

qualitatively sampling these habitats, and methods for 

processing collected samples in the field. These revised 

invertebrate sampling protocols are based on original 

guidance presented by Cuffney and others (1993).

5.1 Invertebrate sample types collected

Two types of invertebrate samples are collected 

in the NAWQA Program. A semi-quantitative sample is 

collected to provide a measure of relative abundance of 

the invertebrate taxa living in the RTH in the reach. The 

RTH theoretically supports the faunistically richest 

invertebrate community and is typically represented by 

a coarse-grained riffle or a woody snag. The 

semi-quantitative RTH sample consists of a series of 

discrete collections (for example, five Slack samples or 

10 woody snag sections) that are processed and combined into a single composited sample. A QMH sample 

is collected to document the invertebrate taxa that are 

present throughout the sampling reach. A discrete 

QMH collection is taken from each of the different 

instream habitats that are present in the reach. These 

discrete collections are then processed and combined 

into a single composited sample. Both RTH and QMH 

samples are preserved and submitted to the NWQL for 

taxonomic analysis.

5.2 Identifying instream habitats for sampling

Before invertebrate sampling begins, crewmembers must identify and document the types of instream 

habitats present in the reach. This information is used 

to determine where RTH and QMH sampling is conducted in the reach. Instream habitat types are broadly 

defined on the basis of a hierarchical framework consisting of three tiers (table 1) that include (1) major 

geomorphic channel units, (2) major channel boundaries, and (3) major channel features (Cuffney and others, 1993). This instream hierarchy is applied to 

identify instream habitats in wadeable and nonwadeable streams. The following explanation of each tier in 

this hierarchy was taken or modified from Cuffney and 

others (1993):

• Tier 1 instream habitat hierarchy defines major 

geomorphic channel units (table 1) present in the 

reach, such as riffles, runs, and pools.

• Tier 2 relates the influence of major channel 

boundaries (table 1) to the distribution of invertebrates (particularly in large rivers) by subdividing 

each of the major geomorphic units into main 

channel, channel margin, and island margin areas 

(Thorp, 1992). Margins, loosely defined as 

instream areas associated with the edges of main or 

secondary channels and islands, are typically depositional, subhorizontal fluvial surfaces, with 

reduced current velocity as compared with the 

adjacent channel area. Margins are influenced 

heavily by the streambanks and typically contain 

elements directly derived from the streambank, 

such as root wads, woody snags, and terrestrial 

vegetation that trails into the water. Channels tend 

to be less influenced by the channel banks and represent the main or secondary flow path of the river. 

The extent of the margin will be influenced by river 

stage, the size of the river, and channel characteristics. For example, margins might be (1) a substantial proportion of the width of small streams but 

only a very small proportion of the width of large 

rivers, (2) greatly reduced on the outside of stream 

meanders but extensive on the inside of stream 

meanders, and (3) abundant at low flows but inaccessible at high flows. It is important to differentiate margins because marginal areas might contain 

richer communities of organisms than channel 

areas as a result of the increased complexity of the 

stream margin habitat. This circumstance is particularly true of large, deep rivers.

• Tier 3 of the instream habitat hierarchy focuses on 

major channel features (table 1) that are important 

in the distribution of invertebrates and that can be 

sampled discretely. The six types of major channel 

features that can be sampled for invertebrates 

include: natural bed, manufactured bed, slough, 

macrophyte bed, woody snag, and bar. Natural bed 

refers to areas where natural bed materials predominate and where macrophytes are not a dominant 

feature. Manufactured bed refers to artificial sub-
Field Protocols 37

strates, such as revetments, levees, pilings, junk 

cars, tires, shopping carts, riprap, dams, weirs, and 

bridge piers. Bars are shallow, gently sloping sand 

or gravel ridges primarily associated with channel 

edges or major changes in water velocity. Bars can 

resemble islands when they are exposed at low 

flows and vegetated. However, islands typically 

have woody vegetation and are at an elevation 

equal to or above that of the surrounding floodplain. Sloughs are remnants of abandoned river 

channels that are connected with the main channel 

even at normal low flows. Sloughs that are isolated 

at low flows tend to diverge biologically and chemically from conditions in the river and are not considered here as an instream habitat. Woody snags 

refer to trees, branches, roots, or other woody 

debris of terrestrial origin that extend into the water 

column either from the streambank or streambed. 

Macrophyte beds are areas where emergent or submergent aquatic plants dominate and invertebrate 

communities are expected to contain organisms 

dependent upon such plants.

Collectively, this three-tiered hierarchy describes 

54 possible habitat types (table 5). However, sloughs 

are restricted to channel and island margins, so only 51 

habitat types are available for qualitative and 

semi-quantitative sampling. Following the guidelines 

for establishing the size and location of the sampling 

reach (see Fitzpatrick and others, 1998) should lead to 

the inclusion of the majority of instream habitat types 

typical of the stream at a given location.

5.3 Semi-quantitative targeted-habitat methods for 

sampling wadeable and nonwadeable streams

The type of RTH sampled may vary with the 

geographic location of the stream and the longitudinal 

position of the sampling reach along that stream. Based 

on the instream habitat matrix presented in table 5, habitat types are assigned priority (fig. 7) to help crewmembers identify the most appropriate RTH that will 

be semi-quantitatively sampled. Elements in each of 

the three hierarchical tiers that collectively define a 

particular habitat type are ranked from highest priority 

(level 1) to lowest priority (level 3 or 5 depending on 

the tier).

In general, invertebrate diversity and abundance 

increase with increasing particle size and then decline 

with particle size greater than the size of cobbles (Minshall, 1984; Allan, 1995). Increasing substrate particle 

size results in more stable substrates and interstitial 

spaces that are available for colonization. Substrate stability and the greater exchange of gasses and water are 

factors explaining why coarse-grained substrates are 

the faunistically richest areas in a stream. In most circumstances, a "riffle, main-channel, natural-bed" habitat type is selected as the RTH for semi-quantitative 

sampling in wadeable streams ("natural bed" in this 

case refers to coarse-grained substrates). When this 

habitat type is not predominant in the reach (for example, a sandy-bottomed Coastal Plain stream) or cannot 

be sampled effectively (for example, nonwadeable 

streams), then woody snags are often selected as the 

RTH.

Major 

geomorphic 

channel units

Major

channel

boundaries

Major channel features

Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Riffle

Main channel

Channel margin

Island margin

Run

Main channel

Channel margin

Island margin

Pool

Main channel

Channel margin

Island margin

Table 5. Generalized matrix of the 51 instream habitats (as defined by major geomorphic channel units, channel 

boundaries, and channel features) that can be present in a sampling reach from which invertebrate samples are 

collected (from Cuffney and others, 1993). Open cells indicate habitats that are possible; shaded cells indicate 

habitats that are not possible
38 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

5.3.1 Method for sampling coarse-grained natural-bed 

substrates

Disturbance-removal sampling is the most 

widely used technique for dislodging organisms from 

coarse-grained substrates. This technique involves 

defining a specific sampling area and then disturbing 

the substrate within that defined area to dislodge invertebrates that are washed into a net downstream. The 

Slack sampler (fig. 8a) is the standard disturbance 

removal sampler used in the NAWQA Program for 

wadeable sites where the water depth does not exceed 

the height of the collection net frame. The Slack sampler consists of a wooden handle that is attached to a 

rectangular net frame (50 by 33 cm) fitted with a 

tapered, 500-μm NitexTM collection net. The cod-end of 

the net, where sample material is trapped, can be fitted 

with a detachable collection receptacle (for example, 

Dolphin bucket fitted with a 500-μm screen). After taking a discrete collection, the receptacle is removed to 

retrieve the sample. Alternatively, the cod-end can be 

folded over two or three times and then secured with a 

large binder clip. Invertebrates are collected from a 

0.25-m2 area immediately upstream of the Slack sampler. The sampling area is determined by using either a 

50-cm-long guide rod or an area template (Cuffney and 

others, 1993). The Slack sampler can be modified in 

several ways to improve operation of the sampler and 

increase the accuracy of the area sampled (fig 8a; 

Invertebrate Tip 1).

Procedure for sampling riffles with the Slack sampler:

1. Select riffles throughout the reach from which 

five discrete collections can be taken. The sampling location for each discrete collection should 

be consistent with respect to substrate type, current velocity, depth, and debris accumulation. 

Avoid locations in riffles that are adjacent to 

other instream habitats, are directly below 

obstructions, or are normally outside of the main 

channel.

Invertebrate Tip 1—Operational modifications 

for the Slack sampler

(a) Bipod stand—Turn the Slack sampler into a 

freestanding device by attaching a bipod stand, 

eliminating the need to have a crewmember 

hold the sampler. The bipod stand is used to 

keep the Slack sampler upright while a collection is taken by one or two crewmembers (see 

Appendix).

(b) Sampling-area template—Use an area template to accurately delineate the sampling area 

upstream of the Slack sampler (fig. 8a). Area 

templates can be constructed out of PVC or a 

lightweight metal (see Appendix). Using an 

area template ensures that the sampling area is 

not exceeded during disturbance removal sampling. The template also ensures a more accurate quantitative sample and provides a defined 

area for better determination of the larger 

rocks that are retained for processing.

(c) Neoprene flange—Attach a strip of neoprene 

along the bottom edge of the Slack sampler 

frame. The flange will help to achieve a tight 

seal against the substrate.

Major geomorphic

channel unit

Major channel

boundary

Major channel

feature

Priority

level Descriptor Prioritylevel Descriptor Prioritylevel Descriptor

1 Riffle 1 Main channel 1 Natural bed

2 Run 2 Channel margin 2 Woody snag

3 Pool 3 Island margin 2 Macrophyte bed

3 Bar

4 Manufactured bed

5 Slough

Figure 7. Priority levels for determining the habitat type that is selected as the richest targeted 

habitat for semi-quantitatively sampling invertebrates (from Cuffney and others, 1993).
Field Protocols 39

2. Acquire each discrete collection by using a Slack 

sampler. Each collection is taken from a 0.25 m2

area immediately upstream of the Slack sampler 

(total area sampled = 1.25 m2).

3. Position the Slack sampler perpendicular to the 

direction of flow and press tightly against the 

stream bottom (see fig. 10a, section 5.5). [Note: 

make sure the water depth does not exceed the 

height of the sampler frame.]

4. Measure water depth, substrate type (dominant 

and codominant), percent substrate embeddedness, and velocity for each discrete collection. 

Record these data on the "Semi-Quantitative Targeted Habitat Invertebrate Sample Field Data 

Sheet."

5. Remove large debris (for example, large cobble 

and woody debris) from the sampling area if > 50 

percent of their surface is within the sampling 

area.

6. Inspect the debris for attached organisms. 

Remove organisms by hand picking, rinsing, or 

gently scrubbing the particle surface (see fig. 10b, 

section 5.5) with stream water in front of the 

Slack sampler opening. Alternatively, the debris 

can be placed in a bucket and inspected at the 

time the sample is processed on the shore.

7. Disturb the sampling area by digging to a depth 

of about 10 cm (or as deeply as practical) with a 

hand rake (see fig. 10c, section 5.5). Avoid collecting excessive amounts of sand and pebbles in 

the collection net by raking in an upstream direction. This technique dislodges organisms and 

allows most of the sand and pebbles to settle out 

in the sampling area. Avoid using a kicking technique to disturb the sampling area because it 

results in more damage to organisms, forces more 

debris into the collection net, and often causes 

areas outside of the sampling area to be disturbed 

(especially when an area template is not used).

8. Retrieve the sampler by lifting it out of the water 

with a slight forward motion to ensure that none 

of the discrete collection is lost.

a. Slack sampler with area 

template and bipod stand.

b. Ponar grabs

c. Ekman grabs

d. D-frame kick net

Figure 8. Samplers used to collect invertebrate samples in the National Water-Quality Assessment Program (Photograph a. by D. 

Sun; b–d by Wildlife Supply Company).
40 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

9. Rinse the collected material into the cod-end of 

the net by carefully splashing water on the outside of the net.

10. Transfer each discrete collection into a 19-L 

(5-gal) white plastic bucket (see fig. 10d, section 

5.5). Inspect the inside of the collection net for 

organisms; place any invertebrates found in the 

discrete collection.

11. Process each discrete collection following the 

steps presented in "Sample Processing Procedures" (Section 5.5).

5.3.2 Method for sampling woody snags

Many wadeable streams throughout the Midwest 

and coastal regions of the United States do not have riffles with coarse-grained substrates present that can be 

semi-quantitatively sampled as the RTH. Under these 

circumstances, woody snags are often selected as the 

RTH that is sampled. Woody snags are submerged sections of wood (branch or log) having a minimum diameter of 1 cm and are colonized by aquatic organisms; 

other organic (for example, leaf packs) and mineral 

debris also might be associated with woody snags.

Procedure for sampling woody snags:

1. Identify at least five (if present) woody snag locations present throughout the reach. Woody snags 

that are suitable for sampling should have been 

submerged for an extended period during which 

there is clear evidence of invertebrate and periphyton colonization.

2. At each woody snag location, select at least two 

snags that will be sampled. [Note: depending on 

the size of the woody snags, additional snags 

might need to be sampled to obtain enough sample material for laboratory processing.]

3. Place a collection net (for example, a Slack sampler) downstream of the woody snag to capture 

and minimize loss of mobile or loosely attached 

invertebrates.

4. Remove the woody snag by using a saw or lopping shears. If the woody snag is too large to cut 

and remove, it can be sampled in its place by 

brushing its surface with a small hand brush.

5. Determine the substrate type where the woody 

snag was positioned. Measure the depth from the 

surface of the water to the woody snag and stream 

bottom, and current velocity. Record these data 

on the "Semi-Quantitative Woody Snag Invertebrate Sample Field Data Sheet."

6. Remove attached organisms by handpicking, 

brushing, and rinsing the surface of each branch 

into a 19-L (5-gal) plastic bucket (see Invertebrate Tip 2). After the initial brushing and rinsing 

process is complete, place the woody snags in a 

separate bucket for about 1 hour. As the snags dry 

in the bucket, missed organisms will crawl out of 

crevices and then can be collected. Add these 

organisms to the rest of the sample material.

7. Combine the sample material collected from each 

woody snag and process the composited sample 

following the steps presented in "Sample Processing Procedures" (Section 5.5).

8. Measure the length and diameter of each woody 

snag to determine the area sampled. If only a portion of the woody snag was sampled, then the 

length and width of the area sampled is measured. 

Record these data on the "Semi-Quantitative 

Woody Snag Invertebrate Sample Field Data 

Sheet." Discard all woody snag sections after 

measurements are obtained.

Invertebrate Tip 2—Using a pump sprayer to 

rinse samples

Use a 1-gallon, home and garden pump 

sprayer to rinse sample material from woody snags. 

Since these sprayers are not high pressure, they are 

not likely to damage organisms. Using these sprayers during sample processing also can save time 

because the user does not have to continually refill a 

squirt bottle. The sprayer also can be used to dispense the 1-percent bleach solution for cleaning 

samplers and nets.
Field Protocols 41

5.3.3 Method for sampling fine-grained natural-bed 

substrates

In situations where coarse-grained natural-bed 

substrates or woody snags cannot be sampled, 

fine-grained natural-bed substrates (for example, sand) 

might be sampled as the RTH. Fine-grained sediments 

are typically sampled by using a grab sampler (fig. 8b, 

c). Grab samplers (for example, Ekman or Petite Ponar 

grabs) can be used in wadeable and nonwadeable 

streams. A hand or power winch is recommended for 

operating a weighted grab in deep water. All screening 

on the grab sampler must have mesh openings of 

500 μm. Make sure all crewmembers understand how 

to safely operate grab samplers.

Procedure for sampling fine-grained sediments with a 

grab sampler:

1. Identify locations throughout the reach where 

five discrete collections of fine-grained sediment 

can be taken. [Note: in some nonwadeable 

streams it might be necessary to determine the 

location of these sediments by using a simple 

sonar unit, such as those used by sport fishermen 

to locate schooling fish (Cuffney and others, 

1993)].

2. Carefully lower the grab sampler to the streambed. In nonwadeable streams, lower the sampler to within about 3 m of the streambed. Stop 

lowering the sampler, then drop it to the bottom. 

[Note: this deployment technique will minimize 

disturbance of the substrate by the descending 

sampler (Cuffney and others, 1993)]. Depending 

on the type of grab sampler being used, the jaw 

release mechanism will disengage when the sampler hits the streambed (Petite Ponar grab) or 

when a weighted messenger is dropped down the 

deployment line (Ekman grab).

3. Recover the grab sampler and inspect to ensure 

that the discrete collection was not lost because 

of debris catching in the jaws of the sampler.

4. Measure water depth, substrate type (dominant 

and codominant), percent substrate embeddedness, and velocity for each discrete collection. 

Record these data on the "Semi-Quantitative Targeted Habitat Invertebrate Sample Field Data 

Sheet."

5. Place the discrete collection in a 19-L (5-gal) 

white plastic bucket. Rinse sediment remaining 

inside the sampler into the bucket.

6. Repeat the collection procedure until all five discrete collections have been taken.

7. Process each discrete collection following the 

steps presented in "Sample Processing Procedures" (Section 5.5).

5.4 Qualitative multihabitat methods for sampling 

wadeable and nonwadeable streams

Discrete qualitative collections of invertebrates 

are made primarily using a D-frame kick net (fig. 8d). 

These collections are supplemented with visual collections that involve handpicking invertebrates from various substrates (for example, large cobbles, woody 

debris, leaf packs, and root wads) because some fragile 

invertebrates are easily damaged (for example, heptageniid mayflies) or are tightly adhered to a substrate 

(for example, some caddisfly larvae and pupae). Grab 

samplers might be used to collect burrowing invertebrates living in fine-grained sediments in nonwadeable 

streams. Sampling effort for collecting QMH samples 

is standardized by time; 1 hour is allocated for QMH 

sampling in each reach. After identifying which 

instream habitats (table 5) are present in the reach, the 

1-hour time limit is divided by the number of these 

habitats to determine the timed sampling effort in each 

habitat. [Note: this sampling time does not include time 

spent inspecting the net or moving around to different 

habitats.] For example, if 10 different instream habitats 

were present in reach A, then the time spent intensively 

sampling each habitat would be 6 minutes.

Qualitative sampling in wadeable and nonwadeable 

streams:

1. Identify the number and type of instream habitats 

present in the reach.

2. Determine the sampling time for each instream 

habitat type by dividing 1 hour by the number of 

habitat types present in the reach.

3. Using a D-frame kick net (with a 500-μm mesh 

net), sample each instream-habitat type by vigorously kicking, jabbing, dipping, or sweeping the 

substrate with the kick net. Handpick invertebrates from various substrates when necessary to 

collect fragile or tightly adhered invertebrates. 
42 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

Use a boat-operated grab sampler to collect burrowing invertebrates living in fine-grained sediments found in nonwadeable streams.

4. Place each discrete collection into a 19-L (5-gal), 

white plastic bucket. If the amount of material 

from each discrete collection is minimal, it might 

be easier to composite these collections before 

processing the sample in the field.

5. Process the QMH sample following the steps presented in "Sample Processing Procedures" (Section 5.5).

5.5 Sample processing procedures

The goal of processing invertebrate samples at 

the study site is to reduce the volume of sample and 

clean the composite sample that is submitted to the laboratory for analysis. Processing a single discrete collection or composited sample in the field can produce up 

to four different sample components: large-rare, 

main-body, split, and elutriate (fig. 9). These four components are derived from processing procedures that 

are implemented to reduce the amount of extraneous 

organic and inorganic debris that accumulates during 

collection or when two or more discrete collections are 

composited. The major tasks associated with processing invertebrate samples in the field are as follows:

1. Rinse and remove large debris (for example, cobbles, filamentous algae, leaves, and twigs) from 

each discrete collection.

2. Elutriate and sieve the discrete collection sample 

to separate invertebrates and organic debris (for 

example, detritus) from inorganic debris (for 

example, mineral sediments). Use 500-μm sieves 

[USA Standard Testing Sieve, American Society 

for Testing and Materials (ASTM) number 35 or 

Tyler equivalent 32 mesh] to elutriate and sieve 

samples. The invertebrates and organic debris 

form the main-body sample component; this component is required for laboratory analysis. The 

inorganic debris forms the elutriate sample component; this component is discarded in the field.

3. Remove large crayfish, hellgrammites, and mussels from each discrete collection. These organisms are removed because they can prey on or 

damage other invertebrates in the sample while 

the discrete collections are being processed. 

These organisms are tracked separately as the 

large-rare sample component. This component is 

produced only if these organisms are present in 

the sample; it is not required for laboratory analysis at the NWQL.

4. Reduce the volume of the main-body sample 

component, if necessary, to an acceptable volume 

(< 750 mL) that facilitates laboratory analysis. 

The sample volume is reduced by splitting the 

main-body sample component in half until an 

acceptable volume is obtained. One-half of the 

final split is designated as the main-body sample 

component and sent to the NWQL for analysis; 

the second half of the final split is designated as 

the split-sample component and is discarded in 

the field or retained at the field office, depending 

on study objectives.

5. Place the large-rare, main-body, and split 

(optional) sample components in labeled, standardized bottle types and preserve with 

10-percent buffered formalin.

6. Record site and sample information for each sample collected on a field data sheet.

Processing an invertebrate sample in the field:

1. Remove large debris (for example, cobbles, filamentous algae, leaves, and twigs) from each discrete collection and inspect for attached 

invertebrates, especially small cryptic invertebrates (for example, microcaddisflies and water 

penny beetle larvae).

a. Carefully remove any attached invertebrates 

by rinsing (fig. 10e) and place them in the 

main-body sample component (see Invertebrate Tips 3 and 4).

b. Discard the large debris.
Field Protocols 43

Discrete invertebrate

collections in separate

buckets

Sieve to rinse out fine sediments

Composite discrete collections

Remove large debris

and visually inspect for 

invertebrates

Main-body

sample component

volume > 750 mL?

Processed

Large-rare sample component

Split sample component

(discard in field or label, preserve,

and retain at field office)

Label sample

components and

log samples into

Bio-TDB

Prepare sample bottles

for shipping and

submit to NWQL for

analysis

Elutriate each discrete

collection

Elutriate sample component

(discard in field)

YES

NO

Split main-body

method until

volume < 750 mL

Place sample components in

bottles and preserve with 10%

buffered formalin

Processed

Main-body sample component

Inspect bucket for

invertebrates

Discrete collections

Detritus and invertebrates retained on

500 mm sieve

“Main-body sample component”

Inorganic debris

in buckets Invertebrates

Eluted organic

detritus and

invertebrates

Remove large rare invertebrates

Mussels, crayfish,

hellgramites

Main-body sample component

Figure 9. Steps followed in the field to process invertebrate samples in the National Water-Quality Assessment Program. 

mL, milliliter; μm, micrometer; <, less than; >, greater than; %, percent; Bio-TDB, Biological Transactional Data Base; NWQL,

National Water Quality Laboratory.
44 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

a. Two crewmembers taking a discrete collection using Slack sampler.

b. Removing attached invertebrates from 

large cobbles with a hand brush just 

upstream of the Slack sampler opening.

c. Using a hand rake to disturb fine substrates

to a depth of 0.1 m within the area template.

d. Transferring the discrete collection into a 

19-L (5-gal) plastic bucket.

e. Rinsing pieces of large organic 

material using a dual wash bucket.

Figure 10. Key steps in the collection and processing of invertebrate samples. (Photographs a-f by 

D. Sun).
Field Protocols 45

f. Elutriating the discrete collection using a 

bucket and sieve.

g. Visually examining elutriate sample 

component for cryptic organisms such 

as caddisflies that make cases out of sand.

h. Picking large-rare organisms from the RTH sample.

i. Sieving to remove fine sediment and evenly 

distribute material prior to splitting.

j. Using the sieve diameter splitting method to reduce 

sample volume to less than 750 mL. 

k. Placing the main-body sample component 

into a standard sample bottle.

Figure 10. Key steps in the collection and processing of invertebrate samples—Continued.

(Photographs g by J. Kennen, h by M. Ayers, i-k by D. Sun).
46 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

2. Elutriate each discrete collection onto a 500-μm 

mesh sieve to separate invertebrates and organic 

debris from the inorganic debris (fig. 10f). Doing 

this will minimize damage to organisms and 

maximize effective elutriation. However, if the 

volume of each discrete collection is < 250 mL, 

then it might be more efficient to composite all of 

the discrete collections and then elutriate the 

composited sample.

a. Place each discrete collection in a separate 

19-L (5-gal) plastic bucket filled about 

one-third full with water.

b. Carefully stir the contents of each bucket by 

hand to suspend as much of the sample material as possible.

c. Swirl the suspended sample material and carefully decant it onto a 500-μm sieve that is held 

over a dishpan. The dishpan provides secondary containment to catch any decanted sample that is spilled outside of the sieve. Pour 

any sample spilled in the dishpan directly onto 

the sieve unless it includes a large volume of 

inorganic debris. [Note: use a large-diameter 

(30-cm) sieve to facilitate subsequent sieving 

of the sample to remove fine sediments.]

d. As decanting proceeds, watch the advancing 

inorganic sediment front to avoid pouring it 

onto the sieve (fig. 10f).

e. Stop decanting when the inorganic sediment 

front reaches the lip of the bucket.

f. If sample is spilled in the dishpan, then pour 

the dishpan water (except fine sediment) back 

into the 19-L (5-gal) bucket. Otherwise, discard the dishpan water.

g. Repeat elutriation about three to five times for 

each discrete collection until the inorganic 

debris (elutriate sample component) appears 

free of detritus and organisms. [Note: heavy 

organisms such as mollusks and some 

case-building caddisflies might settle too fast 

during decantation and will not be able to be 

elutriated.]

h. Set the elutriate sample component contained 

in the bucket aside for later inspection.

i. Repeat the bucket elutriation for each discrete 

collection by elutriating onto the same sieve.

j. The invertebrates and organic debris that are 

retained on the 500-μm sieve represent the 

composited main-body sample component.

3. Inspect the elutriate sample component remaining in the bucket for invertebrates, particularly 

heavy organisms such as mollusks and some 

case-building caddisflies that often do not decant 

onto the sieve during elutriation.

Invertebrate Tip 3—Separating invertebrates 

from filamentous algae and moss

Removing invertebrates from filamentous 

algae and moss is difficult. Doing this after elutriation and compositing makes the removal task even 

more difficult and can even confound laboratory 

processing if large clumps of filamentous algae are 

present in the submitted samples. However, organisms can be effectively removed from filamentous 

algae by gently agitating the algae or moss clumps 

for a few seconds in a bucket filled about one-third 

full with water. Periodically inspect the clumps to 

determine if more agitation is necessary. Pour the 

organisms remaining in the bucket onto the 500-μm 

sieve after elutriation.

Invertebrate Tip 4—Dual bucket sieving and 

washing technique

Large quantities of large debris (for example, 

leaves and woody debris) can be quickly removed 

from the main-body sample component by using a 

dual bucket sieving and washing technique. Prepare 

a 10-L (2.5-gal) dunking bucket by replacing the 

bottom with a 1.3-cm (0.5-in) wire mesh screen 

(fig. 10e); make sure the dunking bucket fits easily 

inside one of the 19-L (5-gal) buckets used to elutriate samples. Fill the 19-L (5-gal) bucket two-thirds 

full of water. Add portions of the main-body sample 

component to the dunking bucket while it is inside 

the 19-L (5-gal) bucket. Carefully work the dunking 

bucket to sieve the fine organic debris and invertebrates, leaving behind the rinsed large debris. Carefully pour the contents of the bucket into a 500-μm 

sieve and discard the large debris retained by the 

dunking bucket after it has been quickly scanned for 

invertebrates.
Field Protocols 47

a. Place a small amount of the elutriate sample 

component in a shallow white tray containing 

enough water to float debris to facilitate detection of invertebrates. Alternatively, the elutriate sample component can be inspected in a 

sieve (fig. 10g).

b. If necessary, sieve the elutriate sample component through a pair of nested large-meshed 

(for example, 2- and 4-mm) sieves to separate 

the larger gravel and pebble material and 

expedite the inspection and removal of invertebrates.

c. Place cryptic invertebrates (for example, 

elmid beetle larvae and case-building caddisflies) in the main-body sample component.

4. Inspect the main-body component for large rare 

invertebrates (for example, mussels, crayfish, and 

hellgrammites) (fig. 10h) that might prey upon or 

damage other invertebrates in the collection.

a. Remove only these organisms and place them 

in a separate bottle identified as the large-rare 

sample component. Match the bottle size to 

the size and quantity (typically no more than 

20) of organisms. [Note: if these organisms 

are not present in the collection, then do not 

prepare a large-rare sample component.]

b. Do not add clumps of sample debris to the 

bottle. This debris often contains other invertebrates, especially early instar specimens, 

and interferes with laboratory analysis.

5. Wash the main-body sample component remaining on the 500-μm sieve by dipping and swirling 

the sieve repeatedly in the stream (fig. 10i) or a 

dishpan. Washing and sieving removes fine sediment from the main-body sample component, 

which facilitates laboratory analysis.

6. Reduce the volume of main-body sample component if it exceeds 750 mL by using the sieve 

diameter splitting method (fig.10j).

a. Ensure that large debris and large-rare organisms have been removed from the main-body 

sample component.

b. Use a standard 20- or 30-cm diameter metal 

sieve marked with six equally spaced (30 

degrees apart) diameters (see fig. 10k). The 

diameter markings are extended up the inside 

walls of the sieve and numbered 1 through 6.

c. Evenly distribute the main-body sample component on the 500-μm sieve by immersing the 

sieve in water, redistributing the sample, and 

then carefully lifting the sieve level out of the 

water so the sample remains in place on the 

sieve (fig. 10i).

d. Roll a die to select randomly which of the six 

diameters will be used to divide the sample in 

half.

e. Using a ruler and putty knife, divide the 

main-body sample component along the randomly selected diameter (fig.10j). Determine 

in an unbiased manner (for example, flipping 

a coin), which half becomes the new, reduced 

main-body sample component; the corresponding half is designated as the split sample 

component.

f. Repeat the sieve diameter splitting method on 

each new main-body sample component until 

a volume < 750 mL is obtained. Discard each 

split sample component that is produced up to 

the final split. Track and record the number of 

splits performed; this information is required 

to determine the total sample abundances for 

individual invertebrate taxa after laboratory 

analysis.

g. Place the final main-body sample component 

in a single appropriately sized wide-mouth 

plastic bottle (for example, NalgeneTM) that 

does not exceed 1 L in capacity (fig. 10k). 

[Note: ensure that sufficient space remains in 

the bottle for adding buffered formalin to preserve the sample.]

h. The final split-sample component may be discarded in the field or preserved and retained at 

the study office if the integrity of the 

main-body sample component is compromised during shipping or laboratory analysis.

7. Preserve the large-rare, main-body, and split sample components with 10-percent buffered formalin.
48 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

a. Add enough 10-percent buffered formalin to 

bring the total volume to within 2 cm of the 

top of the bottle.

b. Cap the bottle and carefully invert it several 

times to mix the sample and buffered formalin.

c. Open the bottle and top off with additional 

10-percent buffered formalin. This step minimizes large void spaces that can cause 

increased agitation of the sample (especially 

during shipping) and potentially damage fragile invertebrates.

8. Place a sample label on the outside of the 

main-body, split, and large-rare sample component bottles.

9. Record collection and sample information on the 

appropriate field data sheet corresponding to a 

particular sample type.

5.6 Field data sheets

Three different field data sheets are used to 

record site and sample information associated with 

invertebrate sample collection. The field data sheets are 

presented in the following order at the end of this section:

• Semi-Quantitative Targeted Habitat Invertebrate 

Sample Field Data Sheet (SQTH; 2 pages)

• Semi-Quantitative Woody Snag Invertebrate Sample Field Data Sheet (SQWS; 2 pages)

• Qualitative Multihabitat Invertebrate Sample Field 

Data Sheet (QMH; 2 pages)

Information that is recorded on each field data 

sheet is organized into sections (table 6) that correspond to data entry screens in Bio-TDB. Except for 

these section labels (for example, "Sample Header"), 

data fields or subsections that are shown in bold face 

are required for entry into Bio-TDB. Consult the 

Bio-TDB User Manual (Ruhl and Scudder, accessed 

April 26, 2002) for additional information on entering 

field data into the data base.

Before leaving the sampling site, the team leader 

should review each field data sheet for completeness 

and accuracy by comparing information on sample bottle labels and field data sheets, especially in the "Sampling Information" section. All missing or incorrect 

information should be completed or corrected. Supplies 

of each different field data sheet (blank and completed) 

should be stored in separate resealable bags for protection from water.

Section This section is used to 

document

Completed on field data 

sheet?

SQTH SQWS QMH

Sample header Where and when a sample 

was collected Yes Yes Yes

Related sampling 

activities

Other sampling activities Yes Yes Yes

Physical site 

conditions

Physical and chemical conditions at the time of sampling

Yes Yes Yes

Sampling 

information

How a sample is collected 

and processed in the field Yes Yes Yes

Instream habitat 

types sampled

Which instream habitats 

were present and sampled 

by indicating the sampling 

gear used

Yes Yes Yes

Supporting 

information

Ancillary sampling information taken in conjunction with discrete 

collections

Yes Yes No

Table 6. Explanation of field data-sheet sections used to record 

information about invertebrate sampling activities

[SQTH, semi-quantitative targeted habitat sample; SQWS, semi-quantitative 

woody snag sample; QMH, qualitative multihabitat sample]
Field Protocols 49

Page # Cumulative Page # .

Date field data reviewed: / / by (initials)

Date entered into Bio-TDB: / / by (initials)

Bio-TDB Sample ID

Semi-Quantitative Targeted Habitat Invertebrate Sample

Field Data Sheet

SAMPLE HEADER

Station: Date (MM/DD/YYYY):

/ /

Waterbody:

Station description:

Station ID: Reach: Start time: h

Collector (leader): End time: h

Sample and photographic notes:

RELATED SAMPLING ACTIVITIES 

(circle all that apply)

Water chemistry Discharge Habitat survey Tissues

Algae survey Fish survey Bed sediment Other (specify):

PHYSICAL SITE CONDITIONS

Clouds: % Wind (circle): Calm Light Moderate Gusty

Precipitation (circle): None Rain Sleet Snow Precipitation Intensity (circle): N/A Light Moderate Heavy

Other weather:

Beginning water measurements Ending water measurements

Time: Time:

Water temperature

(°C):

Water temperature

(°C):

pH: pH:

Dissolved oxygen

(mg/L):

Dissolved oxygen

(mg/L):

Specific conductance

[(μS/cm) @ 25 °C]:

Specific conductance

[(μS/cm) @ 25 °C]:

Turbidity

(NTUs)

Turbidity

(NTUs)

Discharge (ft3/s): Stream stage: ft @ time h

Riparian shading (circle): Shaded Partial Full sun

Water clarity (circle): Clear Slightly turbid Turbid Very turbid

Water color (circle): Black Brown Clear Dark green Light green Yellow

Physical site condition comments:
50 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

SAMPLING INFORMATION

Primary sample or Repeated sampling replicate? (circle one) Sample Number: Mesh size: 500 μm

Number of discrete collections constituting composite: Total area of sample: cm2

Elutriation method (circle): Bucket Other: Sample-splitting method (circle): Not split Sieve diameter

Sample bottle (complete all)

Sample 

component

Preservative Split ratio Bottle sequence Field SMCOD

Large-rare 10% buffered formalin 1/1

Main-body 10% buffered formalin

Subbottle comments:

Invertebrate SQTH Sample Page # Cumulative Page # .

SUPPORTING INFORMATION

Substrate type codes: (1) smooth bedrock/concrete/hardpan; (2) silt, clay, marl, muck, organic detritus; (3) sand; (4) fine/medium gravel; (5) coarse gravel; (6) Very 

coarse gravel; (7) small cobble; (8) large cobble; (9) small boulder; (10) large boulder/ irregular bedrock/irregular hardpan/irregular artificial surface

Velocity meter used (circle one): Pygmy AA Marsh McBirney Other:

Sample 

location 

number

Water depth 

(m)

Substrate

embeddedness

(percent)

Velocity

(ft/s)

Substrate type

Dominant Co-dominant

1

2

3

4

5

INSTREAM HABITAT TYPES SAMPLED

Indicate the habitat types sampled by entering the sampler code used. Enter NA if the habitat type is not present or NC if the habitat type was present but not sampled. 

Blackened cells indicated habitat type not possible. Sampler codes: (2) Slack sampler; (13) Standard Ekman; (17) Petite Ponar; Other (specify): 

Riffle Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin

Run Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin

Pool Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin
Field Protocols 51

Page # Cumulative Page # .

Date field data reviewed: / / by (initials)

Date entered into Bio-TDB: / / by (initials)

Bio-TDB Sample ID

Semi-Quantitative Woody Snag Invertebrate Sample

Field Data Sheet

SAMPLE HEADER

Station: Date (MM/DD/YYYY):

/ /

Waterbody:

Station description:

Station ID: Reach: Start time: h

Collector (leader): End time: h

Sample and photographic notes:

RELATED SAMPLING ACTIVITIES 

(circle all that apply)

Water chemistry Discharge Habitat survey Tissues

Algae survey Fish survey Bed sediment Other (specify):

PHYSICAL SITE CONDITIONS

Clouds: % Wind (circle): Calm Light Moderate Gusty

Precipitation (circle): None Rain Sleet Snow Precipitation Intensity (circle): N/A Light Moderate Heavy

Other weather:

Beginning water measurements Ending water measurements

Time: Time:

Water temperature

(°C):

Water temperature

(°C):

pH: pH:

Dissolved oxygen

(mg/L):

Dissolved oxygen

(mg/L):

Specific conductance

[(μS/cm) @ 25 °C]:

Specific conductance

[(μS/cm) @ 25 °C]:

Turbidity

(NTUs)

Turbidity

(NTUs)

Discharge (ft3/s): Stream stage: ft @ time h

Riparian shading (circle): Shaded Partial Full sun

Water clarity (circle): Clear Slightly turbid Turbid Very turbid

Water color (circle): Black Brown Clear Dark green Light green Yellow

Physical site condition comments:
52 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

SAMPLING INFORMATION

Primary sample or Repeated sampling replicate? (circle one) Sample Number: Mesh size: 500 μm

Number of discrete collections constituting composite: Total area of sample: cm2

Elutriation method (circle): Bucket Other: Sample-splitting method (circle): Not split Sieve diameter

Sample bottle (complete all)

Sample 

component

Preservative Split ratio Bottle sequence Field SMCOD

Large-rare 10% buffered formalin 1/1

Main-body 10% buffered formalin

Subbottle comments:

Invertebrate SQWS Sample Page # Cumulative Page # .

SUPPORTING INFORMATION

Substrate type codes: (1) smooth bedrock/concrete/hardpan; (2) silt, clay, marl, muck, organic detritus; (3) sand; (4) fine/medium gravel; (5) coarse gravel; (6) Very 

coarse gravel; (7) small cobble; (8) large cobble; (9) small boulder; (10) large boulder/ irregular bedrock/irregular hardpan/irregular artificial surface

Velocity meter used (circle one): Pygmy AA Marsh McBirney Other:

Sample 

location 

number

Woody

snag

number

Snag

length

(cm)

Snag

diameter

(cm)

Velocity

(ft/s)

Substrate

Type

Water depth (m) from surface to:

Woody snag Bottom

1

2

3

4

5

6

7

8

9

10

INSTREAM HABITAT TYPES SAMPLED

Indicate the habitat types sampled by entering the sampler code used. Enter NA if the habitat type is not present or NC if the habitat type was present but not sampled. 

Blackened cells indicate habitat type not possible. Sampler codes: (27) Snag collection with Slack sampler; Other (specify):

Riffle Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin

Run Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin

Pool Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin
Field Protocols 53

Page # Cumulative Page # .

Date field data reviewed: / / by (initials)

Date entered into Bio-TDB: / / by (initials)

Bio-TDB Sample ID

Qualitative Multihabitat Invertebrate Sample

Field Data Sheet

SAMPLE HEADER

Station: Date (MM/DD/YYYY):

/ /

Waterbody:

Station description:

Station ID: Reach: Start time: h

Collector (leader): End time: h

Sample and photographic notes:

RELATED SAMPLING ACTIVITIES 

(circle all that apply)

Water chemistry Discharge Habitat survey Tissues

Algae survey Fish survey Bed sediment Other (specify):

PHYSICAL SITE CONDITIONS

Clouds: % Wind (circle): Calm Light Moderate Gusty

Precipitation (circle): None Rain Sleet Snow Precipitation Intensity (circle): N/A Light Moderate Heavy

Other weather:

Beginning water measurements Ending water measurements

Time: Time:

Water temperature

(°C):

Water temperature

(°C):

pH: pH:

Dissolved oxygen

(mg/L):

Dissolved oxygen

(mg/L):

Specific conductance

[(μS/cm) @ 25 °C]:

Specific conductance

[(μS/cm) @ 25 °C]:

Turbidity

(NTUs)

Turbidity

(NTUs)

Discharge (ft3/s): Stream stage: ft @ time h

Riparian shading (circle): Shaded Partial Full sun

Water clarity (circle): Clear Slightly turbid Turbid Very turbid

Water color (circle): Black Brown Clear Dark green Light green Yellow

Physical site condition comments:
54 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

SAMPLING INFORMATION

Primary sample or Repeated sampling replicate? (circle one) Sample Number:

Equal Sampling Effort Procedure: 1 hr total sampling time Mesh size: 500 μm

Elutriation method (circle): Bucket Other: Sample-splitting method (circle): Not split Sieve diameter

Sample bottle (complete all)

Sample 

component

Preservative Split ratio Bottle sequence Field SMCOD

Large-rare 10% buffered formalin 1/1

Main-body 10% buffered formalin

Subbottle comments:

Invertebrate QMH Sample Page # Cumulative Page # .

INSTREAM HABITAT TYPES SAMPLED

Indicate the habitat types sampled by entering the sampler code used; separate multiple sampler codes for a single habitat type with a comma (for example, 7,22,23). 

Enter NA if the habitat type is not present or NC if the habitat type was present but not sampled. Blackened cells indicate habitat type not possible. Sampler codes: (2) 

Slack sampler; (7) D-frame net; (13) Standard Ekman grab; (17) Petite Ponar grab; (22) Visual collection wood; (23) Visual collection leaf debris; (24) Visual collection cobbles; (25) Visual collection other; Other (specify):

Riffle Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin

Run Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin

Pool Natural

bed

Manufactured

bed Slough Macrophytebed

Woody

snag

Bar

Main channel

Channel margin

Island margin
Field Protocols 55

6. FISH SAMPLING PROTOCOLS

The fish sampling protocols described in this 

report present methods for collecting a representative 

sample of the fish community from the stream. Such a 

sample contains most, if not all, species in the stream at 

the time of sampling in numbers proportional to their 

actual abundance. The reach is the representative portion of the stream that is sampled. Each reach contains 

various instream habitats consisting of different geomorphic channel units, substrates, and hydrologic conditions. Fish species are distributed in the stream reach 

according to these instream habitats. No single fish collection gear or method is adequate to sample the diversity of habitats found in the reach. Therefore, two 

complementary methods are used for collecting fish: 

electrofishing and seining. Electrofishing is conducted 

in two separate passes of the reach; the fish collected 

from the first pass are processed before the second pass 

is conducted. Seining is performed after electrofishing. 

Three seine collections (hauls or kicks) are taken and 

combined before processing fish. The fish sampling 

protocols present techniques for applying these sampling methods and procedures for processing fish specimens in the field to determine total length, weight, 

and external anomalies. These revised fish sampling 

protocols are based on the original guidance presented 

by Meador and others (1993) and the QA protocols by 

Walsh and Meador (1998).

6.1 Overview of sampling methods

6.1.1 Electrofishing

Electrofishing is the use of electricity to capture 

fish. A high-voltage potential is applied between two or 

more electrodes that are placed in the water. The voltage potential is created with either direct current or 

alternating current; only direct current is used in the 

NAWQA Program. Direct current produces a unidirectional, constant electrical current. Pulsed direct current, 

a modified direct current, produces a unidirectional 

electrical current composed of a sequence of cyclic 

impulses (Meador and others, 1993).

The frequency of the pulses produced when 

using pulsed direct current can be adjusted by the operator and usually ranges from 15 to 120 pulses per second (pps). High pulse frequencies (> 30 pps) have 

proven to be more effective in collecting fish but appear 

to cause spinal injuries, particularly in trout and salmon 

species (Coffelt Manufacturing, Incorporated, cited in 

Meador and others, 1993). The injury rate depends on 

size; larger fish are more susceptible to injury than 

smaller fish (McMichael and others, 1998). Pulse rates 

< 30 pps have caused low incidence of injury, but are 

generally ineffective in collecting fish. Therefore, a 

pulse-rate range from 30 to 60 pps is recommended to 

provide maximum collection effectiveness with a minimum potential for damage to fish.

Water conductivity also affects the response of 

the fish to the electrical field and is the single most 

important limiting factor in electrofishing effectiveness 

(see Fish Tip 1). Low-conductivity water is highly 

resistant to the flow of electrical current, thereby reducing the amount of electrical current traveling through 

the water and passing through the body of the fish. 

Under such conditions the electrical field is limited to 

the immediate area of the electrode. High-conductivity 

water produces the opposite effect by concentrating a 

narrow electrical field between the electrodes (Meador 

and others, 1993). The conductivity of the water must 

be measured prior to electrofishing to determine the 

appropriate output voltage for effective sampling. 

Although electrofishing is viewed as the single most 

effective method for sampling stream fish communities 

(Bagenal, 1978; Plafkin and others, 1989), it is biased 

toward collection of large-sized fish (Wiley and Tsai, 

1983). Therefore, electrofishing alone should not be 

used to determine fish community structure (Reynolds, 

1983).

Water clarity also affects electrofishing success 

and determines which techniques will be used. In clear 

streams, fish can see the electrofishing crew or boat. 

Fish Tip 1—Settings for electrofishing gear

As a general rule, the output from the electrofishing gear should be about 3,000 watts. Voltage 

times amperage equals wattage. In low-conductivity 

water, high voltage and low amperage are needed to 

stun fish, while in high-conductivity water, low voltage and high amperage are needed to achieve the 

same result. In high-conductivity water, output voltage must be reduced to minimize potential damage 

to the fish. For electrofishing gear with both voltage 

adjustments and ammeters, achieving the proper 

output is straightforward. For electrofishing gear 

without ammeters, some pre-sampling experimentation is necessary.
56 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

Evasion will be the response to both the electrical field 

and the presence of the crewmembers. In turbid 

streams, stunned fish may be difficult to see. Also, 

streams with clay or silt substrates will require additional amperage because the suspended particles create 

a situation similar to increased conductivity.

6.1.2 Seining

Seining is a common method for sampling 

stream fish communities (Bagenal, 1978; Nielsen and 

Johnson, 1983), and it is used to complement electrofishing collections. Unlike electrofishing, seining is a 

highly effective method for sampling small-sized individuals < 10-cm total length (Bayley and Herendeen, 

2000). Seining is always conducted following electrofishing, except for instances when seines are used as 

barriers to fish escaping an electrical current.

Seines are sampling devices that trap fish by 

enclosing or encircling them. Seines are manufactured 

in a variety of dimensions and mesh sizes. The NAWQA 

Program uses 6.4 mm as a standard mesh size for 

seines. Three sizes of seines are commonly used to 

sample fish communities: 3 x 1.2 m; 7.6 or 9.1 x 1.2 m; 

and about 30.5 to 61 x 1.8 m. The 3 x 1.2-m seine is 

referred to as a "common sense" seine (Hendricks and 

others, 1980; Bryan, 1984), a "minnow" seine, or a 

"standard ichthyological collection" seine, and is 

attached to two brails. The 7.6- or 9.1-m seine typically 

has a bag or pocket in the center of the seine (the bunt) 

and, thus, is referred to as a bag seine. As the bag seine 

is pulled through the water, fish are herded toward the 

center of the net and into the bag. A beach seine is typically used along the shorelines of large bodies of water 

and is usually > 30 m long. Because of the greater 

length, larger dimension brails (usually 51 mm x 

51 mm) are required for the beach seine to maximize 

sampling effectiveness and maintain durability.

Water clarity can have a profound effect on seining success. In clear water, fish can see the seine and 

will actively avoid it either by swimming around the 

net or by swimming out of the bag and under any gaps 

between the lead line and stream bottom. The seine 

should be hauled immediately if the crewmembers see 

fish escaping the net. Additional crewmembers are 

used to herd fish back into the seine when the water is 

clear.

6.2 Sampling methods for wadeable streams

6.2.1 Electrofishing techniques for wadeable streams

Backpack and towed electrofishing gears are 

used for sampling fish in wadeable streams. Backpack 

electrofishing (with a single anode) is usually most 

effective in shallow (< 1 m), narrow (< 5 m wide) 

streams. Towed electrofishing gear (multiple anodes) is 

usually more effective in wide (> 5 m) wadeable 

streams with pools deeper than 1 m. Channel width, 

depth, and access should be considered before choosing between backpack and towed electrofishing methods.

Electrofishing techniques for wadeable streams 

require an electrofishing crew consisting of three to six 

individuals. When using backpack electrofishing gear, 

one crewmember is designated as the operator; with 

towed gear, three crewmembers are designated as operators. Regardless of the gear used, two crewmembers 

(or netters) are assigned to collect stunned fish with dip 

nets. One additional crewmember is sometimes needed 

to transfer netted fish into a flow-through holding bottle (or live cage). All crewmembers must wear polarized glasses to enhance their ability to see fish that 

have been stunned by the electrical field. Techniques 

for collecting samples using either backpack or towed 

electrofishing gear are generally similar in riffle-pool 

streams and meandering streams. Regardless of the 

electrofishing gear used, two separate electrofishing 

passes are made in the reach. The fish collected in the 

first pass are processed before the second electrofishing pass begins; fish community data for each pass are 

kept separate.

6.2.1.1 Sampling direction

Sampling begins at the downstream boundary of 

the sampling reach and proceeds upstream. Sampling 

in an upstream direction in wadeable streams is preferred over sampling in a downstream direction (Hendricks and others, 1980). Disturbance caused by 

electrofishing crews walking in the stream increases 

turbidity, thereby greatly reducing visibility and collection success. Also, sampling in an upstream direction 

allows stunned fish to drift downstream, thus facilitating their capture by the netters. The distance of the 

netters from the anode increases with current velocity 

and turbidity.
Field Protocols 57

6.2.1.2 Sampling all instream habitat features

All geomorphic channel units and instream habitat features, such as woody snags, undercut banks, 

macrophyte beds, or large boulders within the wadeable sampling reach, are sampled using pulsed direct 

current. This technique might require electrofishing 

from one shoreline to the other in a "zigzag" pattern, 

while consistently sampling all areas within the reach.

6.2.1.3 Electrofishing riffle-pool streams

Continuous application of electrical current — herding 

fish by the operator:

A common electrofishing technique is to apply a 

continuous electrical current to the water. Fish generally respond to continuously applied electrical current 

by attempting to avoid exposure to the electrical field. 

Thus, continuous application of electricity can result in 

fish moving just ahead and away from the operator. The 

operator should be aware of this response and take 

advantage of natural barriers to herd fish into (for 

example, banks, bars, or shallow riffles) and facilitate 

their capture by the netters. However, upon reaching a 

barrier, fish will turn and attempt to evade the 

approaching electrical current. Therefore, netters 

should be alert when approaching barriers so as to 

avoid missing fleeing fish.

Intermittent application of electrical current — herding 

fish to the operator:

In runs or long pools, fish often evade the weak 

edge of the electrical field by fleeing upstream. One 

technique that is used to counter this response is to use 

an intermittent application of current and then herd fish 

back toward the operator. A crewmember (or two) 

moves upstream of the fish by getting out of the stream 

and walking up the bank. Once ahead of the fish, the 

crewmember returns to the stream and creates a disturbance in the water that drives the fish downstream 

toward the operator. The electricity is turned off as the 

crewmember herds the fish back downstream. When 

the fish are visible and close to the anode, the electricity is turned on to stun them.

Sampling shallow riffles:

In sampling shallow riffles, the operator sweeps 

the anode across the riffle from upstream to downstream while walking across the riffle. Crewmembers, 

positioned downstream of the operator, hold dip nets or 

preferably a small seine on the bottom which allows the 

streamflow to carry stunned fish into their nets. This 

technique minimizes escape and avoidance of the electrical field by benthic fish species (for example, darters 

and sculpins), which commonly inhabit riffles. These 

species may not be seen until the net is examined. 

Sculpins and darters do not have swim bladders. Therefore, unlike other species that do have swim bladders, 

species in these groups do not float when stunned, but 

rather roll along the bottom with the water current.

Sampling rocky riffles, cascades, and torrents:

In sampling rocky, swift riffles, a crewmember 

holds a dip net about 1 m below the anode. The operator works the anode downstream through the swift current between rocks and through small plunge pools 

while the net is maintained on the bottom at a constant 

distance from the anode.

6.2.1.4 Electrofishing meandering streams

Electrofishing in meandering streams can be 

challenging because there are few natural barriers (for 

example, a shallow riffle) to hold herded fish. Therefore, the intermittent use of electrical current is more 

appropriate than continuous current. Intermittent electrical current acts as a herding stimulus to corral fish 

into holding areas along the banks. Block nets may be 

used to isolate the stream reach during electrofishing. 

However, there are several factors to consider before 

using block nets. Proper deployment of block nets is a 

time-consuming activity. The lead line must be 

anchored securely to the stream bottom to prevent fish 

from escaping under the net. The top of the net is usually secured to trees or other structures on the stream 

bank so that it is taut and above the stream surface to 

prevent fish from jumping over the net. The potential 

for entanglement in the net mesh also could result in 

the mortality of small fish. If block nets are used, they 

must be used in the same manner during repeated sampling of the reach (see also Fish Tip 2).
58 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

Sampling Instream Habitat:

A different electrofishing technique is required 

in areas of a meandering stream that have complex 

instream habitat (for example, submerged trees). Fish 

congregating near these habitat features either will disperse or will be difficult to remove after being stunned 

by continuous application of electricity. An effective 

technique for capturing fish associated with complex 

instream habitat is for the operator to approach the habitat feature with the electrical current off. Next, in 

quick succession, the anode is thrust close to the habitat 

feature, the electrical current turned on, and then the 

anode is withdrawn in a sweeping motion away from 

the habitat feature. This technique produces a physiological response by fish called galvanotaxis, where the 

electrical current forces fish to swim towards the anode 

(Meador and others, 1993).

6.2.2 Seining techniques for wadeable streams

Electrofishing in wadeable streams is complemented by various seining techniques using common 

sense and bag seines. Use of a particular seine depends 

on the geomorphic channel units, channel features, and 

instream habitats present in the sampling reach. Areas 

having submerged objects (for example, woody snags 

or cobbles) make seining difficult. Therefore, the 

potential for collecting a representative sample should 

be evaluated before seining these areas. Regardless of 

the seining techniques used, three seine collections are 

taken and combined before processing fish.

6.2.2.1 Seining riffles

Riffle dwelling species (for example, darters and 

sculpins) are sampled by using a kick seining technique 

(Hendricks and others, 1980; Matthews, 1986; Bramblett and Fausch, 1991). Kick seining is an effective 

technique for collecting these species because it 

involves disturbing the substrate and letting the water 

current carry fish into a common sense seine. Two 

crewmembers enter the stream below a riffle and hold 

the seine in a vertical position above the water and perpendicular to the flow at the downstream edge of a riffle. The crewmembers then thrust the brails and lead 

line of the seine to the stream bottom. The brails are 

slightly angled downstream so that the flow forms a 

slight pocket in the seine. Upstream of the seine, a third 

crewmember vigorously disturbs (or kicks) the substrate while moving toward the seine (see also Fish Tip 

3). After reaching the seine, crewmembers lift the seine 

out of the water and sort fish from the collected debris. 

Three kick seine collections are made across the riffle. 

The fish from these collections are combined to form 

the sample.

6.2.2.2 Seining meandering streams

Meandering streams are sampled by using either 

a common sense or bag seine, depending on the channel width and the type of electrofishing gear used. Generally, if the reach is sampled with backpack 

electrofishing gear, then a common sense seine is used. 

If the reach is sampled with towed electrofishing gear, 

then a bag seine is used. Seining downstream has been 

demonstrated to be the most effective technique (Hendricks and others, 1980).

Seining speed should be slightly faster than the 

stream current. Faster seining speeds are avoided 

because water will be pushed in front of the seine, 

thereby forcing fish away from the seine. Maintaining 

the lead line in contact with the bottom and a substantial bow in the seine (accomplished by angling the 

brails back to keep the net bottom well forward of the 

net top) will minimize the potential for escaping fish 

until the net is beached or lifted. When the seine haul is 

finished, the seine is beached by dragging it onto the 

shore. When there is only a small shoreline area to 

beach the seine, the brails are brought close together at 

Fish Tip 2—Crewmembers as barriers to fish 

movement

Use crewmembers as stationary or mobile barriers. Crewmembers are positioned across the reach 

upstream of the electrofishing operator. As the operator approaches, the crewmembers create a splashing 

disturbance in the water that prevents fish from 

swimming past the barrier. Alternatively, crewmembers can become a mobile barrier by herding fish 

downstream to the operator. Use of a small seine as 

an additional barrier improves the efficiency of this 

technique.

Fish Tip 3—Kick seining and electrofishing

If kick seining alone is not effective, try using 

backpack electrofishing gear to improve the capture 

of benthic species. Work the anode back and forth 

while a crewmember disturbs the substrate. Turn the 

power off when the anode reaches the seine.
Field Protocols 59

the shoreline and the lead line slowly pulled into shore 

by hand.

If the seine cannot be beached, the bottoms of the 

brails are brought as close to shore as possible. Then in 

one simultaneous motion, the seine is quickly stretched 

between the brails, lifted out of the water, and carried 

onto shore. Three seine hauls, each covering an area of 

about 30 m, are taken throughout (upper, middle, and 

lower sections) the reach. The fish from these hauls are 

combined to form the sample.

6.3 Sampling methods for nonwadeable streams

6.3.1 Electrofishing nonwadeable streams

Nonwadeable streams are sampled using electrofishing boats. An electrofishing boat crew consists of a 

driver and one or two persons who collect the fish with 

dip nets. The driver should be skilled at maneuvering 

the boat as effectively as possible to allow crewmembers the best opportunity to capture stunned fish. As 

with wadeable electrofishing methods, all crewmembers should wear polarized sunglasses. Two separate 

electrofishing passes, one along each bank, are made 

in the reach. The fish collected in the first pass are processed before the second electrofishing pass begins; 

fish community data for each pass are kept separate.

6.3.1.1 Sampling direction and boat speed

Sampling begins at the upstream boundary of the 

sampling reach and proceeds downstream by maneuvering the boat along one shoreline. The electrofishing 

boat is operated at a speed equal to or slightly greater 

than the current velocity. The shoreline sampled during 

the first pass can be decided by flipping a coin. Sampling is conducted downstream because fish are usually 

oriented upstream (into the current) and will either 

swim into the approaching electrical field or turn to 

escape downstream. In turning to escape downstream, 

fish orient themselves perpendicular to the electrical 

field, thereby exposing a greater surface area of the fish 

to the electrical field that renders them more susceptible to capture. Also, when fish are stunned they are carried downstream by the flow, providing greater 

opportunity for capture. Sampling in a downstream 

direction is more efficient than sampling in an 

upstream direction (Ohio Environmental Protection 

Agency, 1987) in nonwadeable streams (see Fish Tip 

4).

6.3.1.2 Electrofishing Techniques for Nonwadeable 

Streams 

A continuous application of current while drifting downstream generally is used for most large, nonwadeable streams. In areas with clear or shallow water, 

intermittent application of current may be more effective than continuous application. Fish are approached 

with the current off. When the anodes are in position 

near the fish, the current is applied.

6.3.1.3 Boat electrofishing techniques for sampling fish 

from instream habitat 

Instream habitat features along the shoreline are 

sampled by maneuvering the boat close to the habitat 

feature with the electrical current off. As the anode is 

placed near the habitat feature, the electrical field is 

generated and the boat is backed away from the habitat 

feature (see Fish Tip 5). The fish thus are "pulled" 

away from the habitat feature to facilitate their capture. 

Captured fish are placed in a live well on the boat and 

processed after completion of the first electrofishing 

pass.

6.3.2 Seining nonwadeable streams

Nonwadeable streams can be sampled by using a 

beach seine in wadeable shoreline areas, if present. 

Like the bag seine, the exact length of the beach seine 

should be determined after consultation with local fish 

ecologists. Three seine hauls should be taken from 

accessible parts of the upper, middle, and lower secFish Tip 4—Boat position

Position the boat so the bow is angled downstream and toward the bank. This allows the boat 

operator to reverse direction (generally upstream and 

away from the banks) and not pass over stunned fish. 

Periodically, the boat should be slowed to less than 

current speed so that fish drifting with the current 

may be more easily observed.

Fish Tip 5—Current application time

When boat electrofishing, the duration for 

applying electrical current should be increased at 

submerged structures. Fish located in deeper water 

(2–3 m) may require 5–10 seconds of current before 

a response is observed. This duration increases as 

water temperature decreases.
60 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

tions of the nonwadeable sampling reach. The fish from 

these hauls are combined to form the sample.

6.3.2.1 Beach seining

Beach seining is conducted by keeping one end 

of the seine stationary on the shore while the remainder 

of the seine is deployed into the water so that it is 

roughly perpendicular to the shoreline. The seine is 

swung in a downstream direction while pivoting on the 

stationary beach end.

6.3.2.2 Shoreline seining

The shoreline is seined by pulling the seine 

downstream along the bank. Keep one end of the seine 

near the shoreline and the other end extended out from 

the bank and angled downstream. The angle will prevent fish associated with the shallow water near the 

bank from escaping the net by fleeing directly into 

deeper water. To obtain a good seine haul, the lead line 

must remain in contact with the river bottom throughout the haul. If the site is too deep to wade at the 

extremes of the net, another site should be used. If 

another site cannot be found, the seine should only be 

used where the water is shallow enough to ensure constant bottom contact. If seining is not possible or cannot be conducted safely, an alternative method can be 

used.

6.4 Alternative sampling methods

Although electrofishing and seining represent the 

standard methods for collecting a representative fish 

community sample from a reach, there may be occasions when these methods cannot be used effectively. 

For example, electrofishing is ineffective when specific 

conductivity is < 20 μS/cm. Likewise, seining is hampered when a large number of woody snags and other 

types of irregular debris are present in the reach that 

would prevent maintaining the lead line on the stream 

bottom. When physical conditions in the reach prohibit 

the use of these standard methods, alternative sampling 

methods (for example, hoop netting, minnow traps) 

may be proposed. However, proposed alternative sampling methods must be discussed with the study team 

leader and regional biologist on an individual basis to 

ensure appropriate application and compliance with 

State regulations regarding their use. Alternative sampling methods are described in Bagenal (1978), Nielsen 

and Johnson (1983), Bryan (1984), and Meador and 

others (1993).

6.5 Sample processing procedures

The goal of processing collections of fish in the 

field is to collect information on taxonomic identification, length, weight, abundance, and the presence of 

external anomalies (fig. 11) with minimal harm to 

specimens that will be released alive back into the 

stream. The major steps in processing collections of 

fish from each electrofishing pass and group of seine 

hauls or kicks include the following:

1. Sort fish into identifiable and unidentifiable 

groups. Process threatened and endangered 

(T&E) fish species first before other identifiable 

species. Preserve unidentifiable species and 

return them to the field office for identification.

2. Hold and anesthetize fish in a manner consistent 

with minimizing stress or death.

3. Identify and enumerate each identifiable species.

4. Measure total length and weigh at least 30 specimens of each species (excluding T&E species).

5. Examine up to 30 specimens of each species 

(excluding T&E species) for external anomalies.

6. Record data on the Fish Field Data Sheet.

7. Preserve selected specimens for identification in 

the laboratory or vouchering; release all other 

specimens alive back into the stream.

6.5.1 Handling and anesthetizing fish

All captured fish, excluding T&E species, are 

placed in a live cage or boat live well for subsequent 

processing and an effort is made to minimize stress or 

death to specimens. T&E species are identified, enumerated, and released immediately to the stream. 

Regardless of the effort made to minimize handling and 

stress to fish, some mortality will occur (see Fish Tip 

6). However, minimizing mortality involves recognizing which species are sensitive to handling and prolonged confinement and processing them first. 

Therefore, crewmembers should be thoroughly familiar 

with the fish species in their study area. Anesthetizing 

further minimizes stress to the fish and facilitates handling during processing. Carbon dioxide (CO2) (Summerfelt and Smith, 1990) and clove oil (Peake, 1998) 
Field Protocols 61

Can species be

identified in the

field?

YES NO

Preserve

specimens

Identify and

enumerate all

specimens

At least 30 individuals,

each species

Measure total length

Weigh

Examine for external

anomalies

Record data on

Fish Field Data

Sheet

Deposit in voucher

collection

Preserve

specimens

Voucher

Specimens?

Record data on

Fish Field Data

Sheet

At least 30 individuals,

each species

Measure total length

Weigh

Examine for external

anomalies

Anesthetize,

identify, and

enumerate all

species

Is species

threatened or

endangered?

Identify and

enumerate all

specimens

Record data on

Fish Field Data

Sheet

Return all specimens

to stream

Allow fish to

recover from

anesthetic

YES NO

NO YES

Figure 11. Steps followed in the field to process fish collections in the National Water-Quality Assessment Program.
62 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

have been effectively used to anesthetize fish and there 

are no restrictions on their use. After examination, fish 

are placed in a container of ambient stream water and 

allowed to recover from the anesthetic before being 

released back into the stream. Fish should be released 

downstream of the sampling reach to minimize the 

potential for re-sampling and sample bias.

Anesthetizing fish with CO2:

1. Place fish in a 19-L (5-gal) bucket containing 

about 12 L of stream water. [Note: only a few fish 

should be anesthetized at a time to minimize 

potential mortality resulting from prolonged 

sedation.]

2. Add 350 mL of carbonated water or two CO2- 

producing tablets to the bucket. Determining CO2

dosage in the field can be difficult because by the 

time the fish have responded to the sedation, the 

concentration of CO2 may be too high. If the concentration is too high (onset of sedation is rapid), 

the fish should be moved to native water or processed immediately.

3. Leave fish in the bucket until the desired level of 

sedation is achieved (about 2 to 5 minutes).

Clove oil at a concentration of 60 mg/L produced 

stage four anesthesia (total loss of swimming motion 

and weak opercular movement) after about 4 to 5 minutes for walleye; fish recovered within 10 to 12 minutes 

after they were placed in ambient water (Peake, 1998).

Preparing a 30-L solution of 60 mg/L clove oil 

anesthetic:

1. Add 1.8 mL clove oil to 16.2 mL ethanol [Note: 

this solution can be prepared ahead of time and 

stored in a nonbreakable bottle.]

2. Add the clove oil/ethanol solution to 30 L of 

stream water.

6.5.2 Taxonomic identification and enumeration

Fish are taxonomically identified in the field and 

in the laboratory. Identifications are made by a crewmember who is familiar with the fish species commonly found in the study area. Taxonomic 

nomenclature follows that established by the American 

Fisheries Society's Committee on Names of Fishes 

(Robins and others, 1991). An attempt is made in the 

field to identify all fish to the species level. Uncertain 

identifications require that those specimens be 

vouchered for later identification in the laboratory. 

Consult Walsh and Meador (1998) for additional guidance regarding taxonomic identification. 

6.5.3 Length measurements

Length measurements (fig. 12) are determined 

by using a measuring board consisting of a linear 

metric scale on a flat wooden or plastic base with a stop 

at the zero point. Total length must be measured; standard length is optional. Total length measurements are 

recorded individually for up to 30 individuals of each 

species collected from the reach. These 30 individuals 

are selected from the total number of individuals to represent the range of lengths present in the sample for a 

particular species.

Determining total length:

1. Place fish with the body positioned on its right 

side, the head facing the observer's left, and the 

mouth closed (as in fig. 12).

2. Push the snout of the fish against the measuring 

board stop.

3. Measure total length as the distance from the 

closed mouth to the extreme tip of the caudal fin, 

when the lobes of the caudal fin are squeezed 

together.

4. Record total length to the nearest millimeter on 

the Fish Field Data Sheet.

Standard length measurements are optional and 

should only be taken to address specific study objectives. For example, standard length is important in taxonomic studies because it is unaffected by caudal fin 

anomalies. Standard length is the length from the tip of 

the closed mouth to the posterior end of the fleshy caudal peduncle (fig. 12).

Fish Tip 6—Portable aerated containers

Use multiple, portable aerated containers for 

holding and separating each species. Each container 

has a single species or several easily identifiable species. A single aerator pump can be modified to feed 

multiple containers through individual lines and air 

stones.
Field Protocols 63

TOTAL LENGTH

STANDARD LENGTH

0 100 200

MILLIMETERS

6.5.4 Weight measurements

Weight measurements are obtained by using portable electronic and hanging scales. Weight is determined for each fish that is measured for total length. 

However, for individuals of a species weighing < 1 g 

(for example, Pimephales sp.), a batch weight is taken; 

an average individual weight for a given species can be 

determined by dividing the batch weight by the total 

number of individuals weighed in the batch.

Obtaining weights of individual fish: 

1. Level, calibrate, and tare the scale.

2. Place the fish on the scale.

3. Record the weight of the fish to the nearest gram 

on the Fish Field Data Sheet.

Obtaining batch weights of fish:

1. Level, calibrate, and tare the scale.

2. Place all individuals for a particular species on 

the scale.

3. Record the number of individuals weighed and 

the total weight (= batch weight) to the nearest 

gram on the Fish Field Data Sheet.

6.5.5 External anomalies

External anomalies are defined as externally visible skin or subcutaneous disorders or parasites (Ohio 

Environmental Protection Agency, 1987). External 

anomalies might indicate the presence of sublethal 

environmental stresses, intermittent stresses, behavioral 

stresses, or chemically contaminated substrates. External anomalies include deformities, eroded fins, lesions, 

and tumors collectively referred to as "DELT anomalies" (Sanders and others, 1999). The percentage of 

DELT anomalies has been included as a metric in the 

Index of Biotic Integrity (Ohio Environmental Protection Agency, 1987).

Figure 12. Total and standard length measurements of a fish (from Meador and others, 1993).

Fish Tip 7—Suggestions for weighing fish

(1) Use two or three different-sized weighing 

pans (for example, plastic food storage containers, plastic pails, or plastic bottles) to help 

contain fish and keep the scale clean. These 

pans also reduce the amount of direct handling.

(2) Use a sheet of clear plastic wrap to protect the 

electronic scale while working in bad 

weather.
64 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

All fish that are individually measured and 

weighed also are examined for the presence of DELT 

anomalies (fig. 13). Noting and recording other external anomalies, such as anchor worm (Lernaea spp.), 

blackspot (Uvulifer ambloplitis and Crassiphiala

bluboglossa), and popeye disease are optional. However, because of a lack of consistent inverse relation to 

environmental quality across broad geographic areas, 

these optional external anomalies must not be included 

with the DELT anomaly metric. Accurate identification 

of all external anomalies requires adequate training; 

their presence is denoted with a two-letter code (table 

7). Caution should be exercised not to include injuries 

that might have resulted from the collection methods 

(for example, electrofishing).

Deformities are skeletal anomalies that affect the 

head, spinal vertebrae, and fins. For example, scoliosis 

is a type of spinal curvature (fig. 13a). Eroded fins are 

reductions in fin surface area and hemorrhaging along 

fin rays, which can be caused by chronic disease, parasite infestation or poor water quality (fig. 13b). However, eroded fins also can be caused by other factors, 

such as mechanical erosion during spawning. 

Hatchery-raised fish stocked in streams also may have 

eroded fins as a result of prolonged holding under 

crowded conditions in concrete-lined raceways. 

Lesions are defined as tissue alterations that include 

ulcerated, reddened tissue, open sores, or exposed tissue (fig. 13c).

However, obvious predator-caused injuries (for 

example, lamprey scars) should not be diagnosed as 

lesions. Tumors are defined as circumscribed growths 

of tissue growing independently of the structural development of the fish and serving no physiological function (fig. 13d).

Examining fish for DELT external anomalies:

1. Examine up to 30 specimens of each species for 

the presence of DELT external anomalies. To 

minimize the handling of fish that are to be 

released alive, examine for DELT anomalies 

while the fish are being enumerated or measured 

for length and weight.

2. Document any observed DELT anomalies on the 

Fish Field Data Sheet by recording a checkmark 

under the appropriate DELT code (see table 7) for 

each specimen examined.

3. If a specimen was examined and no DELT anomalies were observed, place a checkmark under the 

"AA" (No anomalies) code for that specimen on 

the Fish Field Data Sheet.

4. Other optional anomalies (see table 7) can be 

documented depending on the objectives of the 

study.

Table 7. Two-letter codes used to denote external anomalies in fish (Ohio Environmental 

Protection Agency, 1987)

[DELT, deformities, eroded fins, lesions, and tumors]

Code Description

DELT 

anomalies

AA No anomalies

DE Deformities

ER Eroded fins

LE Lesions

TU Tumors

Optional

AL Anchor worm

BL Black spot

CL Leeches

FU Fungus

IC Ich

NE Blind in one or both eyes, includes missing and grown-over eyes

PA Other external parasites not previously specified

PE Popeye disease
Field Protocols 65

6.5.6 Preserving fish specimens and vouchering

Fish specimens are preserved in the field and 

returned to the laboratory for identification and deposited in a voucher collection. Some species, such as 

those in the minnow genus Notropis, are similar in 

appearance and cannot be positively identified in the 

field. Even taxonomic experts may have difficulty identifying these similar species because confirmatory 

characteristics are either internal (for example, pharyngeal teeth) or require exact counts of meristic characteristics (for example, the number of lateral line scales 

or anal fin rays). Therefore, specimens that are not positively identified in the field are preserved, labeled, and 

returned to the laboratory for later identification. Some 

identifiable specimens also might be preserved and 

deposited in a voucher collection. Walsh and Meador 

(1998) provide guidance and criteria for the selection 

of specimens for vouchering. A confirmation report is 

requested for all fish that are deposited in a voucher 

collection. The confirmation report contains the same 

information as the sample label and lists the confirmed 

identification of each specimen in the voucher collection. After revising the field data sheet, the data are 

entered into Bio-TDB.

Preserving fish specimens:

1. Euthanize specimens with an overdose of anesthetic (for example, CO2).

2. Place all specimens of a particular species or similar looking unknown species, in a sample bottle. 

Ensure that there is sufficient space in the jar for 

adding a necessary volume of 10-percent buffered formalin to preserve tissues.

3. For fish specimens longer than 100 mm total 

length, make a small incision along the right side 

of the body to allow buffered formalin to penetrate the body cavity.

4. Place a completed sample label inside each bottle. If the species is unidentifiable or uncertain, 

make sure each label contains a unique voucher 

ID code (for example, unknown sp. 1, unknown 

sp. 2, Notropis sp. 1, Notropis sp. 2). [Note: 

doing this will facilitate revision of the field data 

a. Example of deformities (DE) showing a 

normal spine (top) and curved spine (bottom). b. Example of severe eroded 

fins (ER).

c. Example of body surface lesions (LE) (note 

reddened area in front of fins). d. Example of raised tumors (TU) on the body surface.

Figure 13. Deformities, eroded fins, lesions, and tumors noted on fish. (Photographs a, c by M. Meador, 

b by Leetown Science Center, and d by M. Mac).
66 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

sheet after positive identification has been made 

and recorded in the confirmation report.]

5. Add enough 10-percent buffered formalin to fill 

the sample bottle. Wear safety goggles and gloves 

to protect from splashing formalin. Tightly seal 

the bottle lid to ensure that the preservative does 

not leak.

6. Leave fish in the buffered formalin for 2 to 7 days 

to ensure proper preservation of tissues.

7. After fish specimens are returned to the laboratory and properly preserved, they are prepared for 

long-term preservation and storage (see Walsh 

and Meador, 1998).

6.6 Field data sheet

A single, two-page field data sheet (see end of 

this section) is used to record site and sample information associated with fish community sampling. Information that is recorded on each field data sheet is 

organized into sections (table 8) that correspond to data 

entry screens in Bio-TDB. Except for these section 

labels (for example, "Sample Header"), data fields or 

subsections that are shown in bold face are required for 

entry into Bio-TDB. Consult the Bio-TDB User Manual (Ruhl and Scudder, accessed April 26, 2002) for 

additional information on entering field data into the 

data base.

Before leaving the sampling site, the team leader 

should review the field data sheets for completeness 

and accuracy. All missing or incorrect information 

should be completed or corrected. The supply of field 

data sheets (blank and completed) should be stored in 

separate resealable bags for protection from water.

Section This section is used to document

Sample header Where and when a sample was collected

Related sampling activities Other sampling activities

Physical site conditions Physical and chemical conditions at the time 

of sampling

Sampling information Sampling methods and effort, and fish specimen data, such as identification, abundance, 

length, weight, and external anomalies

Table 8. Explanation of field data-sheet sections used to record 

information about fish sampling activities
Field Protocols 67

Page # Cumulative Page # .

Date field data reviewed: / / by (initials)

Date entered into Bio-TDB: / / by (initials)

Bio-TDB Sample ID

Fish Community Sample

Field Data Sheet

SAMPLE HEADER

Station: Date (MM/DD/YYYY):

/ /

Waterbody:

Station description:

Station ID: Reach: Start time: h

Collector (leader): End time: h

Sample and photographic notes:

RELATED SAMPLING ACTIVITIES 

(circle all that apply)

Water chemistry Discharge Habitat survey Tissues

Invertebrate survey Algae survey Bed sediment Other (specify):

PHYSICAL SITE CONDITIONS

Clouds: % Wind (circle): Calm Light Moderate Gusty

Precipitation (circle): None Rain Sleet Snow Precipitation Intensity (circle): N/A Light Moderate Heavy

Other weather:

Beginning water measurements Ending water measurements

Time: Time:

Water temperature

(°C):

Water temperature

(°C):

pH: pH:

Dissolved oxygen

(mg/L):

Dissolved oxygen

(mg/L):

Specific conductance

[(μS/cm) @ 25 °C]:

Specific conductance

[(μS/cm) @ 25 °C]:

Turbidity

(NTUs)

Turbidity

(NTUs)

Discharge (ft3/s): Stream stage: ft @ time h

Riparian shading (circle): Shaded Partial Full sun

Water clarity (circle): Clear Slightly turbid Turbid Very turbid

Water color (circle): Black Brown Clear Dark green Light green Yellow

Physical site condition comments:
68 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

SAMPLING INFORMATION

Primary sample or Repeated sampling replicate? (circle one) Sample Number:

Field SMCOD: Method code:

Method codes: Electrofishing—(11A) backpack 1st pass; (11B backpack 2nd pass; (12A) towed 1st pass; (12B) towed 2nd pass; (13A) boat 1st pass; (13B) boat 

2nd pass; Seining—(21A) haul; (22A) kick; (23A) beach.

Model: Output voltage: Seconds:

Beginning time: Ending time: Reach length: Number of seine hauls:

Comments about fish:

Fish Specimen Data

TL=total length; SL=standard length; AA=no anomalies; DE=deformities; ER=eroded fins; LE=lesions; TU=tumors

Name Abundance

ID

Field or 

Lab?

TL

(mm)

SL

(mm)

Wt

(g)

DELT Anomalies 

(check if present) Voucher?

(Y or N) AA DE ER LE TU

Page # Cumulative Page # .
Field Protocols 69

7. POST-FIELD ACTIVITIES

After returning from the field, sampling teams 

undertake several activities prior to shipping samples to 

a laboratory for analysis. These activities include logging site and sample information into Bio-TDB, replacing field labels, preparing packing lists and Analytical 

Services Request (ASR) forms, and preparing samples 

for shipping.

7.1 Logging samples into the Biological Transactional Data 

Base

Sampling teams are required to enter site and 

sample information into Bio-TDB as soon as possible 

after returning from the field. This process will allow 

sampling teams to produce automated sample labels, 

packing lists, and ASR forms in preparation for shipping samples to analytical laboratories. Sampling 

teams are strongly encouraged to demonstrate care 

when entering site and sample information into 

Bio-TDB. Members of a sampling team should practice 

entering samples into the Bio-TDB demonstration data 

base so they can be familiar with the different data 

fields and requirements for each sample medium. 

When a sample is logged into the data base, a Bio-TDB 

sample identification number (different from the 

SMCOD sample number) is assigned to it and then 

recorded on the corresponding field data sheet sample 

header.

7.2 Replacing field sample labels

Labeling samples with complete and accurate 

collection data is an essential step to ensure proper 

sample accounting and laboratory analysis. Careful 

attention to labeling samples accurately also eliminates 

discrepancies that sometimes occur when several different kinds of samples are collected in a study. In the 

field, each bottle or filter for a particular sample is 

given an external label (fig. 2) that uniquely identifies 

its contents. The information on field labels is sometimes difficult to read (especially by laboratory analysts) if it is handwritten, or even lost because improper 

writing ink was used. Therefore, sampling teams must 

replace all field labels with labels produced by the 

automated Bio-TDB label function. Bio-TDB sample 

labels can only be produced if site and sample information has first been entered into the data base. The automated Bio-TDB labels are required because all of the 

information on the label is clearly printed and based on 

information entered into the data base, thereby minimizing discrepancies. Consult the Bio-TDB User Manual (Ruhl and Scudder, accessed April 26, 2002) for 

instructions on preparing automated labels.

7.3 Preparing packing lists and Analytical Services 

Request Forms

Packing lists are an accounting of the samples 

contained in each package and provide a quick means 

of determining the accuracy and completeness of a 

shipment. ASR forms convey important information 

(for example, NWQL labcodes or schedules) about the 

sample and how it is to be processed by the analytical 

laboratory. Laboratories will not log samples into their 

internal tracking systems or begin to process samples 

unless complete ASR forms are received for each sample. Consult the Bio-TDB User Manual for instructions 

on producing automated packing lists and ASR forms.

7.4 Shipping samples to analytical laboratories

Shipping algal and invertebrate samples to analytical laboratories requires careful consideration and 

planning. Several safety and transportation regulations 

relating to how samples are packed, labeled, and 

shipped must be observed by law. Each crewmember 

who is responsible for shipping samples must be 

trained in shipping hazardous materials (for example, 

formalin). This person should obtain the most current 

U.S. Department of Transportation (DOT; see 

http://hazmat.dot.gov, accessed April 26, 2002) and 

transportation agent (for example, Airborne or Federal 

Express) requirements on packing and shipping samples. Failure to comply with these requirements might 

result in the shipment being returned to the customer or 

fines and penalties levied by the DOT or transportation 

agent. Consult USGS NWQL Rapi Note Nos. 01-033 

(2001b) and 01-034 (2001c) for additional information 

regarding safe shipping regulations.

Packing and shipping samples:

1. If necessary, add sample preservative to completely fill each sample bottle to minimize void 

space. [Note: void space increases agitation of 

sample debris that might damage delicate organisms such as mayflies during shipping.]

2. Clean sample bottle threads to ensure that bottle 

lids fit securely. Sand or other debris that is 
70 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

trapped in the threads might cause sample leakage.

3. Stretch a strip of high-quality, plastic tape (for 

example, 3M ScotchTM Brand 471) around the 

lid-bottle seal to prevent or minimize leakage. 

Tape is wrapped in the direction of closing the 

lid. [Note: clear plastic or fiber box tape, masking tape, duct tape, and electricians tape are not 

suitable tapes for sealing sample bottle lids and 

often result in considerable leakage.] Leaking 

sample preservatives represent a safety hazard to 

transportation workers and to laboratory personnel responsible for receiving and processing samples.

4. Protect the Bio-TDB generated sample bottle 

labels during shipping by covering with clear 

packing tape.

5. Place each sample bottle in a sturdy (for example, 

freezer quality) resealable plastic bag and expel 

the air. Doing this provides secondary 

containment of each sample and prevents 

cross-contamination of samples if bottles break 

during shipping and handling.

6. Prepare the exterior of a shipping container (for 

example, cooler) by affixing orientation arrows 

and all required safety and hazard information 

and labels.

7. Line the inside of the container with a layer of 

bubble packing or other suitable shock-absorbing 

material.

8. Line the container with two nested heavy-duty 

trash bags.

9. Place a layer of absorbent material (for example, 

multi-ply paper padding) in the bottom of the 

inner trash bag. Do not use styrofoam peanuts, 

shredded paper, or vermiculite.

10. Tightly pack sample bottles against each other 

inside the trash bag. Ensure that bottles and 

resealable plastic bags are upright. Highlight 

samples on the packing list as they are placed in 

the container. [Note: do not exceed volume limits 

specified for shipping hazardous materials.]

11. Place additional absorbent material around bottles as necessary to fill gaps and close both trash 

bags with a tie.

12. Place a layer of bubble packing on top of the trash 

bags along with internal shipping documents. 

Place shipping documents (safety information, 

packing lists, and ASR forms) for each container 

in a resealable plastic bag and put the bag on top 

of the packed samples. Safety documents include 

24-hour emergency contact information for the 

receiving laboratory and a copy of all applicable 

documentation for shipping hazardous material 

(for example, buffered formalin).

13. Close the shipping container with heavy-duty 

tape.

14. Complete all required hazardous materials shipping paperwork.

SUMMARY

Protocols for sampling algal, invertebrate, and 

fish communities as part of the National Water-Quality 

Assessment (NAWQA) Program were revised based on 

several methodological and procedural modifications, 

additions, and deletions that resulted during the first 10 

years of the Program. The coordinated sampling of 

aquatic biological communities, water chemistry, and 

physical attributes in the NAWQA Program provides 

an integrated approach to monitoring water quality. 

This integrated approach is implemented in a common 

spatial setting by sampling a defined length of stream 

called a "reach." The methods described herein were 

originally developed to address objectives of the 

NAWQA Program, but may be adapted for other 

water-quality investigations.

Several pre-field activities are performed prior to 

going into the field to sample. These activities include 

obtaining necessary collecting permits, organizing supplies and equipment, preparing sample preservatives, 

checking the condition of and repairing sampling gear, 

preparing a field activities plan, and becoming familiar 

with functions of the Biological Transactional Data 

Base. The safety of the sampling teams in the field is a 

high priority. Therefore, all crewmembers must wear 

personal flotation devices when required by a Job Hazard Analysis and be instructed on the safe handling and 

operation of sampling gear, vehicles, and boats used to 
Summary 71

collect samples. Special training and certification is 

obtained for certain operations (for example, operating 

electrofishing gear or a boat).

Algal sampling protocols for periphyton and 

phytoplankton describe methods for collecting composited quantitative and qualitative samples. Up to four 

different sample types might be collected to characterize the algal community in the sampling reach. Three 

quantitative sample types include a richest-targeted 

habitat periphyton sample, a depositional-targeted habitat periphyton sample, and a phytoplankton sample. 

The quantitative samples are collected to measure algal 

community structure, chlorophyll a, and ash-free 

dry-mass. Instead of ash-free dry-mass, phytoplankton 

samples are analyzed for particulate organic carbon. A 

qualitative multihabitat periphyton sample can be collected for characterizing algal taxa richness throughout 

the sampling reach. Methods for collecting these sample types depend on wadeability at the site and on the 

type of algal habitat present in the reach (for example, 

epilithic, epidendric, or epipsammic). For example, the 

SG-92 sampler is recommended for use when collecting a quantitative microalgal sample from epilithic substrates. Samples are processed in the field to prepare 

samples for laboratory analysis. Algal sample processing includes preserving sampling for taxonomic identification and homogenizing and filtering samples for 

chlorophyll a and ash-free dry-mass or particulate 

organic carbon analysis. Field data sheets are used to 

record sampling information (for example, where and 

how samples are collected, and weather and site conditions) and ancillary data from measurements necessary 

for the interpretation of the biological data (for example, light availability and current velocity).

Invertebrate sampling protocols describe methods for collecting a composited semi-quantitative and 

qualitative sample to characterize the invertebrate community in the sampling reach. The semi-quantitative 

invertebrate sample is collected in the richest-targeted 

habitat (typically a riffle or woody snag) that theoretically represents where the highest invertebrate taxa 

richness is found. This sample is collected to describe 

the community structure (for example, relative abundance, and taxonomic and functional metrics) of the 

invertebrate taxa found in the richest-targeted habitat. 

Similar to the algal protocols, a qualitative multihabitat 

invertebrate sample can be collected for characterizing 

invertebrate taxa richness throughout the reach. Methods for collecting these samples depend on wadeability 

at the site and on the substrate particle size (for example, coarse or fine-grained substrates). For example, the 

Slack sampler is used when collecting a semiquantitative sample from wadeable, coarse-grained riffle habitats. Samples are processed in the field prior to 

preparing them for analysis at the laboratory. Invertebrate sample processing includes washing and sieving 

sample debris to remove fine particles, elutriating to 

separate organic and inorganic materials, and preserving invertebrates for identification. Sample splitting 

procedures also are used to reduce the sample to an 

appropriate volume (for example, < 750 mL) for effective performance of laboratory processing methods. 

Field data sheets are used to record sampling information (for example, where and how sample are collected, 

and weather and site conditions) and ancillary data 

from measurements necessary for the interpretation of 

the biological data (for example, current velocity, sample depth, and woody snag sampling areas).

Fish sampling protocols describe methods for 

collecting a representative sample of the fish community from the entire sampling reach. Two complementary sampling methods are used at each site. The 

primary sampling method is electrofishing; the complementary secondary method is seining. The techniques 

for applying these methods depend on wadeability at 

the site and the complexity of instream habitats present 

in the geomorphic channel units (for example, riffles, 

runs, pools) represented. For example, backpack or 

towed electrofishing gear and techniques are used in 

wadeable streams, whereas boat electrofishing gear is 

used in nonwadeable streams. Fish are processed in the 

field and released alive back into the stream whenever 

possible. Fish processing includes sorting, identifying, 

and counting each species, measuring total length, 

weighing, and examining for external anomalies (for 

example, deformities, eroded fins, lesions, and tumors). 

Data from these processing steps are recorded on field 

data sheets along with sampling information. Fish are 

anesthetized to minimize stress during processing and 

to ensure their safe return to the stream. Fish specimens 

are preserved when identification in the field is uncertain or not possible, or when specimens are needed for 

vouchering.

After returning from the field, information 

recorded on field data sheets is entered into the Biological Transactional Data Base from which formatted 

sample labels, packing lists, and Analytical Services 

Request forms are generated. Samples, which are contained in standardized bottles, are packed in approved 

containers to minimize damage during shipping to the 
72 Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program

laboratory for analysis. These packing procedures also 

ensure that samples are not cross-contaminated and 

that shipping and laboratory personnel and the public 

are not exposed to leaking chemical preservatives.
References Cited 73

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Appendix 75

APPENDIX

Operational modifications for the Slack sampler

(a) Bipod Stand

Materials:

• 2–6-ft wooded tomato stakes

• 1–3/8-in x 3-in-long hex bolt and 3/8-in hex nut

• 3–5/16-in cut washers

• 1–5/16-in lock washer

• 1–6/32-in round head machine screw with fitted 

nut and lock washer

• 1–standard No. 20 screw clamp (hose clamp)

• 1–1-in x 6-in-long (1/16-in thick) strip of galvanized steel

Constructing the bipod stand:

1. Drill one end of the 

strip of galvanized 

steel to the diameter 

of the hex bolt and 

the other end to the 

diameter of the 

small machine 

screw.

2. Drill each wooden 

tomato stake 28 cm 

from the top with a 

bit slightly larger 

than the hex bolt 

diameter.

3. Drill the center of the clamp strap to fit the 

machine screw.

4. Bend the strip of steel as indicated in the photograph.

5. Place one washer on the hex bolt, which is then 

passed through the end of steel strip with the 

larger diameter hole and through one of the 

wooden stakes.

6. Add a second washer to the hex bolt and then 

attach the other wooden stake.

7. Add a third washer, followed by the lock washer 

to the end of the hex bolt and secure with the hex 

nut.

8. Attach the screw clamp to the top end of the 

S-shaped steel strip with a machine screw.

9. The assembled bipod should support the upright 

handle of the Slack sampler (see photo and 

fig. 8a). The bipod is implemented by placing the 

Slack sampler in the stream, then sliding the 

opening of the screw clamp down over the top of 

the handle to about 1/3 the distance from the top 

of the handle. The wooden stakes are angled to 

support the Slack sampler. If needed, the screw 

clamp can be tightened to prevent the bipod apparatus from sliding.

(b) Area Template

Materials:

• 1–5-ft length of 1⁄2-in PVC pipe

• 2–PVC T connectors

• some shock cord or 1⁄4-in braided nylon rope

• 2–spring clips

Constructing the area template:

1. Cut the PVC piping into three equal segments.

2. Connect the PVC pipe with the two T-connectors 

to form two right angles. [Note: using T-connectors (instead of elbow connectors) and drilling a 

few holes into the PVC pipe will allow water to 

flow in the frame, preventing it from floating up 

during sampling.]

3. String the shock cord or braided nylon rope 

through all three segments of PVC piping. Keep 

some slack so that the apparatus can be collapsed 

and then tightened and clipped (tied snug if using 

rope) to the frame when in use.

Close-up of bipod attached to Slack 

sampler handle