United States
Environmental Protection
Agency
Environmental Monitorii
Systems Laboratory
26 West Martin L King D,
Cincinnati, OH 45268
Office of Research and Development
Aorll 1995
ENVIRONMENTAL MONITORING
AND ASSESSMENT PROGRAM -
SURFACE WATERS: v
HELD OPERATIONS AND
METHODS FOR MEASURING THE
ECOLOGICAL CONDITIONS OF
WADEABLE STREAMS
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EPA/620/R-94/004
April 1995
ENVIRONMENTAL MONITORING AND ASSESSMENT PROGRAM
SURFACE WATERS:
FIELD OPERATIONS AND METHODS FOR MEASURING THE ECOLOGICAL CONDITION
OF WADEABLE STREAMS
Edited by
Donald J. Klemm and James M. Lazorchak
Bioassessmenl and Ecotoxicology Branch
Ecological Monitoring Research Division
Environmental Monitoring Systems Laboratory, Cincinnati, Ohio
Cincinnati, OH 45244
TAI Contract No. 68-C1-0022
Work Assignment 0-06, James M. Lazorchak, Manager
ENVIRONMENTAL MONITORING SYSTEMS LABORATORY
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
CINCINNATI. OHIO 45268
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DISCLAIMER
This document has been reviewed by the Environmental Monitoring Systems Laboratory -
Cincinnati (EMSL-Cincinnati), U.S. Environmental Protection Agency (USEPA), and approved for
publication. The mention of trade names or commercial products does not constitute endorsement or
recommendation for use.
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FOREWORD
Environmental measurements .are required to determine the quality of ambient waters and the
character of waste effluents. The Environmental Monitoring Systems Laboratory - Cincinnati (EMSL-
Cincinnati) conducts research to:
o Develop and evaluate methods to identify and measure the concentration of chemical
pollutants In drinking waters, surface waters, groundwaters, wastewaters, sediments, sludges, and
solid wastes.
o Investigate and evaluate methods for the Identification and measurement of viruses, bacteria
and other microbiological organisms In aqueous samples and to determine the response of
aquatic organisms to water quality.
o Perform ecological assessments and measure the toxiclty of pollutants to representative
species of aquatic organisms and determine the effects of pollution on communities of indigenous
freshwater, estuarine, and marine organisms, including the phytoplankton, zooplankton,
periphyton, macrophyton, macroinvertebrates, and fish.
o Develop and operate a quality assurance program to support the achievement of data quality
objectives in measurements of pollutants in drinking water, surface water, groundwater,
wastewater, sediment and solid waste.
o Develop methods and models to detect and quantify responses in aquatic and terrestrial
organisms exposed to environmental stressors and to correlate the exposure with effects on
biochemical and biological Indicators.
This field operations and methods manual describes guidelines and standardized procedures for
evaluating the biological integrity of surface waters of streams. It was developed to provide the
Environmental Monitoring and Assessment Program (EMAP) with bioassessment methods for
determining the status and monitoring trends of the environmental condition of freshwater streams.
These bioassessment studies are carried out to assess biological criteria for the recognized beneficial
uses of water, to monitor surface water quality, and to evaluate the health of the aquatic environment.
Thomas A. Clark
Director
Environmental Monitoring Systems
Laboratory • Cincinnati
in
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PREFACE
The Bioassessment and Ecotoxicology Branch, Ecological Monitoring Research Division.
Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency • Cincinnati is
responsible for field and laboratory methods that are used in assessing aquatic ecosystems. Research
areas include the development, evaluation, validation, and standardization of agency methods for the
collection of biological field and laboratory data by USEPA regional, enforcement, and research
programs engaged in inland, estuarine, and marine water quality and permit compliance monitoring, and
status and/or trends monitoring for the effects of impacts on aquatic organisms, including the
phytoplankton, zooplankton, periphyton, macrophyton, macroinvertebrates, and fish. The program
addresses methods and techniques for sample collection; sample preparation; processing of structural
and functional measures by using organism identification and enumeration; the measurement of biomass
and benthic metabolism; the bioaccumulation and pathology of toxic substances; acute, chronic, and
sediment toxiclty. the computerization, analysis, and interpretation of biological data; and ecological
assessments. The Bioassessment and Ecotoxicology Branch also Includes field and laboratory support
of the ecological biomarker research program and transfer of monitoring technology to the regions and
state programs.
This document contains the EMAP-SW field operations and bioassessment methods for
evaluating the health and biological integrity of wadable freshwater streams.
iv
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ABSTRACT
The Environmental Monitoring and Assessment Program (EMAP) is being implemented to provide
for the assessment of status of and trends in the condition of ecological resources within the United States.
In 1993 the Surface Waters Resource Group within EMAP implemented the pilot programs for streams.
This document contains the updated version of the field operations and methods for biological response
indicators (periphyton, benthic sediment metabolism, benthic invertebrates, fish, fish tissue collection,
sediment toxicity sample collection), site protocols, water chemistry, physical assessment methods,
information management, safety and health, and habitat and stream assessment forms that will be used for
the U.S. Environmental Protection Agency (USEPA), Environmental Monitoring and Assessment Program
(EMAP), the USEPA Regional Environmental Monitoring and Assessment Program (REMAP), and the field
procedures for collection chemical indicators of acidification for the EMAP Temporally Integrated
Monitoring of Ecosystems (TIME). This manual was prepared to be user friendly for field personnel that are
responsible for collecting biological, chemical, and physical indicator samples.
v
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EMAP-SURFACE WATERS-STREAMS STUDY
AUTHORS AND CONTRIBUTORS
Section 1: James M. Lazorchak', H. Ronald Preston*, Alan T.Hertiny*. and Donald J. Klemm'
Section 2: Alan T. Herlihy 3 and James H. Green *
Sections: Brian Hill*
Section* Alan T. Herlihy3
Sections: Alan T. Herlihy3
Section 6: Philip R. Kaufmann 3 and E. George Robison'
Section?: Brian H. Hill1
Sections: Brian H. Hill1
Section 9: Philip A. Lewis' and Donald J. Klemm'
Section 10: Frank H. McCormick1
Section 11: Roger B. Yeardley Jr.*
Section 12: Mark E. Smith-
Section 13: Donald J. Klemm', Philip A. Lewis', and James M. Lazorchak'
Section 14: Victoria Rogers3
Section 15: Alan T. Herlihy3 and James H. Green*
'U.S. Environmental Protection Agency, Environmental Monitoring Systems Laboratory, 3411 Church
Street, Cincinnati, OH 45244.
•U.S. Environmental Protection Agency, Wheeling Office, 303 Methodist Building, 11 th and Chaplin
Streets, Wheeling. WV 26003.
Oregon State University, c/o U.S. Environmental Protection Agency, Environmental Research
Laboratory, 200 SW 35th Street Corvallis, OR 97333.
'DynCorp, c/o. U.S. Environmental Protection Agency, Environmental Monitoring Systems Laboratory,
3411 Church Street, Cincinnati. OH 45244.
9 Ogden Governmental Services, c/o U.S. Environmental Protection Agency, Environmental Research
Laboratory. 200 SW 35th Street. Corvallis. OR 97333.
vl
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CONTENTS
Foreword Hi
Preface iv
Abstract v
Authors And Contributors vi
Acknowledgments x
SECTION PAGES
I. Introduction 1 of 5
2. Field Operation Overview 1 of 2
3. Summary Of Daily Field Operations For EMAP Stream Sampling 1 of 2
3.2 Geomorphs 1 of 2
3.3 Biomorphs and Geomorphs 1 of 2
3.4 In Summary: Geomorphs And Biomorphs 2 of 2
4. Initial Site Protocols 1 of 5
4.1 Initial Site Survey . 1 of 5
4.2 Laying Out The Stream Sample Reach 3 of 5
4.3 Procedures For Sampling Dry And Intermittent Streams 4 of 5
5. Water Chemistry 1 of 5
5.1 Introduction 1 of 5
5.2 Collecting Water Chemistry Samples 1 of 5
5.3 Stream Side Chemical Measurements 3 of 5
5.4 References 4 of 5
6. Physical Habitat Assessment 1 of 38
6.1 Introduction 1 of 38
6.2 Thalweg Profile 5 of 38
6.3 Large Woody Debris Measurements 10 of 38
6.4 Channel and Riparian Cross-Sections 12 of 38
6.5 Discharge Measurement 29 of 38
6.6 Equipment List 33 of 38
6.7 References 35 of 38
6.8 Field Forms For Physical habitat Protocols 36 of 38
7. Periphyton Indicator: Field Procedures 1 of 3
7.1 Sampling Rationale 1 of 3
7.2 Field Equipment 1 of 3
7.3 Field Protocols 2 of 3
7.4 Ship Samples To 3 of 3
vi!
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CONTENTS (CONTINUED)
SECTION PAGES
8. Benthic (Sediment) Metabolism: Field Methods 1 of 4
1. Scope and Application lof4
2. Summary of Method 1 of 4
3. Definitions 1 of 4
4. Interferences 1 of 4
5. Safety 1 of 4
6. Equipment and Supplies 1 of 4
7. Reagents and Standards 2 of 4
8. Sample Collection, Preservation, and Storage 2 of 4
9. Quality Control 2 of 4
10. Calibration and Standardization 3 of 4
11. Procedures 3 of 4
12. Data Analysis and Calibration 4 of 4
13. Method Performance 4 of 4
14. Pollution Prevention 4 of 4
15. Waste Management 4 of 4
16. References 4 of 4
17. Tables, Diagrams, Flowcharts, and Validation Data 4 of 4
9. Benthic Invertebrate Indicator 1 of 9
9.1 Introduction 1 of 9
9.2 Procedures for Field Crews 2 of 9
9.3 References 7 of 9
10. Fish Indicator 1 of 14
10.1 Experimental Design 1 of 14
10.2 Methods 1 of 14
10.3 Safety 8 of 14
10.4 Quality Assurance/Quality Control 8oM4
10.5 References 10 of 14
11. Fish Tissue Contaminants Indicator 1 of 6
11.1 Introduction 1 of 6
11.2 Selecting Rsh Tissue Specimens 2 of 6
11.3 Processing Tissue Specimens 3 of .6
11.4 Shipping Tissue Specimens 5 of 6
11.5 Target Analytes 4 of 6
12. Sample Collection and Shipment for Sediment Toxicity Samples 1 of 1
12.1 Sediment collection and Shipment 1 of 1
12.2 Sediment Testing Methods 1 of 1
viii
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CONTENTS (CONTINUED)
SECTION PAGES
13. Safety and Health 1 of 5
13.1 Introduction 1 of 5
13.2 General Precautions 3 of 5
13.3 Safety Equipment and Facilities 3 of 5
13.4 Field and Laboratory Operations 3 of 5
13.5 Disease Prevention 4 of 5
13.6 Chemical Wastes 4 of 5
13.7 References 4 of 5
14. Information Management 1 of 26
14.1 Introduction 1 of 26
14.2 Field Data Forms 1 of 26
14.3 Field Sample Processing 2 of 26
14.4 Sample Labels and Preparation 2 of 26
14.5 Sample Tracking and Reporting 2 of 26
14.6 Supplies 4 of 26
14.6 Forms, Labels, and Checklists 4 of 26
15. Rapid Habitat and Stream Assessment Forms 1 of 5
15.1 Rapid Physical Habitat Assessment Form 1 of 5
.15.2 Stream Assessment Form 4 of 5
Appendices
Summary Of Field Protocols for Indicators
A. Summary of Initial Site Protocols 1 of 1
B. Water Chemistry 1 of 2
C. Physical Habitat 1 of 7
D. Summary Field Protocols for Periphyton 1 of 2
E. Summary Field Protocols for Benthic (Sediment) Metabolism 1 of 2
F. Benthic Invertebrates 1 of 3
G. Field Protocols for Fish Collection 1 of 3
H. Selecting Fish Tissue Specimens 1 of 2
I.. Sample Collection and Shipment for Sediment Toxicrty Samples 1 of 1
ix
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ACKNOWLEDGMENTS
Review comments from the following persons are gratefully acknowledged: Deb J. Chaloud,
Environmental Monitoring Systems Laboratory, Las Vegas, NV 89193; Alan T. Heriihy, Oregon State
University, c/o. U.S. Environmental Protection Agency, Environmental Research Laboratory, Corvallis.
OR 97333; Robert M. Hughes, ManTech Environmental Technology, Inc., c/o U.S. Environmental
Protection Agency, Environmental Research Laboratory, Corvallis. OR 97333; Philip R. Kaufmann,
Oregon State University, c/o U.S. Environmental Protection Agency, Environmental Research Laboratory.
Corvallis, OR 97333; Donald J. Wemm, U.S. Environmental Protection Agency, Environmental Monitoring
Systems Laboratory, Cincinnati OH 45244; Philip A. Lewis, U.S. Environmental Protection Agency,
Environmental Monitoring Systems Laboratory, Cincinnati, OH 45244; Peter M. Nolan, U.S.
Environmental Protection Agency, Region 1, ESD, Biology Section, Lexington, MA 02173; David V: Peck,
U.S. Environmental Protection Agency, Environmental Monitoring Systems Laboratory, Las Vegas, NV
89193; H. Ronald Preston, U.S. Environmental Protection Agency, Wheeling Office, Wheeling VW 26003;
Richard D. Spear, U.S. Environmental Protection Agency, Regional Office II, Edison, NJ 08837.
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EMAP-SW-Streams, Introduction, Field Methods, Section 1. Revision No. 2, March 1994, Page 1 of 5
SECTION 1
INTRODUCTION
1.1 This manual contains an update of the 1993 pBot field operations and procedures for biological
response indicators, physical assessment methods, and research protocols that wOl be used In the 1994
pilot programs for the U.S. Environmental Protection Agency (USEPA), Environmental Monitoring and
Assessment Program (EMAP) and the USEPA Region III Regional Environmental Monitoring and
Assessment Program (REMAP). It also contains the field procedures for collection of chemical indicators
of acidification for the EMAP Temporally Integrated Monitoring of Ecosystems (TIME). The purpose of
the manual is to be a users manual for field teams conducting sampling and snipping of biological
samples. The 1993 pilot laboratory methods have also been updated and are now contained in a
separate document, "1994 Pflot Laboratory Methods Manual For Streams.*
1.2 The EMAP Surface Water Program wDI start its second season of stream pBot activities in the mid-
Appalachian region during 1994 (see Section 2, Field Operations Overview). EMAP Research wDI be
conducted in conjunction with USEPA's TIME and Region 3 REMAP programs. Three hundred eight
Appalachian Ridge and Valley, Blue Ridge, Central Appalachian Plateau, Piedmont and Atlantic Coastal
Plain sites have been selected for this study. The streams wQI be sampled during the 10 week index
period from AprO to July. These sites wDI be sampled by crews of USEPA, U.S Fish and Wildlife Service,
state and contract personnel In each of the operational units of the EMAP, REMAP, and TIME projects.
1.3 The USEPA has designed the EMAP program to determine the current status, extent, changes and
trends in the condition of our nation's ecological resources on regional and national scales. The nation's
ecological resources are a national heritage, as essential to the country now and In the future as they
have been in the past Data indicate that regional and international environmental problems may be
endangering these essential resources. The potential threats include acid rain, ozone depletion.
nonpoint sources of pollution, and climate change.
1.4 The primary goal of EMAP (Whlttier and Pauisen, 1992) Is to provide environmental decision makers
with statistically valid interpretive reports describing the health of our nation's ecosystems. Knowledge
of the health of our ecosystems wfll give decision makers and resource managers the ability to make
informed decisions, set rational priorities, and make known to the public costs, benefits, and risks of
proceeding or refraining from implementing specific environmental regulatory actions. EMAP's
ecological status and trends data wfll allow decision makers to assess objectively, whether or not the
nation's ecological resources are responding positively, negatively, or not at all, to the regulatory
programs put in place ostensibly to benefit them.
1.5 To accomplish ifs goals EMAP is to document the condition of the nation's forest, wetlands,
estuarine and coastal waters (USEPA, 1991 a), Inland surface waters, Great Lakes, agricultural lands, and
arid lands in an integrated manner, on a continuing basis. Although EMAP is designed and funded by
USEPA's Office of Research and Development (ORD), other offices and regions within EPA and other
federal agencies have contributed to its development and wDI participate in the collection and use of
EMAP data. When fully implemented, EMAP will form a comprehensive national monitoring network,
with a large proportion of the data collection and analysis being accomplished by federal, state and local
agencies.
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EMAP-SW-Streams, Introduction, Reid Methods, Section 1. Revision No. 2, March 1994, Page 2 of 5
1.6 The following three objectives guide the EMAP program:
Estimate the current status, extent, changes and trends in indicator* of the
condition of the nation's ecological resources on a regional basis with known
confidence.
Monitor indicators of pollutant exposure and habitat condition and seek associations
between human-Induced stresses and ecological condition.
Provide periodic statistical summaries and interpretive reports on ecological status
and trends to resource managers and tht public.
1.7 The EMAP inland surface waters program (EMAP-SW) Is that part of the overall program which is
monitoring the health of lakes and streams in the United States. The first phase of the program started
with lakes in 1991 (USEPA, 1991b. 1991C, and 1991d), continued in 1992 (USEPA, 19923, 1992b. 1992C.
and 1993), and win continue In 1993, and 1994 (USEPA, 1994). The second phase of the EMAP-SW
program has concentrated on streams in 1993 and wfll again in 1994.
1.8 The EMAP-SW streams pflot of 1993 and 1994 is designed to focus on three classes of streams:
first, second, and third order streams. The 1994 EMAP-SW streams pilot has also been Integrated with
two other USEPA programs, TIME and the USEPA Region ill REMAP pBot
1.9 Prior to fun-scale implementation of EMAP-SW streams, a number of questions must be answered
through a combination of analyses of existing data and data derived from new field activities. There are
two types of field activities to be performed prior to fun-scale implementation, pDot projects and
demonstration projects. PBot projects are intended to answer questions about proposed indicators (e.g.
plot design, indicator sensitivity to various stressors, magnitude of variance components, alternative
methods evaluations, and logistical constraints). PDot studies are not primarily intended to provide
regional estimates of condition but may provide these estimates for a few indicators. Demonstration
projects may be designed to answer many of the same questions but also have the prime objective to
demonstrate the ability to estimate the condition of regional populations.
1.10 In 1993, the stream pBot initiated the USEPA's EMAP, TIME, and REMAP status and trend
monitoring of the ecological condition of the stream population In USEPA Region III. In 1994 this
integrated pilot wO continue to estimate the condition of the entire stream population in a probability
design. For the Region 111 pDot, we have selected stream sample sites with a randomized, systematic
design using the blue lines on 1:100,000 scale USQS topographic maps as the basis for our stream
populatioa For purposes of this pBot, we have restricted our sample population to wadeable streams
(first, second, and third order on the 1:100,000 maps) in the Appalachian Mountain portion of Region III
(north/west of the Blue Ridge, the Piedmont and Atlantic Coastal Plain). The Region III stream pilot
consists of 308 total stream sites visited, 62 EMAP probabBity stream sites, 30 hand-picked reference
sites, and 21 repeat visit sites. One hundred-eleven sites are associated with base EMAP sampling In
the region, 195 sites are associated with the EMAP/HME project (designed to monitor changes in
surface waters associated with changes in acidic deposition), and 84 sites are associated with Region
Ill's REMAP project
1.11 We intend to monitor the physical, chemical, and biological components of ecological condition In
single one-day visits to stream reaches. We realize that assemblages and their physical and chemical
habitat change seasonally and annually at a single site. However, our data indicate that far greater
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EMAP-SW-Streams, Introduction, Field Methods, Section 1, Revision No. 2, March 1994, Page 3 of 5
differences exist between stream sites during similar time periods. Therefore, we believe we can acquire
more information and better assess this spatial variability by maximizing the number of reaches sampled
and by sampling within a specific 'index* period. Our chosen index period Is late spring, specifically late
April through July, twenty-one of the 62 base EMAP sites will be resampled during the index period to
evaluate index period variability.
1.12 A few of the questions to be answered in the 1993/1994 EMAP-SW streams project are as follows:
What Is the plot design for each class of streams for each indicator?
How will each indicator be used to describe condition (good-poor) of streams and how
will acceptable (good) and unacceptable (poor) be determined?
What are differences among different classes of streams?
How many discrete habitats need to be sampled in each plot?
Can teams conduct the field sampling in the time frame described (Late April
through mid July)?
What is the magnitude of regional trends detectable given the indicator
variability and proposed design?
1.13 A special interest component of EMAP-SW is the TIME project The purpose of the TIME project is
to assess the changes and trends in chemical condition in acid-sensitive surface waters of the U.S.
resulting from changes in acidic deposition caused by the 1990 Clean Air Act Amendments. The EMAP-
TIME project has three goals (Stoddard, 1990):
Monitor current status and trends in chemical indicators of acidification In acid-
sensitive regions of the U.S.
Relate changes in deposition to changes in surface water conditions.
Assess the effectiveness of the Clean Air Act emissions reductions in improving the
acid/base status of surface waters.
1.14 The Region III REMAP project was designed to test the EMAP approach on a few of the most
heavily impacted ecoregions of Region III, the mid-Appalachians, the Ridge and Valley, the Central
Appalachians, the Piedment and some of the coastal plain.
1.15 The Region III REMAP project is designed to answer the following questions:
What are biological reference conditions for the Central Appalachian Ridge and Valley
Ecoregion?, .
Do biological communities differ between subregions?
What is the status of mid-Atlantic Highlands stream biota?
Can linkages be established between impairment and possible causes of impairment?
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EMAP-SW-Streams, Introduction, Reid Methods, Section 1, Revision No. 2. March 1994, Page 4 of 5
How can an EMAP-IIke approach be used to design programs to restore and
manage stream resources on a regional scale?
1.16 In order to meet the needs of both EMAP-SW streams and Region III REMAP several different
sampling protocols have been put forward at this time. The protocols have been put forth for the
1993/1994 sample period to represent the needs of three major programs, 1994 EMAP-SW streams
PBot, the EMAP TIME project and 1994 Region III REMAP PDot These protocols are only final from the
standpoint of this pOot activity. They represent information gained from a 2/92 Indicator Workshop of
academic, State, and Federal indicator experts; discussion among the EMAP Stream Team; discussions
with U.S. Geologic Service NAWQA Program leaders; U.S. Pish & Widlife personnel; and discussions
with USEPA Region III State and Regional biologists, and the 1993 pBot field activity. Many
compromises had to be made to satisfy many good comments received from all these discussions. To
address many of these comments the protocols described later wDI be tested in the 1993/1994 sampling
period.
1.17 This, 'Environmental Monitoring and Assessment Program Surface Waters -1994 PDot Operations
and Methods Manual Por Streams,' addresses the field activities for all three sampling programs (EMAP*
SW streams pilot, EMAP TIME, & Region III REMAP). Not all of the protocols win be sampled for or
performed at the approximately 308 total number of sites. Listed below Is a summary of which protocols
will used by the sampling program:
PROTOCOLS/INDICATORS TO BE COLLECTED AT EMAP, REMAP, REMAP
REFERENCE, REMAP TEST & TIME A SITES:
Fish, macrolnvertebrates ( > 595 microns), chemistry, and qualitative physical habitat
PROTOCOLS/INDICATORS TO BE COLLECTED AT EMAP, REMAP REFERENCE, AND
REMAP TEST SITES:
Fish, Fish Tissue, rhacroinvertebrates ( > 595 microns), chemistry, sediment
metabolism, periphyton, and quantitative & qualitative physical habitat
1.18 The remaining sections of the EMAP-SW 1994 PDot Reid Operations and Methods Manual For
Streams provide an overview of field operations, daily activities at each sample site, stream site
verification, site characterization and location. The sections following these are the protocols to be
conducted at each site: water chemistry, physical habitat assessment (quantitative), periphyton, benthic
(sediment) metabolism, sediment toxidty sample collection, benthic Invertebrates, fish, fish tissue
indicator, and qualitative habitat assessment The last two sections are on safety and health as well as
sample labelling & tracking.
1.19 REFERENCES
Stoddard, J.L 1990. Plan for Converting NAPAP Aquatic Effects Long-Term Monitoring (LTM) Project to
the Temporally Integrated Monitoring of Ecosystems (TIME) Project U.S. Environmental
Protection Agency, Environmental Research Laboratory, Corvallis, OR. 97333.
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EMAP-SW-Streams, Introduction, Field Methods, Section 1, Revision No. 2, March 1994. Page 5 of 5
USEPA. 1991 a. Environmental Monitoring Assessment Program (EMAP) Laboratory Methods Manual
Estuaries. D.J. Wemm, LB. Lobring, J.W. Eichelberger, B.B. Potter, R.F. Thomas, J.M.
Lazorchak, G.B. Collins, and R.L Graves (eds.). EPA/600/4-91/024. Environmental Monitoring
Systems Laboratory, U.S. Environmental Protection Agency, Cincinnati, Ohio. 45268.
USEPA. 1991 b. Environmental Monitoring Assessment Program: Surface Water Implementation Plan-
Northeast PDot Lake Survey, Summer 1991. J.E Pollard and KM. Peres (eds.). Environmental
Monitoring Systems Laboratory, U.S. Environmental Protection Agency, Las Vegas, Nevada
89193.
USEPA. 1991 c. Environmental Monitoring Assessment Program: Surface Water 1991 Northeast Pilot
Field Operations and Training Manual. N.G. Tallent-Halsel! and G.D. Merritt (eds.).
Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency. Las
Vegas, Nevada 89193.
USEPA. 1991 d. Environmental Monitoring Assessment Program (EMAP): Field and Laboratory Methods
Manual 1991 Lake Pilot D.J. Wemm and J. M. Lazorchak (eds.). Environmental Monitoring
Systems Laboratory, U.S. Environmental Protection Agency, Cincinnati, Ohio 45268.
USEPA. 1992a. Environmental Monitoring Assessment Program: Surface Water 1992 Northeast Pilot
Field Operations and Training Manual. N.G. Tallent-Halsell and G.D. Merritt (eds.).
Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency, Las
Vegas, Nevada 89193.
o
USEPA. 1992b. Environmental Monitoring Assessment Program: Integrated Quality Assurance Project
Plan for Surface Waters Resources Group, Fiscal Year 1991. D.V. Peck (ed.). EPA/620/X-
91/080. Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency,
Las Vegas, Nevada 89193.
USEPA. 1992C. Environmental Monitoring Assessment Program (EMAP): Field and Laboratory Methods
Manual Used on Probability Freshwater Lakes. D.J. Wemm and J.M. Lazorchak (eds.).
Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency,
Cincinnati, Ohio 45268.
USEPA. 1993. EMAP-Surface Waters 1991 PDot Report D.P. Larsen and S.J. Cristie (eds.).
EPA/620/R-93/003. Environmental Monitoring Assessment Program, U.S. Environmental
Protection Agency, Washington, DC.
USEPA. 1994. Environmental Monitoring and Assessment Program Integrated Quality Assurance
Project Plan for the Surface Waters Resource Group 1994 Activities. Deb J. Chaloud and David
V. Peck (eds.). EPA/600/X-91/080. Environmental Monitoring Systems Laboratory, U.S.
Environmental Protection Agency, Las Vegas, NV 89193 and Environmental Research
Laboratory, U.S. Environmental Protection Agency, Corvallis, OR 97333.
Whlttier. T.R. and S.G. Pauisen. 1992. The surface waters component of the Environmental Monitoring
and Assessment Program (EMAP): An Overview. J. Aquatic Ecosystem Health 1:119-126.
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EMAP-SW-Streams, Field Operations Overiew, Section 2, Revision No. 3, April 1995, Page 1 of 2
SECTION 2
FIELD OPERATIONS OVERVIEW
2.1 The sampling program can be broken down into three elements:
EMAP sites - Probability sites from base EMAP sample grid
EMAP revisits - Repeat visits to 1994 and 1995 base EMAP sites for quantifying interannual and
sampling period variability
TIME sites - Augmented grid probability sites in acid-sensitive MAHA ecoregions.
2.1.1 A breakdown of each of these elements, listing the number of stations by state can be found in Table
2.1. The level of sampling effort varies considerably among the program elements. The full complement of
protocols (Table 2.2) will be measured at EMAP and EMAP revisit sites. At TIME sites, chemistry, benthos.
periphyton, and sediment metabolism will be measured.
2.2 Four sampling teams will be utilized to complete the field work. The EMAP and EMAP revisit sites will
be sampled by 1 four-person teams trained in all of the protocols. The TIME sites will be sampled by 3 two-
person teams. The teams will be made of personnel from DynCorp (Technology Applications Incorporated
(TAI)).
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EMAP-SW-Streams, Field Operations Overiew, Section 2, Revision No. 3, April 1995, Page 2 of 2
TABLE 2.1
MAHA 1994 SITE DISTRIBUTION BY STATE AND ELEMENT
STATE
MD
NY
PA
VA
WV
Total
EMAP
0
0
7
6
3
16
EMAP
Revisits
0
0
7
6
3
16
TIME
6
4
83
24
54
171
TOTAL
6
4
97
36
60
203
TABLE 2.2
MAHA PROTOCOLS BY SITE TYPE
Protocol
Chemistry
Fish
Fish Tissue
Benthos
Sediment Metabolism
Periphyton
Qua). (RBP) Habitat
Quantitative Habitat
Sediment Toxicity
EMAP
X
X
X
X
X
X
X
X
X
EMAP
Revisit
X
X
X
X
X
X
X
X
X
TIME
X
X
X
X
X
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EMAP-SW-Streams, Summary Daily Field Operations, Section 3. Rev. No. 2, March 1994, Page 1 of 2
SECTION 3
SUMMARY OF DAILY FIELD OPERATIONS FOR EMAP STREAM SAMPLING
3.1 The following schedule Is a general outline of the activities that a typical 4-person crew would be
involved in at an EMAP stream site. The field crew Is sub-divided into two sub-crews, the Geomorphs
and the Biomorphs, that reflects their Initial responsibilities more than their expertise.
3.2 GEOMORPHS
1. Arrive at site and verify locations (0.5 h) (Geomorphs)
a. GPS
b. local landmarks
c. determine average stream width for reach length determination
d. photodocumentatlon
3.3 BIOMORPHS AND GEOMORPHS
1. Grab chemistry samples (0.5 h) (Biomorphs)
a. cubltainer
b. 4 syringes
c. discharge
2. Mark lower reach boundary and lay-out 11 transects (1 h) (Geomorphs)
a. 5 transects above and below "X"
b. determine random location for biology samples
c. collect fine sediments for benthic metabolism and seditment toxiclty
d. set-up benthic metabolism
e. bag and label sediment toxiclty sample
3. Collect bugs and periphyton at transect locations (2,0 h) (Biomorphs)
4. Return to lower end of stream reach and begin physical habitat analysis (3.0-3.5 h) (Geomorphs)
a channel morphology-width, depth, slope, substrate, bank slope thalweg
b. canopy
c. woody debris
d. valley profile
5. Process bugs and periphyton (0.5-1.0 h) (Biomorphs)
6. RBP Physical Habitat Sheet, Stream Assessment Sheet (0.25 h) (Biomorphs)
7. Breakdown benthic metabolism study (0.25 h) (Geomorphs)
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EMAP-SW-Streams, Summary Daily Reid Operations. Section 3, Rev. No. 2, March 1994, Page 2 of 2
6. Fishing (2.0-3.0 h) (Geomorphs and Biomorphs)
7. Sample Tracking (0.5-1.0 h) (Biomorphs)
3.4 IN SUMMARY, EACH SUB-CREW IS RESPONSIBLE FOR:
GEOMORPHS BIOMORPHS
Reach verification Chemistry and discharge
Reach determination Macroinvertebrate collection
Marking transects Perlphyton collection
Physical habitat assessment RBP Physical habitat
Benthic metabolism Sample tracking
Fishing Fishing
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EMAP-SW-Streams, Initial Site Protocols, Section 4, Revision No. 2, March 1994, Page 1 of 5
SECTION 4
INITIAL SITE PROTOCOLS
4.1. INITIAL SITE SURVEY
4.1.1 Locating the Chosen Sample Site
4.1.1.1 Stream sample sites were chosen from the blue lines stream network represented on 1:100,000-
scale USGS maps following a systematic randomized design developed for EMAP stream sampling.
Sample sites were then marked with an "X" on finer-resolution 1:24,000-scaie USGS maps. This spot is
referred to as the X-site. Crews should use ail available means to Insure that they are at the correct site,
as marked on the map. including; 1:24,000 USGS map orienteering, topographic landmarks, county
road maps, and global positioning system (GPS) confirmation of site latitude and longitude. Upon
finding the X-site, fill out the stream verification form. The latitude/longitude of the site will be listed on
the stream information sheet that will be available to the crews for each sample site. Record the
latitude/longitude as displayed on the GPS for the X-site on the form. Also record the pertinent GPS
data on strength of signal (2-D or 3-D fix) in the appropriate box on the field form. If possible, wait for
the GPS to get a 3-D foe. This might require moving the GPS to a more open area near the X-site (GPS
works on line of sight to orbiting satellites). If the given latitude/longitude and the GPS
latitude/longitude differ by more than 10 seconds, double check the' vou are at the correct site. Give a
detailed description of the directions to the stream site in the indicate place on the form, use additional
pages, if necessary. We plan on revisiting the exact same sites in four years.
4.1.2. Determining Stream Site Status
4.1.2.1 We fully expect that not all chosen sites will turn out to be streams. On the basis of previous
synoptic surveys in the area, we have found that the maps are far from perfect representations of the
stream network. A significant part of EMAP is the estimation of the actual extent of streams in the area.
Once the crews have reached the marked sample point, evaluate the 'X* she and place it into one of
the following categories. The primary distinction is that the Nontarget categories have no stream
channel or are too deep to wade, the Target categories have a definable stream channel.
NONTARGET
1. No Stream Channel (map error) - Examination of the x-site showed no water body or stream channel.
2. Impounded stream - stream is submerged under a lake or pond due to man-made or natural (e.g.,
beaver dam) impoundments..-If the impounded stream, however, is stQI wadeable. record the
stream as Altered (Target class #4) and sample the stream.
3. Marsh/Wetland - standing water but no definable stream channel. In cases of wetlands surrounding
a stream channel, define the site as Target but restrict sampling to the stream channel.
4. Unwadeable Stream - A stream too deep to be safely sampled by wading following our current
protocols. If over half of the reach length is not safely wadeable, classify the reach as unwadeable
and do not sample it. If more than half of the survey reach is wadeable (e.g., only a couple of deep
pools) classify the reach as target and sample what can be safely sampled.
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EMAP-SW-Streams, Initial Site Protocols. Section 4, Revision No. 2, March 1994. Page 2 of 5
NOTE: In all cases, evaluate the site at the X-site, don't find* another spot to sample.
TARGET
1. Regular Stream • sampleable stream with our current protocols.
2. Intermittent Stream - Row of water is not continual at the site but the channel is wet See Subsection
4.3 for protocols for sampling intermittent streams.
3. Dry Channel • A discernible stream channel is present but there Is no water at the site. See
Subsection 4.3 for protocols for sampling dry streams.
4. Altered Channel • stream channel does not appear the way it Is marked on the map. However, there
is a stream in the field that appears to be the stream marked with the X-site on the map. An
example would be a channel rerouting following a flood event that cut off a loop of stream.
Position a new X-site at the same relative position in the altered channel Make careful notes and
sketches of the changes on the Site Verification form.
INACCESSIBLE
1. Physical Barriers • Crew is physically unable to reach the X-site because of heavy wetlands, steep
gorge or other barrier that prohibits safe entry, explain why in the explanations field.
2. No Permission - Crew is denied access to the site by the landowners, check off "NO" on the stream
sampled box on the site verification form and explain In detail in the explanations field.
4.1.2.2 Record the site class and pertinent site verification Information on the site verification form. If
the site Is in one of the target classes, proceed with sampling. If the site is nontarget or inaccessible.
the site visit is complete and sampling is finished.
4.1.3 Sampling During/After Rain Events
4.1.3.1 We wish to avoid sampling during high flow episodic events. For one, it is often unsafe to be in
the water during such times. In addition, biological and chemical conditions during episodes are often
quite different from those during baseflow. On the other hand, we can not restrict sampling to only strict
baseflow conditions. It would be next too impossible to define this with any certainty at an unstudied
site, and it would also greatly curtaB the sampling window of the survey. Thus, we wfll have to make
some compromise on when to sample/not sample a given system because of episodes. To a great
extent, we wfll depend on the judgment of the field crews. If you get to a site and you think it is unduly
nfluenced by episodic conditions, do not sample the site that day. Reschedule It for another visit The
following are some general guidelines:
1. The major indicator of episodic influence wfll be the condition of the stream itself. If it is running at
bank full discharge or the water seems much more turbid than typical for the class of stream do not
sample it that day. .
2. Do not sample if it is unsafe to wade in the majority of the stream reach.
3. Keep an eye on the weather reports and rainfall patterns. Do not sample a stream during periods of
prolonged heavy rains.
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EMAP-SW-Streams, Initial Site Protocols, Section 4, Revision No. 2, March 1994, Page 3 of 5
4. If the stream seems to be close to normal spring flows, and does not seem unduly influenced by
episodic events, go ahead and sample it, even if it has recently rained or is raining.
4.1.4 Taking Site Photographs
4.1.4.1 Site photographs are valuable In data analysis. If you do take any photographs at a stream,
start the sequence with one photograph of an 8.5 x 11 piece of paper with the stream ID and date in big
letters. After the photo of the stream ID, take two photographs at the X-stte, one in the upstream
direction and one downstream. Take any additional photos you find interesting after these first three
pictures. For pictures of fish or other small objects, place a card with the stream ID and date in the
snapshot
4.2 LAYING OUT THE STREAM SAMPLE REACH
4.2.1 Unlike chemistry, which can be measured at a point, most of the biological and habitat structure
measures require sampling a certain distance of stream to get an accurate picture of the ecological
community. Our pilot studies have suggested that a length of 40 times the channel width is necessary
to get over 90% of the species in the stream reach. Thus, we need to lay out a support reach 40
channel widths long around our X-site to characterize the community and habitat of our chosen sample
point
4.2.2 Upon reaching the site, survey the site according to the following protocol:
1. Determine channel width at the X-site by measuring the wetted width across the channel with a
surveyor's rod or tape measure at three places considered to be of typical" width within 10 meters
of the X-site. Average the 3 readings together and round to the nearest 1 m. Record this width on
the back side of the Stream Verification Form. For dry or intermittent channels, estimate a width
based on the unvegetated width of the channel.
2. Scout out the sample reach. One crew member wfll go upstream and one downstream to check out
the condition of the sample reach. Each person shall proceed unto they can see the stream to a
distance of 20 times the channel width determined in step #1 in the up/down stream direction from
the X-site. The objective of the scouting is to make sure the reach is dear of obstacles that would
bar a wadeable sampling approach.
NOTE: In streams less than 4 m wide, use 150 m as a minimum sample reach length. Each
person would then proceed 75 m from the X-sHe to lay out the reach boundaries.
3. There are some conditions that may require 'sliding* the stream reach around features we do not
wish to sample across. Do not proceed upstream into a lower order stream or downstream into a
higher order stream when laying out the stream reach (order based on 1:100,000 scale maps). Any
such confluences near the sample site wfll be marked on the 1:24,000 scale maps provided to the
crews. If such a confluence is reached, note the distance and flag the confluence as the reach end.
Make up for the loss of reach length by moving ("sliding*) the other end of the reach an equivalent
distance away from the X-site. Similarly, if you run into a lake, reservoir, or pond while laying out the
reach, stop, flag the lake/stream confluence as the reach end, and make up for the loss of reach
length by moving the other end of the reach an equivalent distance from the X-site.'
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EMAP-SW-Streams, Initial Site Protocols, Section 4, Revision No. 2. March 1994, Page 4 of 5
NOTE: Do not slide the reach to avoid man-made obstacles such as bridges, culverts, rip-rap, or
channelization. They are part of the effects that we're attempting to study.
4. Starting back at the X-sfte, measure a distance of 20 channel widths down the middle of the stream
using a tape measure (be careful not to 'cut comers'). At the 20 widths point, find the closest
habitat break and flag that location as the start point (transect A). If there is no apparent habitat
break for 3-4 channel widths (e.g.. a long pool or glide) flag the 20 widths point as the reach end.
and block net ft.
5. Using the tape measure, measure 1/10 (4 channel widths In big streams or 15 m in small streams) of
the required stream length upstream from the start point (transect A). Flag this spot as the next
cross-section or transect (transect B). For transect B. roll the dice to determine if ft is a left (L),
center (C). or right (R) sampling spot A roll of 1 or 2 indicates U 3 or 4 indicates C. and 5 or 6
indicates R (or use a digital wristwatch and glance at the last digit (1-3-L, 4-6-C, 7-9 »R). Mark L
C, or R on the transect flagging. Proceed upstream with the tape measure and lay out the 9
additional transects at 1/10 of the reach length intervals (transects C-K). Alternate the transects in
order as L.C.R after the first random selection,
6. At the last transect (K), which marks the end of the sample reach (40 channel widths), we wish to
end the reach at a habitat break. Thus, mark the last transect at the habitat break closest to the 40
width endpoint As was done for the start point, if there Is no habitat break within 3-4 channel
widths, mark the 40 width endpoint as the reach end. The last transect does not need an L,C or R
location as biology is not collected here. The final reach/transect layout should look like the one in
Figure 1 of Section 6. Physical Habitat Assessment
4.2.2.1 Before leaving the stream, complete a rough sketch map of the stream reach you sampled on
the back of the Stream Verification Form. In addition to any other interesting features that should be
marked on the map. note any landmarks/directions that can be used to find the X-site for future visits.
4.3 PROCEDURES FOR SAMPLING DRY AND INTERMITTENT STREAMS
4.3.1 Layout the survey stream reach as described in Subsection 4.2. For ALL target streams, whether
they are dry, intermittent, or completely flowing, complete the thalweg profile (form SH-01. see
Subsection 4.2) for the entire 40 channel width reach length. For dry locations, record stream
width/depth as 0. The thalweg profile will give us a record of the •water* status of the stream for future
comparisons (e.g., % of length with intermittent pools or no water).
i
4.3.1 At each of the transects (cross sections) laid out for sampling, classify the stream at that spot as:
DRY CHANNEL No surface water anywhere in cross section;
Complete the physical habitat sampling protocols at dry cross sections, use the
unvegetated area of the channel to determine channel width. This will describe
riparian condition, substrate type, and diagnostic physical habitat information.
DAMP CHANNEL Wet spots In cross section but NO flowing water or pods > 10 cm deep;
Do the physical habitat series of forms and collect periphyton samples from the
, wet spots. These are great environments for algae; if there are enough fine wet
sediments for metabolism, collect them as well.
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EMAP-SW-Streams, Initial Site Protocols, Section 4, Revision No. 2, March 1994, Page 5 of 5
ENDURING POOLS: No flowing water but pools > 10 cm deep;
Do complete physical habitat, algae, metabolism, and macroinvertebrate
sampling. This is essentially the standard biological protocol.
FLOWING WATER: Rowing water in cross section, follow standard protocols for all indicators.
4.3.1.1 Thus, physical habitat information is collected in all streams. Intermittent streams will have some
cross-sections with biological measurements and some with none. Totally dry channels will have no
biological sampling. Fish sampling and water chemistry sampling is discussed below.
4.3.2 Water chemistry sampling
4.3.2.1 The water chemistry sample needs to be collected at the X-site. If the X-site is dry but there is
flowing water or a pool with greater than 1 m2 of water surface area, more than 10 cm deep somewhere
along the survey reach, take the water sample and water chemistry measurements at the pool or flowing
water nearest the X-site. Note that the sample wasn't collected at the X-site and where on the reach the
sample was collected on the Sample Collection Form. If there is no spot on the survey reach with
flowing water or a pool greater than 1 m2 and 10 cm deep then do not collect a water sample.
4.3.3 Fish Sampling
4.3.3.1 In intermittent streams, fish sampling will take place in any wet areas along the 40 channel width
reach that are potential fish habitat (electrofishing or seining). Sampling will be restricted to the reach
length laid out previously. No fish sampling will be done in reaches th-t are entirely dry.
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EMAP-SW-Streams, Water Chemistry, Field Methods, Section 5, Revision No. 2, March 1994, Page 1 of 5
SECTION 5
WATER CHEMISTRY
5.1 INTRODUCTION
5.1.1 There are two components to collecting water chemistry information; collecting samples of stream
water to ship to the analytical laboratory, and In situ measurements of specific conductance, dissolved
oxygen, and temperature. At each stream, crews will fill one 4 L cubltalner and four 60 mL syringes with
streamwater. These samples wDI be stored In a coder packed with zlplock bags filled with ice and
shipped to the analytical laboratory at the University of Maine within 24 hours of collection. Streamside
measurements will be made using field meters and recorded on data forms. The primary function of the
streamwater samples and the Streamside chemistry measurements are to determine:
Acid-base status
Trophic condition 'nutrient enrichment)
Chemical Stressors
Classification of water chemistry type
5.1.2 Streamwater from the cubitalners is used to measure the major cations and anions, nutrients, total
iron and manganese, turbidity and color. The syringe samples are analyzed for pH, dissolved inorganic
carbon, and monomeric aluminum species. Syringes are used to seal off the samples from the
atmosphere because the pH, DIG, and aluminum concentrations wDI all change if the streamwater
equilibrates with atmospheric COZ Overnight express mail for these samples is required because the
syringe samples need to be analyzed, and the cubltalner samples stabilized (filtration and/or
acidification) within a short period of time (72 hours). Most of the procedures outlined in this section are
similar to the ones utilized by USEPA in streams for the National Surface Water Survey (Kaufmann et al.,
1988) and have been taken from the Handbook of methods for add deposition studies: Field operations
for surface water chemistry (USEPA, 1989).
5.2 COLLECTING WATER CHEMISTRY SAMPLES
1. Before leaving the base she, package the sample containers (one 4 L cubitainer and four, 60 ml
syringes) and stream sample beakers to prevent contamination; place the syringes in their plastic
container and the cubitainer and beakers in a dean ziplock bag. Label the cubitainer and each of
the four syringes with a bwcode label.
2. In the field, make sure that the syringes and the cubltalner have the same barcode label number.
Make sure the labels are securely attached.
3. Collect the water sample from the X-site in a flowing portion near the middle of the stream. Rinse
the 500 ml sample beaker three times with streamwater, discard the rinse downstream.
4. Remove the cubitainer lid and expand the cubitainer by. pulling out the sides. Fill the beaker with
streamwater and slowly pour 30-50 mL into the neck of the 4-L Cubitainer. Cap and rotate It so that
the water contacts all the surfaces. Discard the water. Repeat the above rinsing procedure two more
times.
NOTE: DO NOT BLOW into the Cubitainers to expand them, this wDI cause contamination.
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EMAP-SW-Streams, Water Chemistry, Reid Methods, Section 5, Revision No. 2, March 1994, Page 2 of 5
5. Fill the cubitainer with streamwater by pouring in 500 mL portions from the sample beaker. Let the
weight of the water expand the cubitainer. Fill the cubitainer to its maximum volume. Rinse the
cubitainer lid with streamwater.
6. Eliminate any air space from the Cubitainer. and cap it tightly. Make sure the cap is tightly sealed
and not on at an angle. Place the cubitainer Inside the 1 gallon ziplock bag It came in and seal the
bag.
7. If a cooler is available, place the cubitainer In the cooler (on ice or streamwater) and shut the lid. If
not, place the cubitainer In an opaque garbage bag and keep at the side of the stream in the
streamwater to keep cool while the rest of the sampling is completed.
8. Collect the 60-mL syringe samples from the same place that the cubitainer sample was collected.
Rinse the syringe by inserting the 60-mL syringe halfway Into the stream, sucking in a 15-20 mL
aliquot, pulling the plunger to its maximum extension and rotate the syringe so the water contacts
all surfaces. Discard the water by depressing the syringe plunger (point downstream). Repeat the
above rinsing procedure two times.
9. Insert the syringe into the stream again and slowly fin the syringe with fresh sample. Try not to get
any air bubbles in the syringe. If more than 1-2 tiny bubbles are present, discard the sample and
draw another one.
10. Invert the syringe (tip pointing up), cap the syringe with the syringe valve, and tap the syringe lightly
-to detach any trapped air bubbles. Make sure the valve to open and expel the air bubbles, and a
small amount of water leave between 50 and 60 mL of sample In the syringe. Close the syringe
valve. If any air bubbles were drawn into the syringe during this process, discard the sample and fill
the syringe again (step 9).
11. Fill all four syringes, make sure that the barcode label number on all four syringes are the same and
match the barcode number on the cubitainer. Place the syringes together In their solid plastic
shipping container and place in the cooler or in the streamwater with the cubitainer.
NOTE: All of the syringe measurements are affected by equilibration with atmospheric carbon dioxide, It
is essential that no outside air contact the samples collected for these determinations.
12. Record the barcode number (Sample ID) on the Sample Collection Form along with the pertinent
stream information (stream name, ID, date, etc.). Note anything that happens during the sampling
process that could influence sample chemistry (heavy rain, potential contaminants), if the sample
was collected at the X-slte, place an Xfor station collected. If you had to move to another part of
the reach to collect the sample, place the letter of the nearest transect In the station collected
columa Record more detaDed reasons and/or Information on the comments line.
13. After carrying the samples out to the vehicles, place the cubitainer and syringe case with syringes in
a cooler and surround with 1 gallon ziplock bags fDIed with ice.
14. Ship the coolers to the analytical laboratory by overnight courier at first opportunity, either that
afternoon or the next day (see Section 14, Information Management, on sample shipment). Place
fresh ice in the ziplocks in the cooler Just before shipping. Write word ICE* on ziplock bags with
indelible marker.
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EMAP-SW-Streams, Water Chemistry, Field Methods, Section 5, Revision No. 2, March 1994, Page 3 of 5
15. At the base site, after sampling, rinse the sample beakers three times with deionized water to
prevent contamination of the next days sample. Place the beakers in a 1-gallon ziplock bag with the
next days cubitainer for transport.
CAUTION: Throughout the water chemistry sampling process it Is Important to take precautions
to avoid contaminating the sample. Many of the streams have a very low ionic strength and can
be contaminated quite easily by perspiration from your hands, sneezing, smoking, bug repellent,
or other chemicals used during sample collection. Thus, make sure that none of the water
sample contacts your hands before going into the cubitainer. Do not apply to oneself any bug
repellant, perfume, or make up until after water sample is collected. Note any problems on field
forms.
5.3 STREAM SIDE CHEMICAL MEASUREMENTS
5.3.1 Introduction
5.3.1.1 The following protocols are intended for in situ measurement of streamwater specific
conductance, temperature, and dissolved oxygen. Specific conductance or conductivity is a measure of
the ability of the water to pass an electrical current which is related to the ionic strength of a solution.
Dissolved oxygen (DO) is a measure of the amount of oxygen dissolved in solution. In natural waters,
minimal concentrations of oxygen are essential for survival of most aquatic organisms. Measures of DO
and temperature are used to assess water quality and the potential for healthy aerobic organism
populations.
5.3.2. Measuring Stream Specific Conductance
1. Check conductivity pen for outward signs of fouling daily. Do not touch the electrodes inside the
probe with any object Always keep the pen's electrode moist by keeping deionized water in the pen
cap. If deionized water is not available, use streamwater or tap water rather than let the electrode dry
out.
2. Perform an initial field quality control check (QCC) before the stream conductivity measurement.
Remove the cap from the bottom of the conductivity pen. Swirl the pen for 3-5 seconds in the
•Rinse' QCC solution (Peck and Metcalf, 1991) and let stabilize for 20 seconds. Record the QCC
solution conductivity on the Field Measurement Form.
3. If the QCC solution measurements were not within 10 uS/cm of theoretical value (see bottle), repeat
the measurement process. If the value is still unacceptable, flag the conductivity data on the field
form. Recheck the QCC solution conductivity back at the base site with fresh bottles of QCC
solution. If tt is still off by more than 10 S/cm, clean the probe as described in the manual, check the
batteries, soak in deionized water for 24 hours, and try again. If this doesn't work, replace the
conductivity pen.
4. Rinse a 250 ml sample beaker three times with stream water and fill with 150 to 200 ml of stream
water for determination of stream conductivity.
5. Swirl the conductivity pen in the beaker for 5-10 seconds. Hold unswirled, for 20 seconds in the
sample to allow for the pen's temperature compensation mechanism to function. Don't let the pen
touch the bottom of the beaker. Record the stream conductivity on the Field Measurement Form.
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EMAP-SW-Streams, Water Chemistry, Field Methods, Section 5, Revision No. 2, March 1994, Page 4 of 5
Alternatively, the pen can be placed directly In a quiescent part of the stream and conductivity read
directly (needs the same equilibration time).
6. Routine Maintenance for conductivity pens
NOTE: Refer to the Instrument manual for probe cleaning Instructions.
a. Store the pens in delonized water by keeping deionized water in the bottom of the pen cap.
b. Before using a probe which has been stored dry, soak the electrodes in deionized water (by
filling the caps) for 24 hours.
c. Always turn off the pens after use by capping them.
d. Replace the bottle of QCC solution with fresh QCC solution every 2 weeks.
5.3.3. Dissolved Oxygen/Temperature
1. Calibrate the dissolved oxygen (D.O.) meter as described In Section 8. Benthic (Sediment)
Metabolism, Subsection 10, Calibration And Standardization.
2. .Place the 0.0. probe In mid-stream at mid-depth at the X-site. Allow the probe to equilibrate and
record the 0.0. and stream temperature on the Field Measurement Form. If the D.O. meter is not
functioning, measure the stream temperature with the field thermometer and record the reading on
the Reid Measurement Form along with pertinent data flags and comments.
NOTE: Stream dissolved oxygen and temperature values may be taken at the same time that the Initial
dissolved oxygen/temperature values are measured for the benthlc (sediment) metabolism studies
instead of the same time as the cubltainer sample Is collected.
5.4 REFERENCES
Kaufmann, P., A. Herlihy, J. Elwood, M. Mitch, S. Overtoa M. Sale. J. Messer, K.Reckhow, K. Cougan, 0.
Peck, J. Coe, A KJnney, S. Christie, 0. Brown, C. Hagley, and Y. Jager. 1988. Chemical
characteristics of streams In the mid-Atlantic and southeastern United States. Volume I:
population descriptions and physico-chemical relationships. EPA 600/3-88/021 a. U.S.
Environmental Protection Agency, Washington, DC
Peck, D.V., and R.C. Metcafl. 1991. Dilute, neutral pH standard of known conductivity and acid
neutralizing capacity. Analyst 116:221-231.
USEPA. 1989. Handbook of methods for acid deposition studies: field operations for surface water
chemistry. EPA 600/4-89/020. U.S. Environmental Protection Agency, Office of Research and
Development, Washington, DC.
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EMAP-SW-Streams, Water Chemistry, Field Methods, Section 5, Revision No. 2, March 1994, Page 5 of 5
Table 5-1. CHEMICAL SAMPLING CHECK-OFF SHEETS
EQUIPMENT TO CARRY IN FIELD FOR WATER CHEMISTRY
125 mL bottle of QCC solution in ziplock bag
Conductivity Pen
Field Forms
One , 250 mL plastic beakers
One, 500 mL plastic beaker
One cubitainer in clean ziplock bag (barcode label attached)
Four, 60 mL syringes in plastic container (each one with barcode label attached)
Four syringe valves in plastic container
Opaque garbage bag
EXTRA EQUIPMENT TO CARRY IN VEHICLE
Cooler with 4-6 one gallon ziplock bags filled with ice
Back-up labels, forms, syringe varves
DAILY ACTIVITIES AFTER SAMPLING
1. Enter chemistry tracking data from today's sample (bar code ID, stream ID, date, etc.) into computer
2. Check that cubitainer lid is on tight and has a flush seal
3. Prepare sample for shipping (label and seal cooler, replace ice as dose as possible to shipping time)
4. Call Overnight shipping company to arrange pick-up of cooler
5. Rinse all sample beakers with deionized water, three times
6. Make sure conductivity pens are rinsed with deionized water and are stored with moist electrodes
7. Barcode label the next days sample containers (cubitiner and syringes), pack cubitainer and sample
beakers in dean ziplock bag, and pack four syringes and syringe valves in plastic container.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 1 of 38
SECTION 6
PHYSICAL HABITAT ASSESSMENT
6.1 INTRODUCTION
6.1.1 Rationale
6.1.1.1 In the broad sense, physical habitat in streams includes all those physical attributes that influence
or provide sustenance to organisms within the stream. Stream physical habitat varies naturally, as do
biological characteristics; thus expectations differ even in the absence of anthropogenic disturbance.
Within a given physiographic-climatic region, stream drainage area and overall stream gradient are likely to
be strong natural determinants of many aspects of stream habitat, because of their influence on
discharge, flood stage, and stream power (the product of discharge times gradient). Summarizing the
habitat results of a workshop conducted by EMAP on stream monitoring design, Kaufmann (1993)
identified seven general physical habitat attributes important in influencing stream ecology. These
include:
Channel Dimensions
Channel Gradient
Channel Substrate Size and Type
Habitat Complexity and Cover
Riparian Vegetation Cover and Structure
Anthropogenic Alterations
Channel-Riparian Interaction
All of these attributes may be directly or indirectly altered by anthropogenic activities. Nevertheless, their
expected values tend to vary systematically with stream size (drainage crea) and overall gradient (as
measured from topographic maps). The relationships of specific physical habitat measurements
described in this EMAP-SW field manual to these seven attributes are discussed by Kaufmann (1993).
Aquatic macrophytes, riparian vegetation, and large woody debris are included in this and other physical
habitat assessments because of their role in modifying habitat structure and fight inputs, even though
they are actually biological measures. The field physical habitat measurements from this field habitat
characterization are used in the context of water chemistry, temperature, and other data sources (e.g..
remote sensing of basin land use and land cover). The combined data analyses will more
comprehensively describe additional habitat attributes and larger scales of physical habitat or human
disturbance than are evaluated by the field assessment alone.
6.1.12 This protocol is intended for evaluating physical habitat in wadeable streams. The protocol is most
efficiently applied during low flow conditions and during times when terrestrial vegetation is active, but may
be applied during other seasons and higher flows except as limited by safety considerations. It is
designed for monitoring applications where robust, quantitative descriptions of reach-scale habitat are
desired, but time is limited. The qualitative nature of the habitat quality rank scores produced by many
currently available habitat assessment methods has not been demonstrated, as yet, to meet the
objectives of EMAP, where more quantitative assessment is needed for she classification, trend
interpretation, and analysis of possible causes of biotic impairment.
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EMAP-SW-Streams, Physical Habitat Assessment. Section 6. Revision No. 2, March 1994, Page 2 of 38
6.1.1.3 The protocol we describe differs from other rapid habitat assessment approaches by employing a
randomized, systematic spatial sampling design that minimizes bias in the placement and positioning of
measurements. Measures are taken over defined channel areas and these sampling areas or points are
placed systematically at spacings that are proportional to baseflow channel width. This systematic
sampling design scales the sampling reach length and resolution in proportion to stream size. It also allows
statistical and series analyses of the data that are not possible under other designs. We strive to make the
protocol objective and repeatable by using easily learned, repeatable measures of physical habitat in place
of estimation techniques wherever possible. Where estimation is employed, we direct the sampling crew
to estimate attributes that are otherwise measurable, rather than estimating the quality or importance of the
attribute to biota or its importance as an indicator of disturbance. We have included the more traditional
\risualdassificationofchannelunitscale habitat types because they have been useful In past studies and
enhance comparability with other work.
6.1.1.4 The time commitment to gain repeatability and precision b greater than that required for more
qualitative methods. In our field trials, two people typically complete the specffied channel, riparian, and
discharge measurements in about 3 hours of field time. However, the time required can vary considerably
with channel characteristics. On streams up to about 4 meters wide with sparse woody debris,
measurements can be completed in as little as 2 hours, whereas crews may require up to 5 hours in large
(> 10m wide), complex streams with abundant woody debris and deep water.
6.1.1.5 The protocol defines the length of each sampling reach proportional to stream width and then
systematically places measurements to statistically represent the entire reach. Stream thalweg depth and
wetted width are measured at very tightiy spaced intervals; whereas channel cross-section profiles,
substrate, bank characteristics and riparian vegetation structure are measured at larger spacings. Woody
debris is talfied along the fuD length of the stream reach, and discharge is measured at one. The tightly
spaced depth and width measures aDow calculation of indices of channel structural complexity, objective
classification of channel units such as pools, and quantification of residual pool depth, pool volume, and
total stream volume.
6.1.2 Components of the Field Habitat Assessment
6.1.2.1 The Physical Habitat Protocol consists of four different components (Table 1). Measurements for
the first three components are recorded on 11 copies of a two-sided field form, with a separate form for
discharge measurements (both forms attached). The procedures for conducting each component are
discussed in four subsections of this training manual The first component (the THALWEG PROFILE) is a
longitudinal survey of depth, width, habitat class, and presence of fine/soft sediment at 100 equally
spaced points along the centerfine between the two ends of the sample stream reach (150 points in some
small streams - see 6.1.3.2). The second component b a continuous tafly of large WOODY DEBRIS along
the reach. The third component, the detailed channel and riparian CROSS-SECTIONS, includes
measures and/or visual estimates of channel cross-sectional dimensions, substrate, fish cover, bank
characteristics, and riparian vegetation structure at 11 equally-spaced stations along the length of the
reach. This third component also includes measurements of the gradient and compass bearing between
stations, providing information necessary for calculating reach gradient, residual pool volume, and
channel sinuosity. The fourth and final component is a measurement of instantaneous DISCHARGE at
one optimally chosen cross-section.
6.1.3 Habitat Sampling Locations on the Study Reach
6.1.3.1 Measurements are made at two scales of resolution along the mid-channel length of the reach;
the results are later aggregated and expressed for the entire reach, a third level of resolution (Rgure 1).
We want to assess habitat and other stream indices over stream reach lengths that are approximately 40
times their average wetted width. This allows us to adjust the sample reach length to accommodate
varying sizes of streams. Subsection 4.2 describes the procedures for locating the X-srte that defines the
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 3 of 38
midpoint of the sample reach. This sampling location is already marked on a 1:24,000 map prior to going
into field. It has precise coordinates of latitude and longitude, and was selected by the EMAP design
group using a randomized systematic sampling design. Subsection 4.3 prescribes the protocol for
delineating a sample reach that is 40 times its width, but no shorter than 150 meters. That section also
describes the protocol for measuring out (with tape measure) and flagging the locations of the 11 channel
gross sections where many habitat measurements will be made. The distance between each of these
cross section is 1/1 Oth the total length of the sample reach.
Table 1. Components of Physical Habitat Protocol
Component Description
1. Thalweg Profile: Measure maximum depth and wetted width, classify habitat, determine presence
of fine/soft sediment at 10-15 equally spaced intervals between each of 11
channel cross-sections (100-150 along entire reach).
2. Woody Debris: Between each of the channel cross sections, tally large woody debris numbers
within the bankfull channel according to size classes.
3. Channel and Riparian Cross-Sections: O 11 cross-section stations placed at
equal intervals along reach length:
- Measure: channel cross section dimensions, bank height, undercut,
angle (with rod and cfinometer); gradient (c&nometer), sinuosity (compass
backsite), riparian canopy cover (densbmeter).
- Visually Estimate*: substrate size class and embeddedness; areal cover
class and type (e.g., woody) of riparian vegetation in Canopy, Mid-Layer
and Ground Cover, areal cover dass of fish concealment features, aquatic
macrophytes and filamentous algae.
- Observe & Record*: human disturbances and their proximity to the
channel.
6. Discharge: In medium and large streams (defined later) measure water depth and velocity (O 0.6
depth with electromagnetic or inpefleMype flow meter) at 15 to 20 equally spaced
intervals across one carefully chosen channel cross-section. In very small streams,
measure discharge with a portable weir or time the filling of a bucket
* Substrate size class and embeddedness are estimated, and depth is measured for 5 particles taken © 5
equally-spaced points on each cross-section. The cross-section is defined by laying the surveyor's rod or
tape to span the wetted channel. Woody debris is tallied over the distance between each cross-section
and the next cross-section upstream. Riparian vegetation and human disturbances are observed 5m
upstream and 5m downstream from the cross section station. They extend shoreward 10m from left and
right banks, fish cover types, aquatic macrophytes, and algae are observed within channel 5m upstream
and 5m downstream from the cross section stations. These boundaries for visual observations are
estimated by eye.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2. March 1994, Page 4 of 38
f
Figure 1. Sample reach layout (plan view).
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 5 of 38
6.1.3.2 The thalweg profile measurements must be spaced evenly over the entire sample reach length.
In addition, they must be sufficiently close together that they do not "miss" deep areas and habitat units
that are in a size range of about 1/3 to 1/2 of the average channel width. Follow these specifications for
choosing the interval between thalweg profile measurements:
i
Channel Width < 2.5m — intervals = 1.0 m
Channel Width 2.5-3.Sm — intervals = 1.5 m
Channel Width > 3.5m — intervals = 0.01 (reach length)
Following these guidelines, you will be making 150 evenly spaced thalweg profile measurements in the
smallest category of streams, 15 between each detailed channel cross section. In all of the larger stream
sizes, you will make 100 measurements, 10 between each cross section. For practical reasons,
width measurements in stream reaches greater than 4m wide (having reach lengths
greater than 150 m) shall be taken only at the 11 cross-section stations. In all the
streams narrower than 4m (having reach lengths of 150 m), widths are measured at all
100 or 150 thalweg profile locations.
6.1.4 Logistics and Work flow
6.1.4.1 The four components of the Physical Habitat data collection are organized into three grouped
activities:
1. Thalweq Profile and Large Woody Debris Tally (Sections 6.2 and 6.3). Two people proceed
upstream from the reach start point (see Section 4.3) making observations and measurements at
the chosen increment spacing (see Subsection 6.1.3). One person b in the channel making
width and depth measurements, and determining whether soft sediment is present under his/her
staff. The other person records these measurements, classifies the channel habitat, and makes
the large woody debris estimates. Each time this team reaches a new cross section flag, they take
out a new copy of one of the 11 Channel/Riparian Cross Section forms. They interrupt the
thalweg profile and woody debris assessment activity to complete each cross-section as it comes.
The thalweg profile is continuous, but is measured in pieces, each recorded on the cross section
form where the Tittle segment begins.
2. Channel/Riparian Cross-Sections (Section 6.4^. One person proceeds with the channel
cross-section dimension, substrate, bank, and canopy cover measurements. The second person
records those measurements while making visual estimates of riparian vegetation structure, fish
cover, and human disturbance specified on the field form. Slope & bearing backsrtes are
accomplished together. Intermediate flagging (of a dBferent color) may have to be used if stream
is extremely brushy, sinuous, or steep to the point that you cannot she for slope and bearing
measures between the 11 points. (Note that the crews could tally woody debris while doing the
backsrte, rather than during the thalweg profile measurements.)
3. Discharge (Section 6.5). Discharge measurements shall be made after collecting the chemistry
sample. They are done at a chosen optimal cross section near the X-srte. However, do not use
the electromagnetic current meter close to where etectrofishing is taking place. Furthermore, if a
lot of channel disruption and sediment must be stirred up, wait on this activity until all chemical and
biological sampling has been completed.
6.2 THALWEG PROFILE
6.2.1 Thalweg" refers to the flow path of the deepest water in a stream channel. The thalweg profile is a
longitudinal survey of maximum depth and several other selected characteristics at 100 to 150 equally
spaced points along the centerline of the stream between the two ends of the stream reach. The
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2. March 1994, Page 6 of 38
EMAP-SW habitat assessment modifies traditional methods by proceeding upstream in the middle, of the
channel, rather than along the thalweg itself. One field person walks upstream (wearing fett-soled waders)
carrying a 1.5 - S meter fiberglass telescoping surveyor's rod (metric) and a 1m metre rule (ca^rated rod or
Dole) Aserondpersononthebankorinthestreamcamesaclipbc^
Debris Forms. Data from the thalweg profile allows calculation of indices of residual pool volume, stream
size, channel complexity, and the relative proportions of habitat types such as riffles and pools.
6 2.2 Use the surveyor's rod and meter stick to make the required depth and width measurements, and
to measure off the distance between sampling points as you proceed upstream. Ideally, each 10th
increment of thalweg measurements wffl bring you in fine with the flag marking a new cross-section profile.
The flag will have been set previously by carefully taping atong the channel, making the same bends that
you do while measuring the thalweg profile. However, you may stifl need to make minor adjustments to
align each 10th measurement with the cross section. In streams with average widths smaller than ZSm,
vou wifl be making thalweg measurements at 1 meter increments. Because the minimum reach length is
set at 150 meters, there will be 15 measurements between each cross section. Use the 5 extra fines on
the thalweg profile portion of each of the 11 field forms to record these measurements.
6 2 3 At each of the 100 increments of length upstream along the mid-channel fine, measure and record
the'greatest depth (to the nearest centimeter) in the channel on either side perpendicular to the center
One (Rgure 1). This deepest point, the thalweg depth", wifi not necessarily be along the mtetemel
line Whon momuring dopth uvfth tho ptrltr ~r r-"y"** "*" "**"*thft wntBI> depth on tne atqe CT Tnc rc°-
to avoid inaccuracies due to the wave formed by the rod in moving water. When placing your rod or staff
on the stream bottom, record the presence or absence of fine, soft seolrnents beneath yours^. Rne
soft sediments are defined here as fine gravel, sand, sift, day or nwck re«% apparent by "feefing-the
bottom with the staff.
6.2.4 At the same channel locations as for thaiwegdeptri. rrieaaure the wetted wk^pemendicular to
the mid^hannel fine. The wetted width boundary is the point at which substrate P^SSSSSUe
surrounded by free water. Width measurements are taken rapidlywhen they are part of the thalweg profile.
Read these width measurements to the nearest 0.1 meter for widths up to about 3 mete^ Thereafter,
read to approximately the nearest 5%. Thisis02mfbrwidthsd4to6m.OJmf6rwKithsd7to8m,and
0.5 m for widths of 9 or 10 m, and so on.
Omit the rapid multiple width measurements when doing th« thalweg profile on stream
reaches greater than 4m wide (le^ those having sample reach lengths greater than 150
m). In these larger streams, widths shall be measured only at th« 11 cross-section
stations. In all the streams narrower than 4m (having reach lengths of 150 m), widths
are measured at all 100 or 150 thalweg profile locations.
6 2.5 While recording the width and depth measurements and the presence of fine sediments, the
second person chooses and records the Habitat Class and the Pool Forming Element Code applicable to
each of the 100 sampling points along the length of the reach (Table 2; Rgure 2). The resulting database
of traditional visual habitat classifications win provide a bridge of common understanding with other
studies. Channel unit scale habitat classifications are to be made at the thalweg of the cross section. The
habitat unit itself must meet a minimum size criteria in addition to the qualitative criteria fisted in Table 2.
Before being considered large enough to be identified as a channel-unit scale habitat feature, the unit
should be at least as long as the channel is wide). For instance, if there is a small deep (pod-Eke) area at
the thalweg within a large riffle area, don't record it as a pool unless it occupies an area about as wrie or
long as the channel is wide.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 7 of 38
Table 2. Habitat Classification at Channel Unit Scale (See Rgure 2). Note that in order for a channel
habitat unit to be distinguished, it must be at least as wide or long as the channel is wide.
Class Code
Pools:
Plunge Pool PP
Trench Pool PT
Lateral Scour Pool PL
Backwater Pool PB
Dam Pool PD
Glide GL
Riffle H
Rapid RA
Cascade CA
Falls FA
Dry Channel OR
Description
Still water, low velocity, smooth, glassy surface, usually deep
compared to other parts of the channel:
Pool at base of plunging cascade or falls.
Pool fike trench in stream center.
Pool scoured along bank.
Pool separated from main flow off side of channel.
Pool formed by impoundment above dam or constriction.
Water moving slowly, with smooth, unbroken surface - tow
turbulence
Water moving, with small ripples, waves and eddies - waves not
breaking, surface tension not broken, sound: "babbling1,
•gurgling'.
Water movement rapid and turbulent, surface with intermittent
wh'rtewater with breaking waves - sound: Contint js rushing,
but not as bud as cascade.
Water movement rapid and very turbulent over steep channel
bottom. Most of water surface broken in short irregular plunges,
mostly Whitewater - sound Roaring.
Free faffing water over vertical or near vertical drop into plunge,
water turbulent and white over high falls, sound: from splash to
roar, depending upon discharge.
No water in channel
Code
N
W
R
B
F
WR, RW, RBW
O
Pool-Forming Element Category
Not Applicable, Habitat Unit is not a pool
Large Woody Debris.
Rootwad
Boulder or Bedrock
Unknown cause (unseen fluvial processes)
Combinations.
Other - note in comments
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 8 of 38
POOL
GLIDE
RIFFLE
RAPID
CASCADE
SIDE CHANNEL
FALLS
Rgure 2. Sketches of channel units (From Frisseil etai.,1986)
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 9 of 38
6.2.6 M'd-Channel Bars. Islands, and Side Channels pose some problems for the sampler conducting a
thalweo profile. These split channel features necessitate some guidance. Bars are defined here as
channel features below the bankfull flow mark that are dry during baseflow conditions (see Section 6.4.3
for definition of bankfull channel). Islands are channel features that are dry even when the stream is
experiencing a bankfull flow. Both bars and islands cause the stream to split into side channels. When a
bar is encountered along the thalweg profile, ft shall be noted as 'BAR* in the comments column and the
active channel is considered to include the bar. Therefore, the wetted width shall be measured as the
distance between wetted left and right banks. H is measured across and over bars and boulders If bars
are present record bar width in the space provided.
6.2.7 If a mid-channel feature is as high as the surrounding flood plain. H is considered an island. Treat
islands different than bars. The way to handle the ensuing channel shall be based visual estimates of the
percent of flow of the side channel as follows:
Side/Main Channel Flow % Action
< 15% Note side channel on form ("--")•
15-35% Note "SC* plus detailed channel & riparian cross-section on the side
channel. (Designate transect as XA, XB, etc on form)
>35% Note *SC* plus detailed channel & riparian cross-section PLUS another
tnatweg profile on the side channel.
62.8 When side channels are present, the comments column of the Thalweg Profile form should reflect
their presence with a continuous 'SC* note. This comment win begin at the point of convergence with the
main channel, their presence off-channel win be continuously noted, and the comments will show the
point of side channel divergence, tf it is a long side channel, a continuing arrow can be used to mark h in
the comments column to save time. When doing width measures with a side channel separated by an
island, include only the width of the main channel in the measures at the time and then measure the side
channel width separately as detailed above. For dry and intermittent streams, where no water is in the
channel, record zeros for depth and wetted width. Record habitat type as dry channel PR).
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision Na 2, March 1994, Page 10 of 38
6.3 LARGE WOODY DEBRIS MEASUREMENTS
&*••*
6.3.1 Methods for large woody debris (LWD) measurement are a simplified adaptation of those described
by Robison and Beschta (1990). This component of the EMAP Physical Habitat protocol allows
quantitative estimates of the number, size, total volume and distrfcution of wood within the stream reach.
LWD is defined here as woody material with small end diameter of at least 10 cm (4 in.) and length of at
least 15m (5 ft). Tally aO pieces of LWD that are at least partially in trie basetbw channel, the'active
channeP (Hood channel up to bankfuO stage), or spanning above the active channel . ' First,, tally
together afl the LWD that is at least partially in the bankfuD channel Zones 1, .or 2. shown in Figure 3.
Then taDy afl the LWD that b not actually within the bantfufl channel, but is at least partially spanning
(bridging) the bank fufl channel (Zone 3 in Figure 3). For both the Zone 1-2 wood and the Zone 3 LWD,
the field form provides 12 entry boxes for tallying debris pieces visually estimated within three length and
four diameter class combinations. Each LWD piece b tallied in only one box There are 12 size classes for
wood at least partially in Zone 1 and 2, and 12 for wood partially within Zone 3. Wood not partially within
those Zones b not tallied. LWD in the active channel b tallied over the entire length of the reach,
including the area between channel cross sections. As in the thaiweg profile, LWD measurements
between each channel cross section and the next one upstream are recorded on the first 10 channel
cross section forms. The location of the large end of the LWD determines the cross section to which it b
assigned.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 11 of 38
ZONE 4
BANKRJLL CHANNEL
ZONE 3
WATER SURFACE
ATBANKFULLFLOW
_
WATER SURFACE
AT LOW FLOW
ZONE 4
Rgure 3. Large woody debris and influence zones (From Robison and Beschta, 1990).
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6. Revision No. 3, April 1995, Page 12 of 36
6.3.2 For each LWD piece, first visually estimate its length and its large and small end diameters in order
to place it In one of the diameter and length categories. The diameter classes are 0.1 m to <0.3m, 0.3m to
<0.6m, 0.6m to <0.8m, and >0.8ra The length dasses are 1.5m to <5.0m, 5m to < 15m, and > 15m.
Sometimes LWD is not cylindrical, so it has no dear •diameter*. In these cases visually estimate what the
diameter would be for a piece of wood with circular cross section that would have the same volume. When
evaluating length, indude only the part of the LWD piece that has a diameter greater than 10 cm (4 in). The
diameter dass on the field form (Rgure 3) refers to the large end diameter. Count each of the LWD pieces
as one tally entry and indude the whole piece when assessing dimensions, even if part of it is in Zone 4
outside of the bankfull channel
6.4 CHANNEL AND RIPARIAN CROSS-SECTIONS
6.4.1 Slope and Bearing
6.4.1.1 The slope, or gradient, of the stream reach is useful In three dffferent ways. First, the overall stream
gradient is one of the major stream classification variables, giving an indication of potential water velocities
and stream power, both of which are in turn important controls on aquatic habitat and sediment transport
within the reach. Second, the spatial variability of stream gradient Is a measure of habitat complexity, as
reflected In the diversity of water velocities and sediment sizes within the stream reach. Lastly, using
methods described by Stack (1989) and other methods currently being developed (Robison and
Kaufmann, 1994), the water surface slope wfll allow us to compute residual pool depths and volumes from
the multiple depth and width measurements taken in the thalweg profile (Subsection 6.2). Compass
Bearings between cross section stations, along with the distance between stations, wfll allow us to estimate
the sinuosity of the channel (ratio of the length of the reach divided by the straight line distance between
the two reach ends).
6.4.1.2 You will measure slope and bearing by "backslting" downstream between cross-section station 2
and 1,3 and 2,4 and 3, etc., up to the 11th cross section (Rgure 4). To measure the slope and bearing
between adjacent stations, use a dinometer, rangefinder compass, tripod, tripod extension, and flagging,
following the following six step procedure. If you are doing these measurements with another person,
replace the tripod" with your co-worker in the following protocol, but be sure that you site to YOUR EYE
LEVEL when backsiting on your co-worker. Also, be sure that he or she is standing AT THE WATER'S
EDGE or IN THE SAME DEPTH OF WATER AS YOU ARE The intent is to get a measure of the WATER
SURFACE SLOPE, which may not necessarily be the same as the bottom slope.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 13 of 38
1
Slope.
move k sde «fcAoime
Measurem^ts
Rgure 4. Channel slope and bearing measurements.
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EMAP-SW-Streams, Physical Habitat Assessment. Section 6, Revision No. 2, March 1994, Page 14 of 38
1. Set up the tripod in shallow water or at the water's edge at Station 1. Standing tall in a position with
your feet as near as possible to the water surface elevation, set the tripod extension with a piece
of flagging at eye level. Remember the depth of water in which you are standing when you adjust
the flagging to eye level.
2. Walk upstream to Station 2 (ten Increments* upstream), where the next detailed cross section is
located.
3. Rnd a place to stand at Station 2 that is at the same depth as where you stood at the previous
station when you set up the eye-level flagging. With the clinometer, site back downstream at your
flagging at Station 1; read and record PERCENT Slope on the field form. Be careful, the
clinometer reads both percent slope and degrees of the slope angle. PERCENT SLOPE IS THE
SCALE ON THE RIGHT HAND SIDE AS YOU LOOK THROUGH THE CLINOMETER
4. Stand now in the in the middle of the channel at Station 2, and site back with your compass to the
middle of the channel at Station 1 and record your compass bearing ('Azimuth'). It does not
matter for these measurements whether or not you adjust your compass bearings for magnetic
declination, but IT IS IMPORTANT THAT YOU ARE CONSISTENT IN THE USE OF MAGNETIC OR
TRUE BEARINGS throughout all the measurements you make on a given reach. Also guard
against recording 'reciprocaT bearings (erroneous bearings 180 degrees from what they should
be). The best way to do this is to know where the primary (cardinaO directions are in the field-
north (0 degrees), east (90 degrees), south (180 degrees), and west (270 degrees) - and insure
;- that your bearings'make sense."
5. Retrieve the tripod from the downstream cross section station (#1), (while doing a debris tally) and
set it up at Station 02 (just like step 1) before proceeding to station 3 to begin a new set of
cross-section measurements and backsitings.
8. When you get to each new detailed cross section station, backsfte on the previous station
repeating steps 2 through 5 above.
6.4.1.3 As stated earlier, ft may be necessary to set up intermediate slope and bearing stations between
the normal 11 stations if you do not have DIRECT UNE-OF-STTE ALONG (AND WITHIN) THE CHANNEL
BETWEEN STATIONS. This can happen if bush is too heavy, or if there are sharp slope breaks or tight
meander bends. Mark these intermediate station locations with a different color of plastic flagging than
used for the normal 11 stations, to avoid confusion. Record these supplemental slope and bearing
backsites in the appropriate places on the field form. It is important also to clearly identify the channel
position (increment number) of these supplemental measurements on the thalweg profile form and the
cross-section & riparian form in their respective comments sections.
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6.4.2 Substrate and Channel Dimensions
6.4.2.1 Substrate size is one of the most Important determinants of habitat character for fish and
macroinvertebrates in streams. Along with bedform (eg. riffles and pools), substrate Influences the
hydraulic roughness and consequently the range of water velocities In the channel. It also influences the
size range of interstices that provide living space and cover for macroinvertebrates, salamanders, and
sculpins. Sediment characteristics are often sensitive indicators of the effects of human activities on
streams. Decreases in the mean sediment size and Increases in the percentage of fine sediments, for
example, may destabilize channels and Indicate changes in the rates of upland erosion and sediment
supply.
6.4.2.2 In this EMAP protocol, substrate size and embeddedness wOl be evaluated at each of the 11
detailed cross sections using a combination of procedures adapted from those described by Wolman
(1954), Bain etal. (1985), and Plafkin eta!., (1989). The basis of the procedure is a systematic selection of
5 substrate particles from each of the 11 channel cross sections (Rgure 5). In the process of measuring
sediment sizes at each channel cross section, you will also make measurements of the wetted width of the
channel and the water depths at each sediment sample point If the wetted channel Is split by a
mid-channel bar (see section 6.2), the five substrate points shall be centered between the wetted width
boundaries regardless of the bar in between. Consequently, sediment particles selected In some
cross-sections may be 'high and dry*. For dry channels, make cross-section measurements across the
unvegetated portion of the channel.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 18 of 38
Rgure 5. Substrate sampling cross-section.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 17 of 38
1. At the cross-section station, lay the surveyors rod across the channel perpendicular to the flow,
with the "zero" end at the left bank (facing downstream). If the channel is too wide for the rod,
stretch the metric tape in the same manner.
2. Divide the width measurement of the channel by 4, and record in the left column "DistLB* of the
form the distances corresponding with 1/4th, 1/2, and 3/4 the distance across the stream, starting
with the left bank at zero. The distance you record to the right bank is the same as the wetted
channel width. (NOTE: this is the same wetted width measurement is also recorded under 'Bank
Measurements' on the form (Section 6.4.3). The sediment sampling points along the rod or tape
will be at the 1/4th, 1/2, and 3/4 positions, and at the water's edge just within the left and right
banks.
3. Place your sharp-ended meter stick at position 1 (Om) at the end of the tape. Read and record the
depth. Pick up the substrate particle (unless it is bedrock or boulder), and visually estimate its
particle sfee. according to the following coded scale. To minimize bias in this method, it is
important to concentrate on correct placement of the measuring stick along the tape and to select
the particle right at the bottom of the stick (not, for example, a more noticeable large particle that is
just to the side of the stick).
RS Bedrock (Smooth) >4000 mm — Smooth surface rock or hardpan (bigger than a
car)
RR Bedrock Rough >4000mm — (bigger than a car)
B L Boulders >250 to 4000 mm — (basketball to car size)
CB Cobbles 64 to 250 mm — (tennisball to basketball)
GC Gravel (Coarse) 16to64 mm— (marbletotennisbair)
GF Gravel (Fine) 2 to 16 mm — (ladybug to marble)
SA Sand 0.06 to 2 mm — (
-------
EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 18 of 38
Rgure 6. Schematic on determining embeddedness.
-------
Y-creams, rayi».'~c
6.4.3 Bank Characteristics
UERJRM
Loc.
ft
\
-------
EMAP-SW-Streams, Physical Habitat Assessment. Section 6. Revision No. 2. March 1994, Page 20 of 38
a.
b ,
. .
Incised
OF
»
Rgure 7. Schematic showing bankfdl channel and incision for channels: (a) not recently incised, and (b)
recently incised into valley bottom.
-------
EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 21 of 38
1. To measure bank angle, lay the surveyors rod or your metef stick down against the left bank, with
one end at the water's edge (left bank defined when facing downstream). Lay the clinometer on
the rod, read and record the bank angle in degrees, using the external scale on the clinometer. A
vertical bank is 90 degrees; undercut banks have angles greater than 90 degrees approaching
180 degrees, and gradually sloped banks have angles less than 90 degrees. Procedures for
measuring bank angle are shown in Rgure 8, as adapted from Platts et aL (1983).
2. If the bank is undercut, measure the horizontal distance of the undercutting.
3. Repeat steps 1 and 2 on the right bank.
4. Hold the surveyor's rod vertical, with Hs base planted at the water's edge. Using the surveyor's rod
as a guide while examining both banks, estimate (by eye) the amount of channel incision as the
height up from the water surface to elevation of the first terrace of the valley floodplain (Note this is
at or above the bankfull channel height). In cases where the channel is cutting a valley sideslope
and has oversteepened and destabilized that slope, the bare "cutbank* is not necessarily an
indication of recent incision. Examine both banks to make a more accurate determination of
channel downcutting.
5. Still holding the surveyors rod as a guide, examine both banks to estimate and record the height
of bankfull flow above the present water level. Spotting the level of bankfull flow during baseflow
conditions requires judgement and practice; even then ft remains somewhat subjective. In many
cases there is an obvious slope break that differentiates the channel from a relatively flat floodplain
terrace higher than the channel. Because scouring and inundation from bankfull flows are often
frequent enough to inhibit terrestrial vegetation, the bankfull channel may be evident by a
transition from exposed stream sediments to terrestrial vegetation. Similarly, it may be identified
by noting moss growth on rocks along the banks. The bankfull flow level may also be seen by the
presence of drift material caught on overhanging vegetation.
6. Measure and record the bankfull channel width the wetted width, and the width of exposed
mid-channel bars fif present). The latter two measurements wfll probably be done while making
substrate measurements across the channel.
7. Repeat steps 1 through 6 upon proceeding to each new cross section.
-------
EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 22 of 38
Using a clinometer to measure
a bank angle of 45°.
Using a clinometer to measure
a bank angle of 145°.
Rgure 8. Measuring bank heighland bank angle (Adapted from Platts et aL. 1983).
-------
6.4.4 Canopy Cover (Densiometer)
IV. CANOPT- COVER MEASURBHEKIS
CENL
FUO
.<•/<*
"s?
*
FUO
6.4.4.1 Riparian canopy cover over a stream is important not only in its role In moderating stream
temperatures through shading, but also as an indicator of conditions that control bank stability and the
potential for inputs of coarse and fine paniculate organic material. Organic Inputs from riparian vegetation
become food for stream organisms and structure to create and maintain complex channel habitat
6.4.4.2 Vegetative cover over the stream shall be measured at each of the 11 detailed cross section
stations. This measurement employs the Convex Spherical Densiometer, model B (Lemmon, 1957). This
instrument is available from only one vendor (see ordering information in section 6.7). The densiometer
must be taped exactly as shown in Rgure 8 to limit the number of square grid intersections to 17. To take
a canopy cover density measurement, the observer looks down on the densiometer held lust above waist
level, concentrating on these 17 points of intersectloa If the reflection of a tree or high branch or leaf
overlies any of the intersection points, that particular Intersection is counted as having cover. The measure
to be recorded on the form is the count (from 0 to 17) of all the intersections that have vegetation covering
them. Therefore, a greater number indicates increasing canopy extent and density. In doing the
measurement. It is important that the densiometer be leveled using the bubble level (Rgure 9).
6.4.4.3 For each of the 11 stations, densiometer measures are taken separately in four directions standing
at the center of the stream.. These measures wfll be used to estimate canopy cover over the channel.
Then, standing at the wetted bank at the left and then the right side of the stream at each station, repeat the
canopy density measurements facing the riparian vegetation (your back towards the stream). These bank
densiometer readings complement your visual estimates of vegetation structure and cover within the
riparian Itself, and are particularly Important In wide streams, where riparian canopy may not be recorded
by the densiometer standing mid-stream.
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EMAP-SW-Streams. Physical Habitat Assessment, Section 6, Revision Na 2. March 1994, Page 24 of 38
6.4.5 Riparian Vegetation Structure
m: VISUAL RIPARIAN
?S;x«i
6451 The previous section (8 .4.4) descAed methods for ^quaitfrymg the cover of canopy over the
and amount of daferent types of riparian vegetation.
level ddsturbafx»dthe^ean corridor. It also indkatea the present and future potential for vanous
types of organic inputs and shading.
6.4.5.2 Observations to assess riparian vegetation apply to the riparian area ™ *******
dovmstreamSmetersfromeachdthellcioss^ecttonst^
from the stream back a distance of 1 0m
X 1 0m riparian plot on each side of the stream (Figure 1 0). If the wetted diarnel b spa by a
bar, the bank and riparian measurements shall be for each side of the channel, not the bar.
-------
EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 25 of 38
TAPE
BUBBLE LEVELED
Rgure 9. Schematic:.Convex spherical canopy densiometer (From Mulvey et al., 1992). In this example.
10 of the 17 intersections show canopy cover, giving a denstometer reading of 10.
-------
EMAP-SW-Streams. Physical Habitat Assessment, Section 8. Revision No. 2, March 1994, Page 26 of 38
a«d( Fish Cover ObservaK
on Qr*
1 on
c^
Rgure 10. Boundaries for visual estimation of riparian vegetation, fish cover, and human influences.
-------
EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 27 of 38
6.4.5.3 Conceptually divide the riparian vegetation into three layers: a CANOPY LAYER (>5m high), an
UNDERSTORY (0.5 to 5m high), and a GROUND COVER layer (75%).
These ranges of percentage areal cover corresponding to each of these codes are also shown on the
Reid Form. When rating vegetation cover types, mixtures of two or more eubdominant classes might afl
be given sparse f1") moderate ("2") or heavy ("3") rankings. One very heavy cover dass wfth no clear
subdominant dass might be ranked'4" with afl the remaining classes efiher moderate C2"), sparse f 1") or
absent CO"). Two heavy classes with 40-75% cover can both be ranked "3".
6.4.6 Fish Cover, Algae, Aquatic Macrophyto*
m>*
> 0 J M 8101
< OJMOhUtL)
* 1 i
h
, fish
6.4.6.1 This portion of the EMAP physical habitat protocol
semi^uantrtatrvely, the type and amount of important types d ^^f»^
Atoneand in combination with other metrics, this information is used to assess h
cover, and channel disturbance.
6462 Observations to assess fish cover and several other in-channel features apply to the
5 TSEZti downstream 5 metersfrom each of the 1l€~~»>n ~ 2S
^^^
-------
EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision Na 3. Apri 1995. Page 28 of 36
6.4.6.3 Filamentous algae pertains to long streaming algae that often occurs in slow moving waters.
Aquatic macrophytes are water loving plants in the stream that could provide cover for fish or
macroinvertebrates. If the stream channel contains live wetland grasses, include these as macrophytes.
Woody debris are the larger pieces of wood that can Influence cover and stream morphology.
Brush/woody debris pertains to the smaller wood that primarily affects cover but not morphology. The tree
or brush within one meter of the surface is the amount of brush, twigs, small debris etc. that Is not in the
water but Is dose to the stream and provides cover. Boulders are typically basketball to car sized
particles. Many streams have artificial structures in them designed for fish habitat Streams may also have
in channel structures for diversions, impoundment and other purposes. Record the cover of these
structures on the form.
6.4.7 Human Influences
6.4.7.1 The field evaluation of the presence and proximity of various important types of human land use
activities In the stream riparian area is used In combination with mapped watershed landuse information to
assess the potential degree of disturbance of the sample stream reaches.
6,4.7.2 At each of the 11 detailed Channel and Riparian Cross-Sections, evaluate the presence/absence
and the proximity of 11 categories of human Influences. Confine your observations to the stream and
riparian area within 5m upstream and 5m downstream from the station (Figure 10). Entries are:
*B for any and all the Items you observe ON THE STREAMBANK. :
'C* for any and all the Items that you observe within 10m from the streambank f CLOSE).
*P* for those items that are PRESENT, but farther than 10m from the bank.
•0' for those NOT PRESENT.
Evaluate and record separately the left and right sides of the channel and banks. For human influence
items outside of the riparian observation plots, mark *P* for each and every station where they are visible,
even if you are observing the same item from station to station.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 29 of 38
6.5 DISCHARGE MEASUREMENT
6.5.1 Background
6.5.1.1 If specified, discharge shall be measured on EMAP-SW sample reaches. Discharge .
measurements are critical in parts of EMAP that focus on trends in the acidity of streams, a characteristic
which is very sensitive to streamflow differences. Discharge should be measured as close as possible to
the EMAP 'X' point where chemicaJ samples are taken, so that the discharge and chemistry are a
corresponding pair. However, different types of channel require different methods of discharge
measurement, and the kinds of places within a channel where the three alternative types of discharge
measurement methods can be employed differ. In streams sufficiently large to use the electromagnetic or
impeller-type water velocity meter, discharge wffl be measured according to the standard Velocity-Area
Method (Linsley et al., 1982) descnbed in Subsection 6.62. In streams that are too small to employ that
method, use the Portable Weir method (Subsection 6.6.3) or time the filling of a volume of water in a
calibrated bucket (Subsection 6.6.4). If none of these methods can be employed, measure the
cross-sectional depth profile and time the movement of a neutrally buoyant object (eg. an orange) through
a measured (and recorded) length of the channel
6.5.2 Velocity-Area Discharge Measurement
6.5.2.1 Electromagnetic Current Mater Calibration Chack
NOTE This procedure was written assuming a Marsh-McBimey Model 2010 electromagnetic current
meter is being used. This procedure may be used, with modfficaiion, with other meters meeting
equivalent specifications.
1. Check meter calibration daily as part of morning routine, then again on site before entering the
stream.
2 Calibration value should be 2,00 ± 0.05. The value obtained during morning calibration should be
recorded in the comments section of the Discharge form. The values obtained onsrte should also
be recorded on the Discharge field form.
3. Once per week the zero value should be checked in a bucket of still water. The probe should sit
for 30 minutes with no disturbance. The value obtained should be 0.0 ± 0.1. The meter zero
should be adjusted if it b outside this range.
6.5.2.2 Calibration and Care of Impeller-Type Current Meter
NOTE This procedure was written assuming a Swoffer Model 2100-14 is being used. It pertains, with
modification, to other meters meeting equivalent specifications. Read the Owners' Manual and the Care
and Operating instmnfinna nrovidpH hy th« Manufacturer REFQRE using the instrument. As a reminder,
some of the DO's and DONTS are:
1. The propeller assembly, including the fibre-optic sensor, rotor bearing, rotor shaft and the
propeller itself, is the single most important part of the instrument Great care must be observed to
insure that this precision instrument gives accurate output.
2. NEVER put the wading rod back into the carrying case, transport ft, or store it with the propeller
rotor assembly still attached.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2. March 1994, Page 30 of 38
3. The sensor and the rotor bearings may have to be cleaned after emersion in turbid water, or if they
sink into soft sediments. Do not scratch the photo-optic sensor and do not use oil on the rotor
bearing.
4. Before using, set the instrument to read In meters per second and calibrate as explained in the
owners'manual
6.5.2.3 Location for Discharge Measurements by Velocity-Area Method
6.5.2.3.1 The basis of this method is that stream discharge is equal to the product of the mean water
velocity and the vertical cross sectional area flowing water. Because velocity and depth typically vary
greatly across a stream, accuracy in field measurements is achieved by measuring the mean velocity and
flow cross-sectional area of many increments across a channel (Rgure 11A). Each increment gives a
subtotal of the stream discharge, and the whole is calculated as the sum of these parts. Discharge
measurements shall be made at only one carefully chosen channel cross section In a stream reach. It is
important to choose a channel cross section that b as much like a canal as possible. Picka cross section
where flow lines are directed downstream to the extent possible (low turbulence and a minimum of
cross-flow). Glides with *U* shaped channel cross auction.^ USAffllty provide the beat conditions for
measuring discharge by the velocity-area method. Try to avojd cross sections with a lot of large boulders.
woodv debris, brush, exposed rocks, and other irregtriaritiM. Do not measure discharge in a pool.
6.5.3.3 Velocity-Area Procedures
1; Facing downstream and beginning on the toft edge of the water, stretch a meter tape across the
chosen stream location perpendicular to stream flow.
2. Measure and record the stream width. Leave the tape tightly suspended across the stream,
approximately one foot above water level
3. Divide the total stream width into 15 to 20 equal-eked intervals. To determine interval width, divide
the total stream wetted width by 15 and round down to a convenient number. This should yield a
sufficient number of intervals.
4. If using the electromagnetic current meter, attach its probe to the wadino rod and check the
internal electronics by turning the switch to 'GAL' If the meter ca&bratas to 2.00 ±0.05 in air,
record this value in the comments section and proceed with the measurements. H using tha
impelleMvpa current meter, instafl its propeder on the sensor boom of the wading rod. Prior to
making measurements, the meter shall love been set and caCbratad to read in meters per
second.
5. Move to the center of the interval number 1, closest to the left bank (looking downstream).
6. Read and record th» stream depth in centime*era at tha canter of tha tntenml
7. Place the current meter probe at 0.6 of the total depth (measured from the surface). Orient the
probe or propeller to face upstream, and make sure that your feet or legs are not deflecting or
impounding flow that passes over the probe or propeller.
8. For the electromagnetic current meter fey. Marsh-MeBimey). use the lowest time constant scale
setting on the meter that provides stable readings. Wait 20 seconds to allow the meter to
-------
EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 31 of 38
equilibrate, then measure and record the current velocity at the center of the interval. For the
impeller-type meter (eg Swoffer 2100V set the control nob at the mid-position of "DISPLAY
AVERAGING". Press 'RESET then "START1 and proceed with the measurements. Wart 20
seconds for the meter reading to stabilize and read current velocity in meters per second.
9. Repeat steps (6) (7)(8) for all intervals across the stream.
6.5.4 Portable Weir Discharge Measurement
6.5.4.1 Discharge in channels too "smalT for the velocity-area method can be measured with a portable,
sharp-crested, 60 degree V-notch weir. Small can be defined as a channel so shallow that the probe can
hardly be placed in the water or the stream is so broken up and irregular due to rocks and debris that a
velocity area method measurement is not possible. The weir must completely block the flow and reroute it
through its notch. H must be high enough to create a small spillway as water flows over its sharp crest. In
other words, donl let the water level downstream interfere with the smooth plunge of the water flowing
over the weir. In addition, the weir must create a relatively flat surface of impounded water upstream for a
distance at least equal to the width of the stream.
6.5.42 Choose a stream cross section that is narrow and easy to block wfth the portable weir. Ideal
sections might be the 'lip* or tail out* below a pod. A bad choice would be right in the middle of a
cascade. Impound the flow with the weir, making sure that water b not flowing beneath or around the side
of the weir. Use mud or stones and plastic sheeting to get a good waterproof seal Insure that the weir is
level from side to side, and that it is perpendicular to the streamflow. When the depth of flow stabilizes,
read and record the height of impounded water above the V-notch (Figure 11B). It may take some time for
the flow to equilibrate (eg., 15 min), as the impounded pond buOds upstream from the weir. For small
discharges, water velocities through the weir will be slow, allowing this height to be measured very dose to
the weir. Larger discharges result in more rapid velocities, producing a pronounced downward curvature
of the water as H approaches the crest of the weir. The height measurement in these cases must be wed
back upstream (say 0.5 m) from the weir, and may necessitate using the ruler and clinometer as a level to
match the pool height against the weir.
IF INSTALLING THE TEMPORARY WEIR REQUIRES A LOT OF CHANNEL DISTURBANCE OR STIRS
UP A LOT OF MUD, WAIT TO DO THIS UNTIL AFTER ALL BIOLOGICAL AND CHEMICAL
MEASUREMENTS AND SAMPLING HAVE BEEN COMPLETED.
6.5.5 Timed Bucket Discharge Measurement
6.5.5.1 Where feasible in very small streams, Discharge can be measured directly, by recording the time it
takes to fill a known volume. This can be an extremely precise and accurate method. Choose a natural
spillway or plunge, or construct a temporary one using a weir, using plastic sheet or on-site materials. '
Using a stopwatch and a calibrated bucket held beneath the spillway, record the filling time and the volume
of water collected in the bucket (see Note in Figure 11 B). Make sure that the entire flow of the stream is
going into the bucket. Repeat the procedure 10 separate times. You may need to time the volumes
obtained from more than one different spillway in the same channel cross section. If so, indicate clearly
the time and volume data replicates that should be averaged together for each spillway.
IF THE TIMED BUCKET DISCHARGE MEASUREMENTS INVOLVE A LOT OF CHANNEL DISTURBANCE
OR STIR UP A LOT OF MUD, WAIT TO DO THIS UNTIL AFTER ALL BIOLOGICAL AND CHEMICAL
MEASUREMENTS AND SAMPLING HAVE BEEN COMPLETED.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 32 of 38
Loca
o**
A. Ve loci la-Area \)ischarae, Measurement
J ^
iO«i*r Level
B. Portable UJtet/feuclcet Discliarae
Rgure 11. Discharge measurements
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6. Revision No. 2, March 1994, Page 33 of 38
6.6 EQUIPMENT LIST
For Physical Habitat Profile Measurements:
1. Surveyors Rod (Metric, Fiberglass, 3-part Telescoping, approx 5m extended about 1.5 m
collapsed) - for width and depth meas's. (about $80)
2. Clinometer (SUUNTO Model PM5/360PC or equivalent) must read both in percent and degrees
slope — for gradient meas's (about $105) .
3. Spherical (Convex) Canopy Densiometer. (Lemmon, Model 8 —only one manufacturer: Robert
Lemmon, Forest Densiometers 5733 S.E. Cornell Dr., Bartlesville, Oklahoma 74006, phone
918-333-2830) (about $80).
4. Tripod (modest price camera tripod, light, telescoping) for backshe on previous cross-section to
obtain gradient (tripod necessary only if Physical habitat work is done by one person aloneV
(about $10-$30)
5. Bearing Compass (Backpacking type, e.g., Silva Ranger, as used in NSS) (About $40)
6. Colored plastic Flagging (2 colors)
7. 'Rebar* to monument site, sheet aluminum label and aluminum nails for to nail into tree.
8. 2 Covered Clipboards (lightweight, with strap or lanyard to hank around neck - Camera straps work
great for straps.)
9. Pencils, soft lead (good "fat lead* mechanical, disposable or whatever works best - need lots of
extras)
10. Chest Waders (1 each person) - with felt-soled boots for safety and speed if waders are the
neoprene 'stocking* type.
11. Hip Wadere (1 aaeh person) ~ with felt soles for safety and speed on slippery rocks.
12. Camera, waterproof 35mm with standard and wide angle lense (small point and shoot would
probably be better as well as cheaper). (About $1004250)
13. Elm (35mm, color slide film, ASA 400 and 100) Will use 10 to 20 frames per stream reach.
14- Fiberolas^ Tape & reel (SO m metric), with good hand crank and handle for measuring cross-
sections. (About $30-$45)
15. Meter stiqfr (cheap wooden ruler with metric calibrations) for bank angle meas's.
16. Hip Chain for measuring of reach and valley transects. (Optional)
Note: The surveyors rod, clinometer, bearing handheld compass, fiberglass tape, and hip chain can be
bought from either Forestry Suppliers or Ben Meadows Company. All other Hems can be bought from
department or sporting goods stores.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 2, March 1994, Page 34 of 38
For Discharge Measurement :
1. Electromagnetic Water Velocity Meter (Mgfc Marsh-McBimey Mod 201D or similar) or Inrmeller-Type
Water Velocity Meter (Mgfc Swoffer Model 2100-14), with adjustable wading rod (metric
calibrations).
2. Portable Weir - 60 degree V-noteh (this b a plastic plate to impound and measure flow in streams
too small for the Velocity-Area Method).
3. Bucket. Plastic with volume graduations - for timed bucket discharge measurements.
4. Stopwatch -1 prefer mechanical over electronic -mechanical ones more reliable in wet situations.
(Used for timed volume discharge measurements.)
Note: The Weirs and Water Velocity meters can be bought from suppliers like Ben Meadows or Forestry
Suppliers. The Meters can also be bought directly from manufacturer.
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EMAP-SW-Streams, Physical Habitat Assessment, Section 6, Revision No. 3, April 1995. Page 35 of 36
6.7 LITERATURE CITED
Bain, M.B., J.T. Finn, and H.E. Booke. 1985. Quantifying stream substrate for habitat analysis studies.
Nor. Amer. Jour, of Fish. Man. 5:499-500.
Frissell, C.A., W.J. Uss, C.E. Warren, and M.D. Hurley. 1986. A hierarchical framework for stream habitat
classification: viewing streams in a watershed contest Environ. Mgmt 10 (2): 199-214.
Kaufmann, P.R. (ed.) 1993. Physical Habitat. Pages 59-69 in R.M. Hughes, ed. Stream Indicator and Design
Workshop. EPA/600/R-93/138. U.S. Environmental Protection Agency, Corvallis, Oregon.
Lemmon, P.E. 1957. A new instrument for measuring forest overstory density. J. For. 55(9):667-669.
Unsley, R.K., M.A. Kohler, and J.LH. Paulhus. 1982. Hydrology for engineers. McGraw-Hill Book Co. N.Y.,
N.Y. 508 p.
Mulvey, M., L Cato, and R. Hafele. 1992. Oregon nonpoint source monitoring protocols stream
bloassessment field manual: for macroinvertebrates and habitat assessment. Oregon Department
of Environmental Quality Laboratory Biomonitoring Section. 1712 S.W. 11th Ave. Portland,
Oregon, 97201. 40 p.
Plafkin, J.L, M.T. Barbour, K.D. Porter, S.K. Gross, R.M. Hughes. 1989. Rapid bioassessment protocols
for use in streams and rivers: benthic macroinvertebrates and fish. EPA/440/4-89/001. U.S.
Environmental Protection Agency, Assessment and Watershed Protection Division, Washington,
DC.
Platts, W.S., W.F. Megahan, and G.W. Minshall. 1983. Methods for evaluating stream, riparian, and biotic
conditions. USDA For. Serv., Gen. Tech. Rep. INT-183,71 p.
Robison, E.G. and R.L Beschta. 1990. Characteristics of coarse woody debris for several coastal streams
of southeast Alaska. USA. 47(9): 1684-1693.
Robison, E.G. and P.R. Kaufmann. 1994. Evaluating two objective techniques to define pools in small
streams, pgs 659-668, Jn R.A. Marston and VA Hasfurther (eds.) Effects of Human Induced
changes on hydrologlc systems. Summer Symposium proceedings, American Water Resources
Association,. June 26-29,1994, Jackson Hole, Wyoming. 1182 pp.
Stack, B.R. 1989. Factors influencing pool morphology in Oregon coastal streams. M.S. Thesis, Oregon
State University. 109 p.
Wolman, M.G. 1954. A method of sampling coarse river-bed material. Trans. Am. Geophys. Union
35(6):951-956.
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EMAP-SW-Streams, Physical habitat Assessment, Section 6. Revision No. 2, March 1994, Page 36 of 38
6.8 APPENDIX: FIELD FORMS FOR PHYSICAL HABITAT PROTOCOL
-------
:,; j: .:;...;. ; ; > CHANNEL/RIPARIAN CROSS SECTION & THALWEG PROFILE FORM,;:,:r^vJsi'.ii^^
STREAM NAME:
STREAM 0:
8UMI IBACXMTEt
1. 8m
-lo6..;L
LFT •',..
.CHI ..]::.- •..;•;.:
CTH "'.I' ->i:
RCT* ,'•', . : .
ROT ;-;L:o.-
I9TRATEMEA1
X.XXM
S
•
DATE OF VISIT: / / 94
TEAM O |drd«|: 12346878
« S^/SloiMi.Wkit);:-.;-;::- _
DETTH
XXX CM
^ " .
SacciAss
. CODE
:
EMtED.
0-100% FWO
. SUBSTRATE SIZE CLASS COOES
RS • •EDROCK (SMOOTH) •(UfcOEft IM* A CAR)
RR « KDROCK IROUOM) MUMMER THAN A CAR)
BL - MXADHt |260 TO 4000 MM) •(•ASXETBAlL TO CAR)
CB * eomt |S4 TO 260 MM) •(TEN** BAU TO MWETMUi
QC - COARSE ORAVEL 118 TO 84 MUl *(MAMttJ TO TtNMS lAUl
OF - tMEOM
SA - MNP (0
FN - ftr/cu
WD -WOOD
OT v OTHBI I
LOC. B
-FT
*°T 1 L
kVa 12 TO 18 MM|*(lAQYBUa TO MARSIE)
).08To2MM|*|o«Tn*upTouDv*uq$aE) -•
Y.MUCK (NOT
ANY sac)
COMMENT)
wnrvi
^
r. % BEARJNQ (BAdtsnc): ::; | •
In. FISH
COVER/
OTHER
,»_.»_
MACMTNVm
lAAMIOVOBMB
> o.i M BIOI
< 0.3 M (SMALL!
OVOMAHOMO VBO.
< 1 M V KJ*FACS
UHOBKUT BANK*
BOWBOW
STWCTUMB
0-AMtNT
1- MOOBMTC (10-46*1
3- HlAVT |40.7«*t
4» Vnv HfAW t> 70*t
CHAMNtU
Fuo
... i. BANK MEASUREMENTS .^ - . .- ..;;. ._ .y;,. Y-;',
ANK ANOLE
0-360
•
3AMCFUU. WIDTH • .v
NOSED HT. = ' ••:; J(v
SANKFUiHT. :
WETTED WOT*
^•.RWBTH
| . . : . .- .
'&D^&'&&&
m
m
m
m
m
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-------
Section 6, Physical Habitat Assessment. Page 38 of 38
Remidecl l*j (initial).
THALWEG PROFILE & WOODY DEBRIS FORM-STREAMS
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-------
EMAP-SW-Streams, Periphyton-Field, Section 7, Revision No. 2, March 1994, Page 1 of 3
SECTION 7
STREAM PERIPHYTON: FIELD PROCEDURES
7.1 SAMPLING RATIONALE
7.1.1 Periphyton are algae, fungi, bacteria, protozoa, and associated organic matter associated with channel
substrates. Periphyton are useful indicators of environmental condition because they respond rapidly and
are sensitive to a number of anthropogenic disturbances, including habitat destruction, contamination by
nutrients, metals, herbicides, hydrocarbons, and acids.
7.2 FIELD EQUIPMENT
1. Large funnel (15-20 cm diameter).
2. Scrape area delimiter (3.8 cm diameter pipe, 3 cm tall).
3. Stiff-bristle toothbrush with handle bent at 90° angle.
4. Wash bottle.
5. Collection bottle to catch removed periphyton.
6. 60 mL syringes with 3/8* hole bored into the end.
7. 50 mL centrifuge tubes or similar sample vials.
8. Formalin.
9. Glass-fiber filters (0.45 urn average pore size) for chlorophyll Ł.
10. Pre-ieached, pre-ashed, weighed glass-fiber filters (0.45 urn average pore size) in numbered
pans for ash free dry mass (AFDM).
11. Forceps for filter handling.
12. Millipore-type filtration apparatus with stainless steel filter base, and Nalgene funnel and suction
flask.
13. Nalgene Hand-operated vacuum pump (need one additional pump as a backup).
14. Aluminum foO.
15. Ice chest
16. Dry ice or other means of freezing sample.
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EMAP-SW-Streams, Periphyton-FJeld, Section 7, Revision No. 2, March 1994, Page 2 of 3
7.3 FIELD PROTOCOLS
7.3.1 Collection
1. Periphyton samples wfll be collected using a random-systematic procedure. The location (left
middle, or right 1/3 of the channel) of the first sample (Transect B) will be chosen randomly.
Subsequent samples (Transects C-J) wfll be collected sequentially from the left, middle, then right
1/3 of the channel, resulting In three samples from each side and middle.
2. Periphyton are collected, using the appropriate method, from flowing (riffles) and slack water
(pools) habitats.
3. Rock and wood samples which are small enough (< 15 cm diameter) and can be easily removed
from the stream are collected by placing the substrate in a funnel which drains into a sample bottle.
A defined area of substrate surface (12 cm2)is enclosed, and attached periphyton is dislodged with
30 seconds of brushing with a stiff-bristled toothbrush. Care must be taken to ensure that the
upper surface of the rock is the surface that is being scraped. .
4. Loosened periphyton is then washed, using stream water from a wash bottle, from the substrate
into the 500 mL sample bottle.
5. Soft-sediments are collected by vacuuming the upper 1 cm of sediments confined within the 12
cm2 sampling ring into a 60 mL syringe.
6. All samples, regardless of substrate type, are composited by habitat (riffle or pool) and mixed
thoroughly.
7. Record total volume of composited sample before proceeding to the next
stepl
8. Four subsamples wfll be taken from each composite sample. These are:
a. Identification /Enumeration
1) Withdraw 50 mL of mixed sample and place in a labelled sample vial
(50 mL centrifuge tubes work wetl). Cover label with dear tape.
2) Preserved sample with 2 mL of 10% formalin. Gloves should be worn.
3) Tightly cap tube and tape with electrical tape.
b. Chlorophyll a
1) Withdraw 25 mL of mixed sample and filter onto a glass-fiber filter
(0.45 um pore size) using a hand-operated vacuum pump. (Note: for
soft-sediment samples, allow grit to settle before withdrawing sample).
-------
EMAP-SW-Streams, Periphyton-Reld, Section 7, Revision No. 2, March 1994, Page 3 of 3
2) Fold filter so that the sample on the filter surface is folded together,
wrap in aluminum foil, and affix the tracking label to the outside, and seal
with clear tape.
3) Freeze filter as soon as possible by placing it in a freezer.
4) Store frozen for laboratory analysis.
c. Ash Free Dry Mass (AFDM)
1) Withdraw 25 ml of mixed sample and filter onto a pre-leached, pre-
weighed glass-fiber filter. (Note: for soft-sediment samples, allow grit
to settle before withdrawing sample).
2) Do not fold this filter. Return filter to it's numbered container, wrap in
aluminum foil, and affix tracking label to outside, and seal with dear tape.
3) Freeze filter as soon as possible by placing f in a freezer.
4) Store frozen for laboratory for analysis.
d. Alkaline/Acid Phosohatase
1) Withdraw 50 ml of mixed sample and place in a labelled sample vial
(50 ml centrifuge tubes work well). Cover label with clear tape.
2) Tightly cap tube and tape with electrical tap a.
3) Freeze sample as soon as possible by placing it on dry ice.
4) Store frozen for laboratory analysis.
NOTE: ALL SAMPLES MUST BE CAREFULLY LABELLED WITH THE APPROPRIATE
BAR-CODE AND ADHESIVE TAGS!
7.4 SHIP SAMPLES TO:
Dr. James M. Lazorchak
U.S. EPA
3411 Church Street
Cincinnati, OH 45244
Phone: (513) 569-7076
Fax: (513) 569-7078
-------
EMAP-SW-Streams, Benthlc Metabolism, Reid Methods, Section 8, Rev. No. 2, March 1994, Page 1 of 4
SECTIONS
BENTHIC (SEDIMENT) METABOLISM: FIELD METHODS
1. SCOPE AND APPLICATION
1.1 Ecosystems are complex, self-regulating, functional units defined by rates and processes, such as
energy flow or material cycling. Functional indicators are those metrics that measure energy flow and
material transformation within the ecosystem.
1.2 The method outlined here is designed for headwater to mid-order streams, though ft may be adapted
for larger rivers or lakes.
2, SUMMARY OF METHOD
2.1 The method measures changes in dissolved oxygen (DO) concentrations of the overlying water within
microcosms containing small amounts (ca 10 mL) of sediments as a means of assessing benthic microbial
community activity. Sediments are collected from depositional habitats along a study reach defined by 40X
channel width. Following Incubation, DO Is remeasured and the sediments saved for ash free dry mass
(AFDM) analysis. Respiration, the decline In DO within each microcosm, is adjusted for AFDM, yielding a
measure of community respiration /g AFDM. Organic carbon turnover time can be calculated from the
empirical relationship between organic carbon (estimated as AFDM X 0.5) concentration of the sediment
and oxygen consumption.
3. DEFINITIONS
3.1 AFDM - ash free dry mass
3.2 DO - dissolved oxygen
4. INTERFERENCES
4.1 Conditions which may interfere with this method are unknown. The method has been performed
successfully in waters containing high metal concentrations, high conductivity, and moderate salinity.
5. SAFETY
5.1 Gloves should be worn when collecting, compositing, mixing, and subsampling sediments.
6. EQUIPMENT AND SUPPLIES
6.1 Ice chest for incubating centrifuge tubes
6.2 1 L Nalgene beaker to support centrifuge tubes during incubation
6.3 Scoop sampler for sediments.
6.4 50 mL, screw-top, centrifuge tubes.
-------
EMAP-SW-Streams, Benthic Metabolism, Reid Methods, Section 8, Rev. No. 2. March 1994, Page 2 of 4
6.5 YSI Model 58 Dissolved Oxygen meter with Model 5905 Stirring BOD probe.
6.6 Spare batteries for D.O. meter.
6.7 Permanent markers for labelling tubes.
6.8 Sample labels and field data sheets.
6.9 Zip-lock sandwich bags to contain replicates from each treatment and site.
6.10 Ice chest with dry ice for sample freezing (or other means of freezing samples).
7. REAGENTS AND STANDARDS
None.
8. SAMPLE COLLECTION, PRESERVATION, AND STORAGE
8.1 Collect sediment samples by gently scooping the top 4 cm of soft, surface sediments from depositional
habitats (pools, eddys, backwaters) within the study reach. Samples are collected near Transects B-J, as
defined by the Physical Habitat reach delineation. If soft sediments are abundant collect samples at or near
each transect If soft sediments are scarce, collect them wherever you can within the reach. A total of 3
L of sediments are needed for metabolism and sediment toxicity analyses.
8.2 Fill ice chest 2/3 fun with stream water and record temperature and dissolved oxygen (D.O.).
8.3 Thoroughly mix composited sediments.
9. QUALITY CONTROL
9.1 All field work conducted during this study will be done by personnel having prior experience with the
designated methods, or under the direct supervision of experienced personnel.
9.2 Sediment metabolism wOI be sampled at each site on all site visits.
9.3 Quality Assurance Objectives:
9.3.1 Precision-Precision of measurements is assured by carefully following the method protocols,
including calibration of DO meter.
9.3.2 Completeness-Valid data are required from 80% of the sites visited.
9.3.3 Representativeness-Sediment metabolism is being determined for depositional areas of streams.
Replicate samples from these areas in a stream segment (defined by fish survey) are composited and mixed
to provide a representative site sample.
9.4 Documentation and Review: Sediment metabolism field data are recorded and checked for
completeness and accuracy before leaving the site. All data sheets are inspected for completeness.
-------
EMAP-SW-Streams, Benthic Metabolism, Reid Methods, Section 8, Rev. No. 2, March 1994, Page 3 of 4
accuracy, and legibility before proceeding to the next sample. Raw data sheets are retained in a file, and
the data is entered into an ASCII file.
10. CALIBRATION AND STANDARDIZATION
10.1 Before the field season, meters will be calibrated against the WinWer titration method of DO
determination.
10.2 Dissolved Oxygen Meter Calibration (for YSI model 58, with YSI model 5905 stirring BOD probe)
10.3 Each day, before leaving lab, motel, or base station:
10.3.1 Check meters batteries to ensure that meter and stirring probe are operational.
10.3.2 Check probe membrane to ensure that it is not frayed or torn and that there are no bubbles under
the membrane. If membrane is not intact or has entrapped bubbles, It should be replaced according to the
manufacturer's directions.
10.4 Upon arriving at each she:
10.4.1 Zero meter according to manufacturer's directions.
10.4.2 Calibrate meter using the water-saturated atmosphere method described in the meter's operations
manual.
11. PROCEDURES
11.1 Remove any visible organisms from the sediment before placing approximately 10 mL of sediment
in each of 5 labelled, 50 mL screw-top centrifuge tubes. Labels are covered with clear tape.
11.2 Pill each tube to the top (no head space) with stream water from the ice chest and seal.
11.3 Pill 1 additional tube with stream water only and seal.
11.4 Incubate in closed ice chest for 2 hours.
11.5 Measure D.O. in each tube.
11.6 Respiration is the difference between initial and final D.O.
11.7 Decant overlying water and save sediment.
11.8 Seal tubes and place on dry ice as soon as possible.
11.9 Store frozen for laboratory analysis.
11.10 If you are collecting samples for sediment toxicitv tests, save remaining composite sediment sample
by double-bagging in labelled, zip-lock freezer storage bags. Label is covered with clear plastic tape.
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EMAP-SW-Streams, Benthic Metabolism, Reid Methods, Section 8, Rev. No. 2, March 1994, Page 4 of 4
11.11 Store sediment toxicity sample chilled (but not frozen!) for laboratory analysis.
11.12 NOTE: All samples must be carefully labelled with the appropriate bar-code and adhesive tags.
Tags need to be covered with dear tape.
11.13 Ship samples to: Dr. James M. Lazprchak
US EPA-EMSL
3411 Church Street
Cincinnati, OH 45244
Phone 513/569-7076
Fax 513/569-7078
12. DATA ANALYSIS AND CALIBRATION
N/A
13. METHOD PERFORMANCE
N/A
14. POLLUTION PREVENTION
t-
N/A
15. WASTE MANAGEMENT
1.5.1 For normal respiration measurements, no waste Is produced. Water may simply be decanted into
the stream from which it came.
16. REFERENCES
N/A
17. TABLES, DIAGRAMS, FLOWCHARTS, AND VALIDATION DATA
None.
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EMAP-SW-Streams, Benthlc Macroinverte. Reid Meth., Sec. 9, Rev. No. 2, March 1994, Page 1 of 9
SECTION 9
BENTHIC INVERTEBRATE FIELD METHODS
9.1 INTRODUCTION
9.1.1 Rationale
9.1.1.1 This protocol is intended for evaluating the biological integrity of wadeable streams of EPA
Region III of the United States for the purpose of detecting stresses on community structure and
assessing the relative seventy of these stresses. It is based on the Rapid Bioassessment Protocol III -
Benthic Macroinvertebrates (RBP) published by the U.S. Environmental Protection Agency (Plafkin et al.,
1989) and adopted for use by many states. The two man kick net procedure of the RBP is replaced in
this protocol with a kick net modified for use by one person.
9.1.2 Importance of Monitoring Benthic Assemblages
9.1.2.1 Benthic invertebrates inhabit the sediment or live on the bottom substrates of streams. The
benthic macroinvertebrate assemblages in streams reflect overall biological integrity of the benthic
community such that monitoring these assemblages are useful In assessing the status of the water body
and monitoring trends. Benthic communities respond to a wide array of stressors in different ways so
that it Is often possible to determine the type of stress that has affected a macroinvertebrate community
(Klemm et al., 1990). Because many macroinvertebrates have relatively long life cycles of a year or
more and are relatively immobile, macroinvertebrate community structure Is a function of past
conditions.
9.1.3 Sampling Design
9.1.3.1 The protocols described here employ a randomized, systematic spatial sampling design that
minimizes bias in positioning the sampling stations. Sampling stations are randomly located along the
transects established according to the Physical Habitat Assessment Protocol. Eleven transects will be
systematically marked off equal distance apart In proportion to stream size. Macroinvertebrate
assemblages wfll be collected at the nine interior transects at either the l/4th (left), 1/2 (middle), or 3/4
(right) points on each transect as determined by the throw of a die. If the die comes up a one or two
the sample wfll be collected at the 1/4 point, a three or four would indicate the 1/2 point, and a five or
six would indicate the 3/4 point Left and right are determined when facing downstream.
9.1.4 Components of the Macroinvertebrate Indicator Protocols
9.1.4.1 There are two Macroinvertebrate Indicator Protocols. The BASE Protocol wfll be utfiized at all
EMAP-SW and REMAP Sites. It consists of the collection of nine samples with the 595 urn modified kick
net and combining the riffle samples in one bucket and the pool samples in another bucket The
combined riffle samples become the composite riffle sample and the combined pool samples become
the composite pool sample. The SORTING-IDENTIFICATION Protocol describes the procedure that
must be followed during the sorting and Identification of the samples (see the Laboratory Manual).
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EMAP-SW-Streams, Benthlc Macroinverte. Reid Meth., Sec. 9, Rev. No. 2, March 1994, Page 2 of 9
9.1.5 Objectives of the Study
9.1.5.1 The primary objective of the 1994 EMAP-SW pilot study Is to determine if nine modified kick net
samples separated into a composited riffle sample and a composited pool sample will adequately
characterize a stream segment In 1st and 2nd order streams throughout EPA Region III. A second
objective is to determine the relative Importance of the riffle and the pool sample in assessing stream
condition.
9.2 PROCEDURES FOR FIELD CREWS
9.2.1 Preparing for the Sampling Day
9.2.1.1 Before leaving the Base Site for the day's sampling site(s). Check the conductivity meter and
00 meter for proper operation and be sure the batteries are well charged. Go over the equipment
checklist (Table 1) to be sure all equipment is loaded in the vehicle and in good working condition.
Place one gallon of ethand per site in a cooler which has vermiculite on the bottom. Check maps for
all sampling sites to be visited. DO NOT place barcodes on the jars before going into the field because
doiny *o causes tracking problems if any labeled jars are not used at the site they were prepared for.
9.2.2 Upon Arrival at the X-sfte.
9.2^.1 Record the time, weather conditions, and initial observations in the field note book and fill in the
Information requested at the top of the Checklist (Table 2). While the habitat assessment team marks off
the 11 transects, the benthos crew wOl do the water chemistry following the Protocol for Water
Chemistry. By the time the water chemistry measurements are completed and the water chemistry
samples are collected, the habitat assessment crew should have the downstream transects marked off
so that the benthos crew can begin to collect the benthic invertebrate samples.
9.2.3 BASE Protocol for Collection of the Macroinvertebrate Composite Samples
9.2.3.1 Carry benthos sampling gear to the first sampling station. The first sampling station is transect
B (the first transect upstream from the downstream end of the study segment and wQI be marked by flag
bearing the letter B). If the transects have not been established by the physical habitat crew, the
benthos crew w9l need to set up the transects following the Physical Habitat Protocols.
9.2.3.2 From a random numbers table or roll of a die pick a number (if not already done by the Physical
Habitat Crew) between one and three to determine if the sample wQI be taken at the 1/4 (left side), 1/2
(middle), or 3/4 (right side) point of the transect (1 or 2 - 1/4, 3 or 4 - 1/2,5 or 6 - 3/4). After
randomly picking sampling point at the first transect systematically sample from left middle, right for the
remaining eight transects. In streams only two nets wide pick left or right or if the stream is only one
net wide collect entire stream width. Be sure to cirde R. C. or L on field forms for each transect
sampled.
9.2.3.3 Determine if the stream at this station consists of a riffle, run, or pool and collect the sample
following the protocol for that habitat (See Subsection 9.2.3.4 • Sampling Protocol for Riffles and Runs or
Subsection 9.2.3.5 - Sampling Protocol for Pools). For the purpose of this section any area where there
is not sufficient current to collect a sample by the riffle protocol, would be defined as a glide-pool and
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EMAP-SW-Streams. Benthic Macroinverte. Field Meth., Sec. 9, Rev. No. 2, March 1994, Page 3 of 9
sampled according to the protocol for pools. If it is impossible to sample the station with the modified
kick net following either of these protocols, spend about 60 seconds hand picking a sample from about
0.5 m2 of substrate at the station and combine it with the other samples in either the "riffle" or 'pool'
bucket. Repeat the procedure to collect a sample from each of the nine established interior transects
(transects B-J). M of the riffle samples will be combined in one bucket and the pool samples will be
combined in another bucket.
9.2.3.4 Procedures for Collecting Macroinvertebrates from Riffles and Runs with the Modified Kick Net
Sampler
9.2.3.4.1 A modified Kick Net with 595 urn mesh openings and 0.5 m nylon bag (Wfldco # 425-C50) will
be used to collect macroinvertebrates in riffles and runs. Begin with the most downstream station so as
not to disturb the organisms upstream of the sampling location. This procedure requires only one person
and is the preferred macroinvertebrate collecting method for streams with flowing water (a second
person is often used to keep time and record information on the field forms).
9.2.3.4.2 The following steps are used to collect the sample from riffles and runs:
1. Attach the four foot pole to the sampler. Care should be exercised to be sure the handle is
on tight or the net might become twisted in strong current causing the loss of part of the
sample.
2. Position the sampler quii/cl/ and securely on the stream bottom so as to eliminate gaps
under the frame with the net opening upstream. Large rocks that prevent the sampler from
seating properly should be avoided.
3. Hold the sampler in position on the substrate while checking for heavy organisms, such as
mussels and snails in an area of about 0.5 m2 in front of the net These organisms will have to
be hand picked and placed in the net.
4. Hold the sampler securely while kicking the substrate vigorously for 20 seconds (Use
stopwatch) in an area of about 0.5 m2 in front of the net
5. After 20 seconds sampling at the station (transect), hold the net in place with the knees and
pick up any loose rocks in the sampled aiea and rub off organisms in front of the net Also any
additional mussels and snails found should be placed in the net Remove the net from the water
with a quick upstream motion to wash the organisms to the bottom of the net
6. Rinse net contents into the •riffle" bucket about half full of water (about one or two gallons)
by inverting the net in the water.
7. Inspect the net for dinging organisms. With forceps remove any organisms found and place
them in the bucket.
8. Urge objects (rocks, sticks, leaves, etc.) in the bucket should be carefully inspected, and
any organisms found should be washed into the bucket before discarding. Remove as much
detritus as possible without losing any organisms.
-------
EMAP-SW-Streams, Benthic Macroinverte. Reid Math., Sec. 9, Rev. No. 2, March 1994. Page 4 of 9
9. If there is too tittle water to collect the sample with the kick net randomly pick up 10 rocks
from the riffle and pick and wash the organisms off them into the riffle bucket
10. Combine all the riffle samples In the same'riffle* bucket
11. After all nine stations are sampled and the riffle samples combined in the 'riffle* bucket
obtain a composite riffle sample as described in Subsection 9.2.3.6.
9.2.3.5 Procedures for Collecting Macroinvertebrates from Pods with the Modified Kick Net Sampler
9.2.3.5.1 A modified Kick Net Sampler with 595 pm mesh openings and a 0.5 m nylon bag attached to
a four foot pole is used for collecting macroinvertebrates from pools. This sampler, described as a
modified kick net is available from WQdco (Catalog number 425-C50).
9.2.3.5.2 The following steps are used to collect the sample from pools and slow water areas:
1. Attach the four-foot pole to the Kick Net sampler. Care should be exercised to be sure the
handle is on tight or the net might become twisted while dragging It through the water causing
the loss of part of the sample.
2. If the pool Is too deep (much over one meter) to safely sample at the designated spot move
downstream tfll a safe sampling spot is found.
3. Inspect the stream bottom for any heavy organisms, such as mussels and snails. In the 0.5
m2 area to be sampled. These organisms will have to be hand picked and placed Into the net
or bucket
4. While disturbing about 0.5 m2 of the substrate by kicking vigorously with the feet collect a
20 second sample by dragging the net repeatedly through the disturbed area just above the
bottom while continuing to kick. Keep moving the net all the time so that the organisms trapped
in the net win not escape,
5. After 20 seconds, hold the sampler between the legs with the net partially submerged and
pick up any loose rocks in the sampled area and rub or brush any organisms found on them
into the net Also recheck the area for any additional snails or dams and place them in the net
6. Rinse net contents into the 'pool* bucket about half filled with water (about one or two
gallons) by inverting the net In the water.
7. Inspect the net for dinging organisms. With forceps remove any organisms found and place
them in the bucket
8. Large objects in the bucket should be carefully inspected and any organisms found must be
washed into the bucket before discarding the objects. Remove as much detritus as possible
without losing any organisms.
-------
EMAP-SW-Streams, Benthlc Macroinverte. Field Meth., Sec. 9, Rev. No. 3, April 1995, Page 5 of 9
9. If there is too little water to use the kick net, stir up the substrate with the gloved hands and
use the U.S. 30 sieve to collect the organisms from the water in the same way the net is used in
larger pools.
10. Combine all the pool samples in the same 'pool' bucket
11. After all nine stations are sampled and the pool samples are combined in the 'pool* bucket,
obtain a composite pool sample as described In Subsection 9.2.3.6.
9.2.3.6 Procedures for Obtaining a Composite Sample
9.2.3.6.1 Nine transects wUI be collected from all the sites. All the riffle samples from each site will be
combined and all the pool samples will be combined in the field and a composite sample of each obtained
as follows:
1. Save the entire combined sample as the composite sample for that habitat (riffle or pool) at
that site and record on the sample collection form the number of samples that make up the
composite sample.
2. Pour the entire contents of the bucket through a U.S. Standard 30 sieve. Remove any large
objects and wash off any clinging organisms onto the sieve before discarding.
3. Using a squirt bottle, rinse all the organisms from the bucket onto the sieve. This is the
composite sample for that habitat (riffle or pool) for the site.
4. Estimate the total volume of the sample and determine how large a jar will be needed for the
sample (quart, half gallon, or gallon). Do not use more than one jar for each of the samples
unless it cannot be avoided.
5. Wash the contents of the sieve to one side by gently agitating the sieve in the water and
wash the sample into a jar using as little water from the squirt bottle as possible Oar should not
be over 1/4 full of water). If the jar is too full pour off some water through the sieve until the
jar is not more than 1/4 full or use a second jar if a larger one is not available. Carefully
examine the sieve for any remaining organisms and with forceps place them in the sample jar.
6. Place a waterproof label with the following Information in the jar.
1. Stream Number
2. Type of sampler and mesh size used
3. Habitat type (riffle or pod)
4. Name of stream
5. Date of collection
6. Collectors initials
7. Number of transect samples composited
7. Replace the cap and with grease pencil put site number and sample type (Riffle or Pool) on
the cap. If two jars are used be sure to mark them as such.
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EMAP-SW-Streams. Benthlc Macroinverte. Reid Meth.. Sec. 9, Rev. No. 2, March 1994, Page 6 of 9
9.2.4 After Collection of benthlc Invertebrate Sample(s)
9.2.4.1 Return samples and the sampling gear to the vehide.
9.2.4.2 Assist the rest of the field crew with the fish collection.
•
9.2.5 Upon Returning to Vehide
9.2.5.1 Each sample to assigned a unique prenumbered barcoded label to be used for tracking
purposes. This unique barcode number must be placed on the jar and the label containing the barcode
number must be filled in with a waterproof pen or pencfl. This unique number must appear on all
data sheets and must not be removed from the jar except as described in the Sample Sorting and
identification Protocol (See Benthlc Invertebrate Laboratory Manual). If more than one jar Is used for the
sample this number must appear on each jar (use the extra labels and write the number on the label).
DO NOT use two different Barcode Numbers on two jars containing one single sample. Either at the
vehide or at the motel, enter the barcoded numbers into the computer.
9.2.5.2 AH samples are preserved In ethand as follows:
a. If jar to more than 1/4 fuD of water, pour off enough to bring It to less than 1/4 full
using the U.S. Standard No. 30 sieve so as not to lose any organisms.
b. FBI Jar nearly fuU with 95% ethand so that the concentration of ethanol is 70%. If
there is a small amount of water in the sample, it may not be necessary to fill the jar
entirely full to reach a 70% concentration. It is very Important that suffident ethanol be
used to reach 70% concentration.
c. Transfer any organisms on the sieve back Into the jar with forceps.
9.2.5.3 Check to be sure that the prenumbered stick-on barcoded label Is on the jar and cover the
entire label with dear, waterproof tape. Check to be sure the waterproof label Is in the jar and properly
filled in as described In the Protocol for Obtaining the Composite Sample (Subsection 9.2.3.7). Be sure
the Inside label and outside label describe the same sample. Replace the cap and seal It with electrical
tape. Check to make sure the cap is properly marked with site number and habitat type (pool or riffle).
CQPV Barcoded Sample ID Number from the sample far onto the Sample Collection Form-Streams and
indicate which Is the pool sample and which Is the riffle sample.
9.2.5.4 Complete the check off sheet (Table 2) and enter any additional pertinent Information in the field
note book. Race the samples In a cooler or other secure container for transporting. Check to see that
ail equipment to In the vehide.
9.2.6 After Each Week of Sampling
9.2.6.1 All samples must be transported or shipped to the designated depot site at the end of each
week of sampling or at other agreed upon intervals. Tracking numbers of all samples transferred to the
depot must be entered into the computer. A printout of the sample tracking numbers (barcodes) of the
samples shipped must be induded with the shipment and a copy will be mailed to the Indicator Lead
along with the computer diskette each week.
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EMAP-SW-Streams, Benthic Macroinverte. Field Meth., Sec. 9, Rev. No. 2, March 1994, Page 7 of 9
9.2.6.2 When the shipment arrives at the depot the tracking numbers on the jars will be checked with
the numbers on the printout. Any discrepancies will be cleared up by contacting the field crew
responsible for the shipment as soon as possible. The samples will be picked up by Region III biologists
and transferred to EMSL • Cincinnati at the end of the sampling period. All samples will be checked
when they arrive at Cincinnati to be sure all samples listed on the shipping label are in the shipment.
The samples will be shipped to the designated taxonomist(s) as soon as possible after they arrive at
Cincinnati so that the analyses can be completed and the data analyzed within the required time frame.
A copy of the computer printout listing all samples shipped will be included with the shipment and a
copy will be sent to the Indicator Lead at EMSL-Cincinnatl.
9.3 REFERENCES
Klemm, D.J., P.A. Lewis, F. Fulk, J.M. Lazorchak. 1990. Macroinvertebrate field and laboratory methods
for evaluating the biological integrity of surface waters. EPA/600/4-90/030. U.S.
Environmental Protection Agency, Environmental Monitoring Systems Laboratory, Cincinnati, OH
45268.
Plafkni, J.u, M.T. Barbour, K.D. Porter, S.K. Gross, R.M. Hughes. •.'.•89. Rapid bioassessment protocols
for use in streams and rivers: Benthic macroinvertebrates and fish. EPA/440/4-89/001. U.S.
Environmental Protection Agency, Assessment and Watershed Protection Division, Washington,
DC. 20460.
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EMAP-SW-Streams, Benthic Macroinverte. Reid Meth.. Sec. 9. Rev. No. 2, March 1994, Page 8 of 9
TABLE 1. EQUIPMENT UST FOR MACROINVERTEBRATE SAMPLING CREWS
1. Modified Kick Net with 595 pm mesh openings and dosed bag (WUdco # 425-C50)
2. Spare nets for the Kick Net Sampler or extra sampler
3. Sample jars, one quart, one per site, US Plastic #66165
4. Sample jars, half gallon, one per site
5. Sample jars, one gallon, one per site
6. Maps of sampling sites
7. Labels, waterproof, 6 per site
8. First aid kit
9. Rubber gloves, heavy rubber, two pair
10. Rain gear for each person
11. Pencils, No. 2 soft lead, a dozen
12. Grease pencBs, two
13. Knife, pocket with at least two blades
14. Scissors, one pair
15. Forceps, two pair
16. Clear waterproof Tape
17. Electrical tape, four rolls
18. Labels, prenumbered barcoded. stick-on type, 6 per site
19. Compass • -
20. Wash bottle, one liter capacity
21. Sieve, U.S. Standard 30
22. Two Buckets, plastic, eight to ten quart capacity
23. Sieve bucket, 595 pm mesh openings (optional)
24. Kim wipes in small ziplock bag
25. Field check list sheets
26. Ethanol. one gallon per site. In proper container
27. Funnel, with large spout
28. Waders, hip length, pair for each person
29. Computer for sample tracking
30. Cooler for transporting ethanol and samples
31. Watch with timer or a stop watch
32. Pole attachment for Kick Net Sampler, four foot length
33. Camera and film
34. Small spatula
35. Reid note book • pocket size
36. Dice for random numbers determination
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EMAP-SW-Streams, Benthic Macroinverte. Field Meth., Sec. 9. Rev. No. 2, March 1994, Page 9 of 9
TABLE 2. MACROINVERTEBRATE SAMPLING PROTOCOL CHECKLIST FOR BASE PROTOCOL
Date: Time: Site Number.
Name of Stream: Location:
Crew ID (Circle): A B C D E F G; Collector:
1. Weather and time recorded on Habitat Assessment Data Sheet.
2. Initial observations in field notebook.
3. Composite riffle sample collected with label inside.
4. Composite pool sample collected with label Inside.
5. Correct barcode and label on all jars and sealed with clear, waterproof taoe.
_ 6. All samples preserved.
, 7. Caps sealed with tape.
. 8. Samples logged In computer.
.. 9. Sample jars in cooler or otherwise secured.
10. All equipment accounted for and secured in vehicle.
Signature: Time:
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APPENDIX
I. SAMPLE COLLECTION AND SHIPMENT FOR SEDIMENT TOXICITY SAMPLES
-------
SAMPLE COLLECTION AND SHIPMENT FOR SEDIMENT TOXICITY SAMPLES
SEDIMENT COLLECTION AND SHIPMENT
1. Use the sediment left over from the benthic (sediment) metabolism indicator in Section 8, Benthic
(Sediment) Metabolism: Field Methods.
2. Mix sediment well with a stainless steel or plastic mixing spoon.
3. Fill a 1 gallon ziplock bag with at least 1 L of sediment.
4. Close bag. making sure seal Is secure.
5. Fill out ID label; place label and sediment inside another ziplock bag. Seal.
6. Place these bags inside a half gallon wide-mouth, screw top plastic jar. Place a label with stream ID on
outside of the jar. Make sure cap seals tight.
7. Hold sediments samples on ice (do not freeze!) for laboratory analysis.
8. Ship samples to: DynCorp, C/0 U.S. EPA
3411 Church St.
Cincinnati, OH 45244
Phone (513) 569-7095
FAX (513) 569-7081
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APPENDIX H, FISH TISSUE, REVISION NO. 2, MARCH 1994, PAGE 2 OF 2
SELECTING FISH TISSUE SPECIMENS - 1994 EMAP-SW STREAMS PILOT
B) if fewer than 5 fish of any size are available, you may serxl as few as 3 fish that are at or at least
near the minimum desired size (120 mm).
C) If an acceptable secondary target species sample (by the above criteria) is not available send only
tt,e Primary target species sample. If neither a primary or secondary spedes sample that meets these
criteria is available, use your best judgement in sending some type of fish sample (may be mbced
species).
SPECIMENS
• Keep hands, work surfaces, and wrapping materials dean and free of potential contaminants (mud.
fuel, formalin, sun screen. Insect repeUant etc.)
• Measure total welaht of Individuals for primary target spedes and count the total number of
XS^ tang* (TU o^s«»nd«y target sped.. Individual Record ail of
this information on the Fish Tissue Sample Tracking Form.
. Write the ba/^e number^)*** on the Fish Tissue Senipli Traddng Fern Make sure that the form
to filed out completely.
Individually. Once wrapped, place eac sampe n k or garbage bag.
• Expel excess air and seal the bag(s). Wrap dear tape ejourid the bag(s) to seal and make a
surface for each sample label.
lab*. will somrtmn « * met. should alwy» b. a total on Hit tear tag.
• Place tabeledzlphx* or oari»g.b.gts) Wo .eiond ftortc tag and «>l and toM second bag(s)
(repeat previous two steps).
• Place doubie^agged sample(s) In coder wtth dry tee unfl attpmem.
PROBLEMS?. QUESTIONS? - CM1: Roger Y-nJey, »13) S6&.7093 (* ta. vdcema. also)
-------
APPENDIX
H. SELECTING FISH TISSUE SPECIMENS
-------
APPENDIX H, FISH TISSUE, REVISION NO. 2. MARCH 1994, PAGE 1 OF 2
SELECTING FISH TISSUE SPECIMENS - 1994 EMAP-SW STREAMS PILOT
if OOSSibte. obtain one sample each, containing the desired weight or number (see beJow)-of similarly
sized individuals*, from the primary and secondary target species lists (2 composite samples total):
I, PC'MARY TARGET SPECIES
Small adult fish DESIRED
fin priority ordert WEIGHT
1 ) Blacknose Oace 50** - 400 g
2) Another Dace species 50** - 400 g
3) Creek Chub or Failflsh 50** - 400 g
4) Slimy ScuJpln/Mottied Scuipin 50** - 400 g
5) Stoneroller 50** - 400 g
6) A Darter species 50** - 400 g
7) A Shiner species 50** - 400 g
A) Choose the highest priority target species from the abovellst thalhas at least enough HjndMduato
to attain the minimum weight (50 g). Get as much weight of flsh as posstte wtthln the desdred weight
range (50^00 g). Use scale provided to determine weight With dean hands, place the fish In fresh
aluminum fofl (dull side towards flsh) before placing fish in weighing container.
(B) If fewer than the desired number of Individuals of aja primary target species are collected, send
Individuals of a small nontarget spedes if 50 g or more are available.
• . Getting a sufficient sample amount Is a higher priority than
weigw represents the minimum amount needed for laboratory «**
minimum weight if more fish are present They should send a. many flsh a. possible up to 400 g
weight goal.
]| SECONDARY TABngT SPECIES
Collect and save a sample of secondary target spedes if such a sample of desired rmmber of Individuals
of desired size Is available. Collect similar sized IndMduato if enough are present
*
Larger adult flsh DESIRED DESIRED
fln priority ordert SSZ HUMSEfl
1) White sucker > 120 mm 5
2)Hogsucker > 120 mm 5
3) A Bass spedes >120mm 5
4) A Trout spedes >120 mm 5
5) A Sunfish spedes >l20mm 5
6) Carp > 120 mm 5
^^
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APPENDIX G, FISH, REVISION N0.3, APRIL 1995, PAGE 2 OF 3
Code Categories
N NONE
D Deformities
E Eroded fins
L Lesions
T Tumors
F Fungus
B Black spot disease
I 'Ion"
A Anchor worm
W Leeches
X Exopthalmia"
M Missing
0 Other
FIELD PROTOCOLS FOR FISH COLLECTION
EXTERNAL ANOMALIES
No anomalies present.
Skeletal anomalies of the head, spine, and body shape.
Reductions of fin surface area or hemorrhage of fin rays.
Open sores or exposed tissue.
Irregular cell growth which are firm and cannot be easily broken.
Filamentous or "fuzzy" growth on the fins or body.
Parasite that appears as small black cysts on the fins and body.
Appears as white spots on the fins and body.
Parasitic infection characterized by a worm embedded in the flesh of the
fish.
Worms which have anterior and posterior suckers. They may attach
anywhere on the body.
Popeye disease" Is an anomaly seen as the bulging of the eye.
Eyes missing, blindness or deterioration of the lens should also be noted.
Anomalies or parasites not specified.
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APPENDIX G, RSH. REVISION N0.3, APRIL 1995, PAGE 3 OF 3
FIELD PROTOCOLS FOR RSH COLLECTION
VOUCHERING FISH SPECIES AND UNIDENTIFIED SPECIMENS
Category 1. Large species, difficult to identify, OR uncommon In region. Preserve 1 -2 adults plus 2-5 YOY and
juveniles. Document with photograph.
American Eel White Sucker Shads
Sturgeon Longnose Sucker Bullhead catfish
Paddlefish Hogsucker Channel catfish
Gars ' Quillback Esoclds
Bowfin Carpsuckers Aforonespp.
Mooneye and Goldeye Moxosfomaspp. Drum
Carp Buffalo fishes Salmonids
Walleye and Sauger M/cropferusspp. Grapples
Category 2. Small fish OR difficult to identify species. Preserve up to 25 adults, juveniles and YOY per site.
Lampreys Troutperch Sculpins Madtoms
Cyprinids Chubsucker Sunfish Sticklebacks
Darters Topminnows Silversides
a. Place subsample of 25 individuals of total catch into a kill jar containing 10% buffered formalin
solution.
b. When specimens are dead, transfer to a nylon bag containing a waterproof label with site ID,
barcode, date, crew ID and species name.
c. Repeat process for all species identified.
d. If fewer than 25 individuals are collected, preserve entire sample.
e. Place all ziplock bags in labelled voucher jar.
Category 3. State and federal species listed as protected. Photograph and release. Voucher if specimens
have died.
NEW YORK VIRGINIA (All SC) Grass pickerel
Mooneye Spotfinchub Redsidedace
Round whitefish Yellowfin madtom Satinfin shiner
Silver chub Orangefin madtom Popeye shiner
Black redhorse (SC) Roanoke logperch New River shiner
Longear sunfish ' Kanawha minnow
Eastern sand darter WEST VIRGINIA (ALL SC) Cheat minnow
Bluebreast darter Mountain brook lamprey Longnose sucker
Longhead darter Shovelnose sturgeon Mountain madtom
Gilt darter Paddlefish
Mooneye MARYLAND
Potomac sculpin
PENNSYLVANIA Slimy sculpin Striped Bass
Eastern sand darter Shield darter Maryland darter
Crystal darter
-------
APPENDIX
G. FIELD PROTOCOLS FOR FISH COLLECTION
-------
APPENDIX G. FISH. REVISION N0.3. APRIL 1995, PAGE 1 OF 3
FIELD PROTOCOLS FOR FISH COLLECTION
1. Site Selection
a. Determine channel width.
b. Survey sample reach.
c. Determine if reach requires block nets.
d. If conductivity is below 10 uS or if flow, depth or turbidity make it unsafe to electrofish, crew may
elect to use seine only or not sample. THIS IS A SAFETY DECISION.
e. In case of emergency, determine location of means of easy egress from stream.
2. Electrofishing
a. Set unit to 300VA and pulsed DC. Select initial voltage setting. Start generator, set timer, and
depress switch to begin fishing.
b. Fish in an upstream direction, parallel to the current. Adjust voltage and waveform output
according to sampling effectiveness and mortality fish specimens.
c. With switch depressed, sweep electrodes from side to side in the water. Sample available cut-bank
and snag habitat as well as riffles and pools.
d. Netters follow operator and net fish. Deposit fish in buckets. Block with seines in riffles, pools and
snags.
e. Continue for 40 channel widths. Record total time spent collecting and shocking time on data
sheets.
f. Identify and release any threatened and endangered species.
g. Identify and measure (SL, TL, BD) sport fish and very large specimens, record external anomalies,
and release unharmed.
h. Identify other specimens. Determine number of Individuals in species, measure largest and
smallest Individuals, and voucher as described in Voucher Protocol.
I. Retain a subsample of target species for Fish Tissue Contaminants analysis.
3. Seining will be used in conjunction with electrofishlng and in sites where stream is too deep for
electrofishing to be conducted safely.
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APPENDIX F, BENTHIC INVERTEBRATES, REVISION NO. 2, MARCH 1994, PAGE 2 OF 3
TABLE III. PROCEDURES FOR POOLS USING THE MODIFIED KICK NET SAMPLER
1. Attach four-foot pole to the sampler.
2. Inspect about 1/2 square meter of bottom for any heavy organisms, such as mussels and snails.
which have to be hand picked and placed in the net
3 While disturbing about 1/2 square meter of substrate by kicking, collect a 20 second sample by
dragging the net repeatedly through the area being disturbed. Keep moving the net all the time so that
the organisms trapped in the net will not escape.
4. After 20 seconds remove the net from the water with a quick upstream motion to wash the organisms
to the bottom of the net.
5. Rinse net contents into a small bucket of water (about one or two gallons) by Inverting the net In the
water.
6.lnspect the net for clinging organisms. With forceps remove any organisms found and place them in
the bucket
7. Urge objects in the bucket should be carefully inspected for organisms which are washed into the
bucket before discarding.
8. Combine this sample with the other pod samples In the -pool' bucket
9. After all stations are sampled and all pod samples are combined together In the -pooT bucket obtain
a composite sample as described In Table IV.
TABLE IV. PROCEDURE FOR OBTAINING THE COMPOSITE SAMPLE
,. Pour the contents of the riffle bucket throug|i. US.•"*"» «Ł;JSJ* *" bttl"1''
rinsing « well to be sure all organisms are washed from th» bucket onto the sieve.
wash contents of the sieve to one side by gently agteHno. In watsr and wash Into jar using as IMe
ItXn?ZS&ŁŁT«%Sslble. (MMy ~mln. the s^v, tor any remaining onjanism, and
place them in the jar.
3. Place property filed out waterproof label In the jar and replace the cap.
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APPENDIX F, BENTHIC INVERTEBRATES, REVISION NO. 2, MARCH 1994, PAGE 3 OF 3
TABLE V. SAMPLE PRESERVING AND LABELING
1. Fill in special prenumbered bar coded label and place on jar. All additional jars used for a sample
must be labeled with same number. Enter this number which wfll be used for tracking purposes in the
computer.
2. Preserve samples in ethanol as follows: a. If jar Is more than 1/4 fun of water, pour off enough to
bring it to less than 1 /4 full using proper sieve to retain organisms, b. FBI jar nearly full with 95%
ethanol so that the concentration of ethanol Is 70%. If there is a small amount of water, in the sample, it
may not be necessary to fill the jar entirely full to reach a 70% concentration, c. Transfer any organisms
on the sieve back into the jar with forceps.
3. Check to be sure waterproof label Is in jar with the required information on it
4. Check to be sure that the prenumbered stick-on bar coded label Is the on jar and
agrees with the inside label. Cover the entire label with dear, waterproof tape.
5. Seal the caps with electrical tape.
a Complete the check off sheet and place samples in cooler or other secure
container for transport.
7. Secure ail equipment in the vehicle.
TABLE VI. MACROINVERTEBRATE SAMPLING PROTOCOL CHECKLIST
Date: Time: Site No.:
Stream Name & Location:
Crew ID: 1 234587 8; Collector
_ i. Habitat Assessment Sheet Completed with weather conditions described.
_ 2. Initial observations in field notebook.
_ 3. Composite riffle sample collected with waterproof labet Inside.
_ 4. Composite pool sample collected with waterproof label Inside.
_ 5. Bar code and label on ail jars covered with dear waterproof tape.
__ 6. All samples preserved.
__ 7. Caps sealed with tape.
_ 8. Photos of site.
9. Samples logged In computer.
__ 10. Sample jars stored for travel.
__ 11. All equipment accounted for and secured in vehlde.
Time sampling completed
-------
APPENDIX
F. BENTHIC INVERTEBRATES
-------
APPENDIX F, BENTHIC INVERTEBRATES, REVISION NO. 2, MARCH 1994, PAGE 1 OF 3
TABLE I. BASE PROTOCOLS FOR COLLECTING MACROINVERTEBRATES
1. Do the water chemistry.
2. Locate first sampling station (second flag) from downstream end of the study segment and roll die to pick
left (1), middle (2), or right side (3) of transect to sample. If stream narrower than three nets, pick left or
right. If wide enough for only one net, then sample entire stream width. After first transect systematically
sample remaining transects left, middle, or right so that three samples are collected on left, middle, and right
at the site.
3. If riffle or run use protocol in Table II. If pool use protocol In Table III or hand pick for 60 seconds if kick
net cannot be used.
4. Go to next upstream station and repeat Combine all riffle samples in one bucket and pool samples in
another. Check net after each sample for dinging organisms and transfer to bucket
5. After a sample is collected from each of nine interior transects and ail samples are combined In the proper
bucket obtain a composite sample as described in Table IV.
6. Assist with the fish collection.
7. Preserve and label each sample as described In Table V.
TABLE II. PROCEDURES FOR RIFFLES AND RUNS USING KICK NET SAMPLER
1. Attach four foot pole to the sampler.
2. Position sampler quickly and securely on stream bottom with net opening upstream.
3 Hold the sampler in position on the substrate while checking for snails and dams in an area of about 0.5
square meter in front of the net; kick the substrate vigorously for about 20 seconds in front of the net
4. Inspect and rub off with the hands any organisms dinging to the rocks, especially those covered with
algae or other debris,
5. Remove the net from the water with a quick upstream motion to wash the organisms to the bottom of net
6. Rinse net contents Into the 'riffle' bucket containing one or two gallons of water by Inverting the net In
the water.
T.lnspect the net for dinging organisms. With forceps remove any organisms found and place them into
the bucket
8. Large objects (rocks, sticks, leaves, etc.) in the bucket should be carefully inspected for organisms
before discarding.
9. Combine all riffle samples in the "riffle" bucket
10. After all stations are sampled and all riffle samples combined in the 'riffle' bucket obtain a composite
sample as described in Table IV.
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APPENDIX E, BENTHIC (SEDIMENT) METABOLISM, REVISION NO. 2, MARCH 1994, PAGE 2 OF 2
SUMMARY FIELD PROTOCOLS FOR BENTHIC (SEDIMENT) METABOLISM
1.3 Ship samples to: Dr. James M. Lazorchak
US EPA-EMSL
3411 Church Street
Cincinnati, OH 45244
Phone 513/569-7076
Fax 513/569-7078
-------
APPENDIX
E. SUMMARY FIELD PROTOCOLS FOR BENTHIC (SEDIMENT) METABOLISM
-------
APPENDIX E, BENTHIC (SEDIMENT) METABOLISM, REVISION NO. 2, MARCH 1994, PAGE 1 OF 2
SUMMARY HELD PROTOCOLS FOR BENTHIC (SEDIMENT) METABOUSM
1.1 Dissolved Oxygen Meter Calibration (for YSI model 58, with YSI model 5905 stirring BOD probe)
1 Each day, before leaving lab, motel, or base station: a) check meter's batteries to ensure that meter
and stirring probe are operational and b) check probe membrane to ensure that It Is not frayed or torn and
that there are no bubbles under the membrane. If membrane is not Intact or has entrapped bubbles, it
should be replaced according to the manufacturer's directions.
2. Upon arriving at each site: a) zero meter according to manufacturer's directions and b) calibrate meter
using the water-saturated atmosphere method described In the meter's operations manual.
1.2 Sediment Collection and Experimental Set-up
1 Collect sediment samples by gently scooping the top 4 cm of soft, surface sediments from depositions
habitats (pools, eddys. backwaters) within the study reach. Samples are collected near Transects B.J. as
defined by the Physical Habitat reach delineation. If soft sediments are abundant, coUect samples at or near
each transect If soft sediments are scarce, collect them wherever you can within the reach. A total of 3
L of sediments are needed for metabolism and sediment toxidty analyses.
2. FB ice chest 2/3 full with stream water and record temperature and dissolved oxygen (D.O.).
3. Thoroughly mbc composited sediments.
4. Remove any visible organisms from the sediment before pacing approdmatelytOmL of sediment In each
of 5 labelled. 50 mL screw-top centrifuge tubes. Labels are covered with dear tape.
5. Fill each tube to the top (no head space) with stream water from the ice chest and seal.
a F3l 1 additional tube with stream water only and seal
7. incubate in dosed Ice chest for 2 hours.
8. Measure 0.0. in each tube.
9. Decant overlying water and save sediment
10. Seal tubes and freeze as soon as possible.
11. Store frozen for laboratory analysis.
13. Store sediment toxicity sample chilled (but not frozen!) (or laboratory analysis.
' NOTE AH samples must be carefully, labelled with the appropriate barcode and adhesive tags. Tagsneed
to be covered with dear tape.
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APPENDIX D, PERIPHYTON, REVISION NO. 2. MARCH 1994. PAGE 2 OF 2
SUMMARY FIELD PROTOCOLS FOR PEHIPHYTON COLLECTION
2) Fold filter so that the sample on the filter surface is folded together
wrap in aluminum foil, and affix the tracking label to the outside, and seal
with dear tape.
3) Freere fiUer as soon as possible by placing It In a freezer.
4) Store frozen for laboratory analysis.
C. Ach Free Drv M??s fAFDMl -
1) Withdraw 25 mL of mixed sample and fitter onto a pleached, pre-
wdgnedtfass-ffcer filter. (Note: for soft-sedlment samples, allow gnt
to settle before withdrawing sample).
9\ nft not fold this filter. Return fitter to It's numbered container, wrap in
'aluminum !ol a^al tracking lab* to outsKe. and seal with dear tap,
3) Freeze filter as soon as possible by placing It In a freezer.
4) Store frozen for laboratory for analysis.
d. Albino/Acid Phosnhatase
2) Tightly cap tube and tape with electrical tape.
3) Freeze sample as soon as possible by placing it on dry ice,
4) Store frozen for laboratory analysis.
NOTE:ALLSAMPLESMUSTBECAREFUU.YUBELLE0WmTHEAPPROPR.ATE
BAR-CODE AND ADHESIVE TAGS!
1.2 Ship Samples To:
Dr. James M. Lazorchak
U.S. EPA
3411 Church Street
Cincinnati, OH 45244
Phone: (513) 569-7076
Fax: (513) 569-7078
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APPENDIX
D. SUMMARY FIELD PROTOCOLS FOR PERIPHYTON
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APPENDIX D, PERIPHYTON, REVISION NO. 2, MARCH 1994, PAGE 1 OF 2
SUMMARY HELD PROTOCOLS FOR PERIPHYTON COLLECTION
1.1 Collection
1. Periphyton samples will be collected using a random-systematic procedure. The location (left, middle.
or right 1 /3 of the channel) of the first sample (Transect B) will be chosen randomly. Subsequent samples
(Transects C-J) will be collected sequentially from the left middle, then right 1/3 of the channel, resulting
in three samples from each side and middle.
2. Periphyton are collected, using the appropriate method, from flowing (riffles) and slack water (pools)
habitats.
3. Rock and wood samples which are small enough (< 15 cm diameter) and can be easily removed from
the stream are collected by placing the substrate in a funnel which drains into a sample bottie. A defined
area of suostrate surface (12 cirr^ls enclosed, and attached periphyton Is dislodged with 30 seconds of
brushing with a stHf-brisfled toothbrush. Care must ba taken to ensure that the upper surface of the rock
is the surface that Is being scraped.
4. Loosened periphyton is then washed, using stream water from a wash bottle, from the substrate into the
500 mL sample bottle.
5. Soft-sediments are collected by vacuuming the upper 1 cm of sediments confined within the 12 cm2
sampling ring into a 60 mL syringe.
6. All samples, regardless of substrate type, arc- composited by habitat (riffle or pool) and mfaced thoroughly.
7. Record total volume of composited sample before proceeding to the next step!
8. Four subsamples wffl be taken from each composite sample. These are:
a. Identification /Enumeration
«i^ —
1) Withdraw 50 mL of mixed sample and place In a labelled sample vial
(50 mL centrifuge tubes work well). Cover label with dear tape.
2) Preserved sample with 2 mL of 10% formalin. Gloves should be worn.
3) Tightly cap tube and tape with electrical tape.
b. Chloronnvll a
1) Withdraw 25 mL of mixed sample and filter onto a glass-fiber fBter
(0 45 urn pore size) using a hand-operated vacuum pump. (Note: for
soft-sediment samples, allow grit to settle before withdrawing sample).
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APPENDIX C, PHYSICAL HABITAT, REVISION NO. 2, MARCH 1994, PAGE 6 OF 7
FIELD SUMMARY: HABITAT CLASSIFICATION PICTURES
Rgure2. Sketches
of channel unte (From FriaaeU etaL,1986)
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APPENDIX C, PHYSICAL HABITAT, REVISION NO. 2, MARCH 1994, PAGE 7 OF 7
HELD SUMMARY: P-HAB PROBLEM AREAS
Bars: dry at baseflow, inundated at bankfuil flow.
Measure wetted width across and over bars, but record bar width In the column provided on thaiweg profile
or cross-section form.
Islands: as high as the surrounding flood plain; dry even at bankfuil flow. .
Measure only the width of the main channel between island and shore In the measures; then if required.
measure the side channel width separately (record on another form). Handle the side channels created by
Islands as follows:
* visually estimate the percent of flow in the side channel
• If < 15% - Note side channel with ^SC" In side channel column.
* If 15-35% - Note *SC* plus detafled channel & riparian Cross Sections on the side channel. Label
additional new Cross Section Transect Forma as "XA", *XB*. etc. corresponding to nearest main
channel Transect location.
* If >35% - Note *SC" plus detafled channel & riparian profiles PLUS another thaiweg profile on the
side channel. As above, label additional new Cross Section Transact and Thaiweg Profile Forms
as -XA-, -XB-, etc. corresponding to nearest main charms! Transact location.
When side channels are present, write "SCT In the span provided on the Thaiweg Profile form. The thaiweg
form should reflect their presence with a continuous "SO" note until channels converge. If It Is a long side
channel, a continuing arrow can be used to mark It In the comments column to save time.
and intermittent Streams, where no water Is In thi charmafc
Record zeros for depth and wetted width*
Record habitat type as dry channel fDR").
Make ail Channel Cross-Section measures across the unvegetated portion of the channel.
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APPENDIX C, PHYSICAL HABITAT, REVISION NO. 2, MARCH 1994, PAGE 4 OF 7
FIELD SUMMARIES: SUBSTRATE and WOODY DEBRIS SIZE CLASSES
Substrate size class and embeddedness are estimated, and depth is measured for 5 particles taken @ 5
equally-spaced points on each cross-section. The cross-sectlon is defined by laying the surveyor's rod or
tape to span the wetted channel.
ffllHSTBATE SIZE CLASSES:
RS Bedrock (Smooth) > 4000 mm - smocth surface rock or hardpan (bigger than a car)
RR Bedrock Rough >4000 mm — (bigger than a car)
BL Boulders > 250 to 4000 mm - (basketball to car size)
CB Cobbles 64 to 250 mm - (tennisball to basketball)
GC Gravel(Coarse) 16 to 64 mm — (marble to tennisball)
GF Gravel (Fine) 2 to 16 mm — Oadybug to marble)
SA Sand .06 to 2 mm - ( 0.1 m (2.4 In)
— — Length Ł.i.Sm(2. 5 ft) -count only part with
TWQ Tallvs:
above MM *» <*• is
will eventually fall into channel).
Categories for Tally (12 potential combinations):
Diameter (large end): 0.1 to <0.3 m SSiw
0.3 to <0.6 m . (1 to 2 It)
0.6to<0.8m (2 to 2,6
>0.8m (>2.6tt)
>ism (>*««)
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APPENDIX C, PHYSICAL HABITAT, REVISION NO. 2, MARCH 1994, PAGE 5 OF 7
HELD SUMMARY: HABITAT CLASSIFICATION AT CHANNEL UNIT SCALE
Table 2. Habitat Classification at Channel Unit Scale (See figure 2). Note that in order for a channel
habitat unit to be distinguished, it must be at least as wide or long as the channel Is wide.
Class Coda
Pools:
Plunge Pool PP
Trench Pool PT
Lateral Scour Pool PL
Backwater Pod PB
Oam Pool PO
Glide GL
Riffle RI
Rapid RA
Cascade CA
Fails FA
Dry Channel OR
Description
Still water, low velocity, smooth, glassy surface, usually deep
compared to other parts of the channel:
Pool at base of plunging cascade or falls.
Pool like trench in stream center.
Pod scoured along bank.
Pool separated from rain flow off side of channel.
Pool formed by impoundment above dam or constriction.
Water moving stowiy, win smooth, unbmtmn nuHsc* - tow
turbulence
Water moviig, win smafl npplea, waves and eddies - waves no*
breaking, surface tension not broken, sound: •babbfing',
•gurgling*.
WfltflT IIM*"""*"* "T"* n"*4 •*«*"**"*. «•»*««• "&** intermittent
YihrtHwgtBfwflh breaking waves - sound: Continuous noshing,
but not as loud as cascade.
Water umwnant rapid and very turbulent over steep channel
bottom. lytastdwstBf surface broken in short Irregular plunges.
mostly whaewater- sound: Roaring.
Free faffing water over vertical or near vertical drop into plunge,
water turbulent and whto over Wgh falls, sound: from splash to
roar, depending upon dbcharga.
No water in channel
Code
N
W
R
B
F
WRRW.RBW
O
Pool-Forming Bement Category
Not Applicable, Hat** Unit is not a pool
Large Woody Debris.
Rootwed
Boulder or Bedrock
Unknown cause (unseen fluvial processes)
Combinations
Other - note in comments
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APPENDIX C, PHYSICAL HABITAT, REVISION NO. 2, MARCH 1994, PAGE 2 OF 7
FIELD SUMMARY: COMPONENTS OF P-HAB PROTOCOL
Width, Depth Profile, Hab Classes, Woody Debris:
At 10 (15) equally spaced Intervals between each of 11 channel cross-sections (100 or 150 along
entire reach):
- Measure max. depth (Thaiweg') and wetted width.
- Classify habitat and pool-forming elements.
- Determine presence of soft/fine sediment at thalweg measurement points.
Between each of the channel cross sections, tally all Large Woody debris within and above the
bankfull channel according to size class. In the tally boxes P^* ™ ** form- "ru*e se^
tally for LWD wholly or partially within the bankfull channel and then for LWD only bndgmg above
the channel.
NOTE: If initial width estimate is <2.5 m, then 150 thalwefl injaff^8""*^ ?^° ?±^n^I
TTso m reach. If width is 2.5 to 3.5 m, then make 100 thalweg measurements at i j m Intervals. In all
other Sses 100 measurements are made at an interval 1/IOOtti the length ofthe sample reach. II the
chann^widih is initSy estimated greater than 4.0 m (and the reach length therefore exceeds 150 m), then
omit the multiple width measures at the 100 thalweg Intervals.
Channel and Riparian Cross-Sections:
Measurements- Bankfull tooth, oankfull height, Indston height, wetted width, bar width, undercut
tenka^eTiih^nd clinometer); gradient (dlnometer), sinuosity (compass backste), ripanan
canopy cover (densftometer).
Discharge:
and
In succession.
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APPENDIX C, PHYSICAL HABITAT, REVISION NO. 2, MARCH 1994, PAGE 3 OF 7
RELO SUMMARY: RIP. VEG., HUMAN DISTURB., IN-CHANNEL COVER:
• Observations upstream 5 meters and downstream 5 meters from each of the 11 cross-section stations.
* For Rip. Veg. and Human Disturbances, include the visible area from the stream back a distance of 10 m
(30 ft) shoreward from both the left and right banks. If the wetted channel Is split by a mid-channel bar, the
bank and riparian measurements shall be for each side of the channel, not the bar.
* Three vegetation layers:
CANOPY LAYER (>5m high)
UNDERSTORY (0.5 to 5m high)
GROUND COVER layer (<0Ł high).
• Canopy and Understory Vegetation Types:
(Qeciduous. Coniferous, Mtod, or^one) In each of the two taller layers (Canopy and Understory). "Mixed*
if more than 10% of the area! coverage made up of the alternate type.
* Area! Cover Scores for Vegetation and In-Channel Coven
0: (absent - zero cover)
1: (sparse - cover <10%)
2: (moderate - cover 10-40%)
3: (heavy - cover 40-75%)
4: (very heavy - cover >75%).
• Scoring Human Disturbances:
B: The human activity or structure is ON THE STREAMBANK
C: CLOSE to the Bank (within 10m)
P: PRESENT, but farther than 10m from the bank.
0: NOT PRESENT.
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APPENDIX
C. PHYSICAL HABITAT
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APPENDIX C, PHYSICAL HABITAT, REVISION NO. 2, MARCH 1994, PAGE 1 OF 7
HELD SUMMARY: P-HAB LAYOUT AND WORKFLOW
1. Habitat Sampling Layout:
Thaiweo interval: 1.0 m for streams <2.5 m wide (from initial estimate).
1.5 m for streams 2.5 - 3.5 m wide
0.01 (reach length) for streams >3.5 m wide
100 thaiweg meas' increments in each sample reach, except 150 in streams <2Łm wide
Channel /Riparian Cross Section Transect every 10th thalweg interval (every 15lh for channels <2.5m wide).
Eleven of them, marked "A" thru "K".
2. Work Row:
* At downstream start point (Cross-Section 'A"), one person makes channel dimension, substrate, bank, and
canopy densitometer measurements. Second person records those measurements whfle making visual
estimates of riparian vegetation structure, fish cover, and human disturbance. No bearing or slope at first
cross section.
• Proceed upstream between Cross Sections A and B, making measures at each Thalweg distance interval.
One person in channel measures width (when required), thalweg depth, and determines presence of
fine/soft sediment at thalweg. The other person records those measurements, classifies channel habitat
and makes large woody debris estimates.
• When you complete 10 thalweg Intervals and reach one of 11 pro-marked cross section flags, stop and
take out a new cross-section form for Cross Section V. Repeat all the Channel/RIpanan measurements
at this new location. In addition, do the dope & bearing backsftes together. lrt^^ete*9 <
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APPENDIX B, WATER CHEMISTRY, REVISION NO. 2, MARCH 1984, PAGE 2 OF 2
SUMMARY OF SITE PROCEDURE FOR WATER CHEMISTRY
COLLECT WATER SAMPLE
A. Make sure cubitainers and syringes are labelled and have the same barcode ID.
B. Rinse the 500 mL sample beaker three times wfth streamwater from mid-stream.
C. Rinse cubitainer three times wtth 25-50 mL of streamwater using sample beaker.
D. Fill cubitainer wtth streamwater using the 500 mL sample beaker. Cap the cubitainer and
make sure that the seal Is tight.
DO NOT EXPAND CUBITAINER BY BLOWING IN IT.
E. Rinse each of the four. 60 mL syringes three times wtth 10-20 mL of streamwater.
F. Rl each of the four syringes with streamwater frorn mid-stream by slowly pulling out the
plunger. If any air gets into the syringe, discard the sample and draw another.
G Invert the syringe (tip up) and cap the syringe wtth a syringe varve. Open the valve, tap the
&
transport container
and wriund wtth +6 one
I!. IN SITU MEASUREMENTS
A. Conductivity
1 Measure and record the conductivity of the QCCaohJtan.
i Mewure stream conductivity In sample beaker or In orescent part of stream.
B. Dissolved Oxygen/Temperature
1. Calibrate DO nr>eter foUc)wlng nwter Irnlnictions.
2. Measure DO/Temperature In mid-stream.
-------
APPENDIX
B. WATER CHEMISTRY
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APPENDIX B, WATER CHEMISTRY, REVISION NO. 2, MARCH 1994, PAGE 1 OF 2
WATER CHEMISTRY SAMPUNG CHECKLIST
I. EQUIPMENT TO CARRY IN FIELD FOR WATER CHEMISTRY
Rinse/Test 125 mL bottles of QCC solution in zipiock bag
Conductivity Pen
Reid Forms
One. 250 mL plastic beakers
One, 500 mL plastic beaker in dean zipiock bag
One cubitalner in dean zipiock bag (barcode label attached)
Four, 60 mL syrinnes in plastic container (each one with bar code label attached)
Four syringe valves in plastic container
Opaque garbage bag
II. EXTRA EQUIPMENT TO CARRY IN VEHICLE
Cooler with 4-6 one gallon zipiock bags fined with ice
Back-up labels, forms, syringe valves
III. DAILY ACTIVITIES AFTER SAMPLING
1. Enter chemistry tracking data from today's sample (bar code 10, stream ID, date, etc.) into computer.
2. Check that cubitainer lid is on tight and has a flush seal.
3. Prepare sample for shipping (label and seal cooler, replace ice as dose as possible to shipping time).
4. Call Overnight shipping company to arrange pick-up of cooler.
5. Rinse all sample beakers with deionized water, three times.
6. Make sure conductivity pens are rinsed with deionized water and are stored with moist electrodes.
7. Barcode label the next days sample containers (cubitainer and syringes), pack cubitainer and sample
beakers In dean zipiock bag. and pack four syringes and syringe valves In plastic container.
-------
APPENDIX
A. SUMMARY OF INITIAL SITE PROTOCOLS
-------
APPENDIX A, INITIAL SITE PROTOCOLS, REVISION NO. 2, MARCH 1994, PAGE 1 OF 1
SUMMARY OF INITIAL SITE PROTOCOLS
1. Find the stream location in the field corresponding to the "X* on 7.5* topo map (X-site). Crews
should use all available means to insure that they are at the correct site, as marked on the
map. including; 124.000 USGS map orienteering, topographic landmarks, county road maps.
and -global positioning system (GPS) confirmation of site latitude and longitude.
2. Classify the site AT THE X-SITE. as:
NONTARGET No Stream Channel
Impounded stream
Marsh/Wetland
Unwadeable Stream (> 50% of reach is unwadeable)
TARGET Regular Stream
Intermittent Stream
Dry Channel
Altered Channel (stream channel different from map representation)
INACCESSIBLE Physical Barriers (physically unable to reach the X-site)
No Permission
Record class on Site Verification form, do not sample Nontarget or Inaccessible sites. Take
samples from Target sites as discussed in field operations manual
3. Measure the stream width at three typical" places within 10 m of the X-sfte. Average and
round the width to the nearest meter. Record width on the stream site verification form.
Layout a sample reach with a length of 40 times the stream width. If the stream is less than 4
m wide, use 150 m as the sample reach length.
4. Do a reconnaissance of the sample reach.
5. Proceed downstream half the required reach length, measure the distance with a tape
measure down the middle of the stream.
6. Rnd the dosest habitat break to the downstream reach endpoint, mark it as the reach start
point (transect A). If no habitat break is present within 3-4 channel widths of the endpoint,
mark the endpoint as Transect A and block net tt.
7. Proceed upstream marking 10 more sample transects (sftea B-K) at 1/10 Intervals along the
calculated reach length (every 4 channel widths or 1JS meters in small streams). Assign each
transect a Left, Center or Right sample site by throwing a die.
8. For the last transect (K), end the sample reach at the habitat break nearest the measured 40
channel width endpoint If no habitat break is present for 3-4 channel widths, end the reach at
the endpoint.
NOTE: If there is a lake/pond or a stream order change (100,000 map based) along the survey
reach, end the sample reach at the barrier. Make up for the loss of stream length by adding
length to the other end of the reach ('slide' the reach). Locations where the stream order
changes will be noted on the topo maps provided to the field crews. Do not 'slide' the reach to
.WHT-^, c—rr" f «m,-'n»*«n' ,«tn.ictUT0* culverts and the like.
-------
APPENDICES
SUMMARY OF FIELD PROTOCOLS FOR EMAP INDICATORS
A. SUMMARY OF INITIAL SITE PROTOCOLS
B. WATER CHEMISTRY
C. PHYSICAL HABITAT
D. SUMMARY FIELD PROTOCOLS FOR PERIPHYTON
E. SUMMARY FIELD PROTOCOLS FOR BENTHIC (SEDIMENT) METABOLISM
F. BENTHIC INVERTEBRATES
G. FIELD PROTOCOLS FOR FISH COLLECTION
H. SELECTING FISH TISSUE SPECIMENS
I. SAMPLE COLLECTION AND SHIPMENT FOR SEDIMENT TOXIOTY SAMPLES
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EMAP-SW-Streams, Fish Indicator, Field Methods, Section 10. Revision No. 2, March 1994, Page 1 of 14
SECTION 10
FISH INDICATOR
10.1 EXPERIMENTAL DESIGN
10.1.1 The EMAP Surface Water Program will continue Its stream pDot activities in the mid-Atlantic
region during FY 94. Research will be conducted in conjunction with USEPA's TIME and Region 3
REMAP programs. One hundred ninety-nine Appalachian Ridge and Valley, Blue Ridge, Central
Appalachian Plateau, Piedmont and Atlantic Coastal Plain sites will be selected for study. These sites
will be sampled by teams of USEPA, LJSFWS, state and contract personnel In each of the operational
units of the EMAP, REMAP, and TIME projects. The streams will be sampled during the 12 week Index
period. The objective is to collect a representative sample of the fish assemblage by methods designed
to 1) collect all except very rare species in the assemblage and 2) provide a measure of the relative
abundance of species in the assemblage.
10.1.2 Data will be summarized (means, standard deviations, etc.), analyzed for correlations among
biotic and physico-chemical data (both parametric and distribution-free), and tested for site differences
using analysis of variance. In addition, multivariate analyses of the data wfll be compared to metrics
(e.g., diversity, evenness, relative abundance, IBI) to determine which approach yields the most
Information about the status of stream ecosystems. Cumulative distribution frequencies of community
metrics will be evaluated to determine success of sampling method in obtaining a representative
qualitative sample of the fish assemblage. Reference sites and historical databases will be used as a
basis for this evaluation.
10.2 METHODS
10.2.1 Protocols for Fish Sampling
10.2.1.1 Site Selection. Sampling sites have been selected according to the base EMAP grid design.
The candidate reference streams were identified by Region 3 and state biologists. Geographic
Positioning System technology and USGS 7.5 minute base maps wfll be used to locate the latitude and
longitude of each EMAP sampling reach (x-site). The x-site will be used as the center of the sampling
reach.
10.2.1.2 Fish wfll be collected according to time and distance criteria. Collection time should continue
for no less than 45 minutes and no greater than 3 hours for a distance of between 150 - 500 meters in
order to obtain a representative sample over a distance equivalent to 40 channel widths. Homogeneous
(or large systems) without dearly-defined habitat types should be sampled wherever best fish habitat is
found. Sampling gear wDI consist of backpack electrofishing equipment supplemented by block netting
and seining in habitats where flow, substrate and structure affect capture of benthlc species. 'These
protocols are designed for use in wadeable streams.
10.2.2 At the sampling site
a. Once x-site has been located, determine fish sampling reach as a function of mean channel
width taken at the x-slte (40 channel widths). The x-site wfll serve as the midpoint of the sample
reach (except in cases indicated in Section 6. Physical Habitat Assessment).
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EMAP-SW-Streams, Rsh Indicator, Reid Methods. Section 10, Revision No. 2, March 1994, Page 2 of 14
b. Walk the length of the sample reach to determine pool depths, habitat composition, barriers
and obstructions which may impede or aide in fish capture. Determine if reach requires block
nets be placed at upstream and downstream ends of stream. (This wfll usually be required only
in those places where sample reach is a large continuous pool.)
c." If the downstream or upstream limits of the sampling reach (pro-determined linear distance
from x-site) fall in the middle of a habitat unit, extend the planned fishing reach to the bottom of
the downstream habitat or the top of the upstream habitat If no habitat break is evident within
20 m of either endpoint (e.g., a long pool or glide), block net at the endpbint Record total
length of stream reach sampled on data sheets, noting such exceptions to the protocol on the
site description and fish field data forms.
d Decision to use electrofishing equipment wfll depend on size of site, flow, conductivity and
turbidity. If conductivity is below 10 t*S or If flow is too high, site too deep or water Is too turbid
to assure safe footing or locate stunned fish, crew may consider use of seine only or determine
that site Is TJnsampleable". THIS IS A SAFETY DECISION.
e. In case of emergency, determine location of means of easy egress from stream.
10.2.3 Electroflshlnq
10.2.3.1 A generator-powered backpack electrofishing unit with a 45° titt cutout switch wfll be used.
Always wear waders and heavy-duty insulated rubber gloves when working with electricity In
water. Wear polarized sunglasses to aid vision. All four crew members wffl participate in electrofishing.
a. Set unit to 300VA and pulsed DC. Based on stream conductivity, select Initial voltage setting
(150-400V for high conductivity >300»S; 500-800V for medium conductivity 100uS; 900-1100V
for low conductivity <100nuS waters). Determine that all crewmembers are wearing waders and
gloves and are dear of the anode. Start generator, set timer, and depress switch to begin
fishing. Starting at the bottom of the most downstream riffle, pod, or other habitat type from the
x-site and fish in an upstream direction, parallel to the current
Adjust voltage and waveform output according to sampling effectiveness and Incidental
mortality to specimens. Backpack unit is equipped with an audio alarm when output
voltage greater than 30V Is present It also serves as an Input current indicator for pulse
cycles greater than SHz. It begins as a strong continuous tone and begins to beep
slowly at currents of 1.25 amps. It beeps faster as Input current increases. In case of
overload (in excess of 3 amps), the beep becomes very rapid and the LED overload
Indicator comes on Release the anode switch and adjust voltage and waveform and
continue fishing.
b. With switch depressed, slowly sweep electrodes from side to side in the water In riffles and
pods. Sample available cut-bank and snag habitat as well as riffles and pods. Move wand in
and out of large snags or deep cuts or release the electrode switch, move the wand away
slightly, depress the switch again and sweep the wand away from the cover to draw fish out Into
open.
In streams wider than can be spanned by sweeping the electrode from side to side, it may be
necessary to work from the mkJIIne of the stream reach to the banks. At the bottom of the
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EMAP-SW-Streams, Fish Indicator, Reid Methods, Section 10, Revision No. 2, March 1994, Page 3 of 14
reach, start by samplintg from the midline to the LEFT BANK (FACING DOWNSTREAM). At
each subsequent transect, switch to the opposite side from the one which has just been fished.
In stretches with deep pools, fish the margins of the pool as much as possible, being extremely
careful not to step into deep water.
c. Netters follow along beside or slightly behind person operating shocker on the anode side
and net stunned fish which are then deposited In separate buckets or holding tanks. (Water
should be changed periodically.) Minnow seines (4m x 2m x 0.5cm) and kick nets (2m x 2m x
0.5cm) may be used to block in riffles, pools and snags.
d. Continue unto 40 channel widths or a segment of at least 150 m has been sampled.
Sampling should continue for at least 45 mln. Distance sampled should not exceed 500 m;
sampling time should not exceed 3 hours. Record total time spent collecting and shocking
time on data sheets.
e. If fish show signs of stress (loss of righting response, gaping, gulping air, excessive mucus),
stop at intervals and work up catch. Be sure to release fish downstream to reduce likelihood of
resampling. This should only be necessary on very warm days, in long reaches or H very large
numbers of fish are collected.
10.2.4. Seining
10.2.4.1. Seining will be used in conjunction with electrofishing to ensure sampling of those species
which may otherwise be underrepresented by an electrofishing survey alone (eg. darters, sculpins).
Seines wfll be used as block or kick nets to selectively Isolate sections of the stream being electrofished
(eg. snags, riffles, cutbanks). This method has proven effective in collecting sculpins, darters, dace and
other benthic cyprinids. Seining may also be used in sites where stream is too deep for electrofishing to
be conducted safely. In those sites, seining in riffles and pool margins wOl provide some indication of
the assemblage of common species.
a Riffle klcknet sampling consists of placing the net perpendicular to the current with the
leadline on the bottom. The net (2m L x 1.25m W x .6cm mesh) is tilted slightly to form a
pocket for trapping fish. Starting about 2m upstream, crewmembers disturb the substrate by
shuffling their feet and overturning rocks. Seiners raise the net and tt is carefully examined for
fish.
b. Pool seining consists of pulling the net (3m x 2m x .6cm mesh) back and forth across the
pool, using the shore and other natural habitat breaks as barriers. Another technique is to pull
the net along in a downstream direction and sweeping toward the shore. Net is examined as
above. •
c. Block nets may be used in very large pools to limit fish escape or as seines. Large nets are
typically deployed parallel to the current and swept to shore.
10.2.5. Sample Processing
10.2.5.1 Work as quickly as possible to minimize stress to live fish. Alt members of crew could work to
separate fish into families or obvious 'morphotypes* (two dorsal fins vs. one, sucker mouth, catfish.
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EMAP-SW-Streams, Rsh Indicator, Reid Methods, Section 10, Revision No. 2, March 1994, Page 4 of 14
trout). Once the rough sort has been completed, one person should identify, measure and examine the
fish while another writes.
a. Sort fish by species into small buckets and containers. Taxonomic identification should be
performed only by trained ichthyologists familiar with the fish species of the region.
b. To minimize handling, threatened and endangered species should be identified,
counted and returned Immediately to the stream. Consult the APS Publication 'Protected
Rshes of the United States and Canada*, J.E. Johnson, 1978. If conditions permit and stress
to individuals wil be minimal, photograph fish for voucher purposes. Indicate if photographed
on data sheet If protected fish have died, they should be vouchered in formalin. At the earliest
possible time, the appropriate state officials should be notified.
Try to process one species completely before going on to the next However, where there are
many individuals of easily Identified species, processing may be facilitated by keeping a tally
count of the number of Individuals on the second line of the identification column for each
species and totalling the tally once processing is complete. Each fish must be examined
Individually but small spedes may be handled in small manageable groups to speed processing.
c. Sport fish and very large specimens (suckers, etc.) should be identified, measured to the
nearest mm (standard length, total length, body depth, Fig. 10.1), examined for external
anomalies (Table 10.3.), and released. Record all information on field data sheets. Separate a
subsample of the target species for the fish tissue contaminants analysis. Voucher specimens
(up to 25) of smaller individuals of each species should be kept (see Section 10.2.7 for voucher
protocols). If only large Individuals are collected, photograph each species and indicate if
photographed on data sheet
d. Identify other species. Measure the largest and smallest Individuals (standard length and
total length) to provide a size range for the species. Use the space beneath line for the
common name on the data sheet to tally the number of individuals. Examine individuals in the
sample for external anomalies and note proportion and type of anomalies observed keeping a
running tally count In the column for comments and anomalies. Once all Individuals have been
processed, record total number under count and the species code beneath the common name.
Separate a subsample of the target species for the fish tissue contaminants analysis. Voucher a
subsample of 25 Individuals from each species identified (see Section 10.2.7 for voucher
protocols). If fewer than 25 Individuals are collected of a species, keep all of the specimens.
Count the number of unidentifiable specimens (If a large number, estimate number of individuals
and note as comment) and preserve them in the jar labeled UNKNOWN. Record all information
on field data sheets.
e. Preserve voucher and unidentifiable ("unknown") specimens in 10% formalin solution (37%
formaldehyderwater). Start with a concentrated solution of formaldehyde and dilute to tinal
volume.
f. Affix labels containing stream ID number, date, team ID and sample type to appropriate jars
from that site. Enter barcode ID in the space provided on the field data sheet
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EMAP-SW-Streams, Fish Indicator, Field Methods, Section 10, Revision No. 2, March 1994, Page 5 of 14
Scala Sinol* Anu
Utml Lira
Standard Ungth
Fort Un«th
•
Toul
Seal*
Figure 10.1. Fish measurement With mouth dosed and fish on right side, slide fish to touch 'bump
board*. Read scale to nearest mm. (modified from Lagler, 1956).
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EMAP-SW-Streams, Fish Indicator, Reid Methods, Section 10, Revision No. 2, March 1994, Page 6 of 14
10.2.6 External Anomalies
10.2.6.1 External anomalies may result from sublethal environmental or behavioral stress, diseases, and
toxic chemicals. Readily identified external anomalies Include deformities, eroded fins, lesions, tumors,
diseases and parasites.
Categories and Codes:
NONE (Code - N). No anomalies present
Deformities (D) are skeletal anomalies of the head, spine, and body shape.
Eroded fins (E) appear as reductions of fin surface area or hemorrhage of fin rays.
Lesions (L) are open sores or exposed tissue.
Tumors. (T) are areas of irregular cell growth which are firm and cannot be easily broken open
when pinched. (Masses caused by parasites usually can.)
Fungus, (F) may appear as filamentous or fuzzy* growth on the fins or body.
Black soot disease (B) is caused by a parasite and appears as small black cysts on the fins and
body.
'Ich' (I) is caused by a protozoan infestation and appears as white spots on the fins and body.
Anchor worm (A) is a parasitic infection characterized by a worm embedded in the flesh of the
fish.
Leeches (W) are annelid worms which have anterior and posterior suckers. They may attach
anywhere on the body.
Exopthalmia (X) or 'popeye disease* is an anomaly seen as the bulging of the eye.
Missing (M) eyes, blindness or deterioration of the lens should also be noted.
Other (0) anomalies or parasites not specified.
Caution: Spinal cord injury, lesions, and exopthalmia or missing eyes may occasionally result
from the sampling process, especially when electro-fishing. Care should be taken in the early
stages of electrofishing to use the most effective combination of voltage and pulse width while
minimizing injury to fish.
10.2.7 Voucher Protocol
10.2.7.1 With the exception of very large individuals of easBy identified species, voucher collections of ail
species will be made to provide a permanent, archived, historical record of fish collections. Voucher
material falls into three categories.
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EMAP-SW-Streams. Fish Indicator, Reid Methods, Section 10, Revision No. 2, March 1994, Page 7 of 14
pateaorv 1. Large easily identified species OR adults may be difficult to identify .QH the species Is
uncommon in region. Preserve 1-2 small (<150 mm TL) adult Individuals per site plus 2-5 YOY and
juveniles. If only large adults are collected, reserve smallest individual untfl voucher procedure is
complete and preserve ONLY if space is available. In specimens > 100mm make a small slit on the
lower abdomen on the RIGHT side. Document with photograph.
American Eel White Sucker Buffalo fishes Drum
Sturgeon Longnose Sucker Bullhead catfish Carp
Paddlefish Hogsucker Channel catfish Salmonlds
Gars Quillback Esocids Grapples
Bowfm Carpsuckers Me/one spp. Mlcropterus spp.
Mooneye and Goldeye Moxosfoma spp. Shads Walleye and Sauger
Category 2. Small to moderate-sized fish OR difficult to identify species. Preserve up to 25 adults,
Juveniles and YOY per site.
Lampreys Troutperch Sculplns Madtoms
Cyprinids Chubsucker Sunfish Sticklebacks
Darters Topminnows SDversWes Mudminnows
Category 3. Species of 'special concern*. These are state and federal species listed as protected.
Photograph and release. If specimens have died, voucher them, note on datasheet and notify
appropriate state official as soon as possible. Listed species are protected unless otherwise noted. SC
« special concern. It is unlikely crews wDl encounter protected species in Maryland.
NEW YORK VIRGINIA (AH SC) ' Satinfln shiner
Mooneye Spotfinchub New River shiner
Round whltefish YeHowfln madtom Kanawha minnow
Silver chub Orangefln madtom Cheat minnow
Black redhorse (SC) Roanoke logperch Longnose sucker
Longear sunflsh Mountain madtom
Eastern sand darter WEST VIRGINIA (Ail SC) Crystal darter
Bluebreast darter Mountain brook lamprey Shield darter
Gilt darter Shoveinose sturgeon Slimy sculpln
Longhead darter Paddlefish Potomac sculpln
Mooneye
PENNSYLVANIA Grass pickerel MARYLAND
Eastern sand darter Bedside dace Maryland darter
Popeye shiner Striped bass
a. Once alt Individuals of a species have been processed, place voucher subsample in kDl jar
containing a strong (approx 20%) formalin solution. (Use the "Unidentified* jar for this.)
Individuals > 150mm SL should be silt on the lower abdomen of the RIGHT side.
b. When specimens are dead, transfer to a perforated zipiock bag containing waterproof label
with site ID, barcode, species name and code. Place In 'Voucher* jar In 10% formalin. BE
SURE THAT JAR IS LABELED WITH VOUCHER LABEL with site ID, barcode and date.
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EMAP-SW-Streams, Fish Indicator. Reid Methods, Revision No. 3, April 1995, Page 8 of 13
c. Continue until all species are processed. Seal voucher jar with electricians' or clear tape.
Check that jar Is correctly labeled. Enter BARCODE ID In appropriate place on field data sheet.
d. Place all Individuals of unidentified fish Into UNKNOWN jar with label with site ID, barcode and
date. Seal jar with tape. Enter BARCODE ID in appropriate place on field data sheet
e. Transport to storage depot at end of week. Store In a cool, ventilated space.
10.3 SAFETY
10.3.1 Primary responsibility for safety while electrofishir.g rests with the fish team leader. All crew
members should receive training in First Aid and CPR. Electro-fishing units have a high voltage output and
may deliver dangerous electrical shock. WhDe electrofishing, avoid contact with water unless sufficiently
insulated against electrical shock. Use chest waders with non-slip soles and water-tight rubber (or
electrician's) gloves that cover to the elbows. If they become wet inside, stop fishing until thoroughly
dry. Avoid contact with anode at all times. Cathode is also a potential shock hazard. At no time
while electrofishing should a crewmember reach into the water for any reason. If ft Is necessary for a
crewmember to reach into the water to pick up a fish or something which has been dropped. DO SO ONLY
WHEN THE ELECTRICAL CURRENT HAS BEEN INTERRUPTED AND THE ANODE REMOVED FROM
THE WATER. Do not resume electrofishing until crew is sure that all is clear. The electrofishing
equipment provided is equipped with a 4? tilt switch which interrupts the current Do not make any
modifications to the electrofishing unit which would make ft impossible to turn off the electricity.
10.3.2 General safety guidelines should be observed. If waders or gloves develop leaks, leave the water
immediately. Avoid operating electrofishing equipment near people, pets or livestock. Discontinue any
activity in streams during thunderstorms or heavy rain. Rest if crew becomes fatigued. ,
10.3.3 Formaldehyde is an extremely caustic agent and may cause severe Irritation on contact of vapors
or solution with skin, eyes or mucus membranes. It Is a potential carcinogen. Contact with vapors or
solution should be avoided. Wear gloves and safety glasses and always work in a well-ventilated area. In
case of contact with skin or eyes, rinse Immediately with large quantities of water. Store stock solution in
sealed containers in safety cabinet or cooler lined with vermicullte.
10.3.4 Ethanol has been shown to have deleterious effects on the central nervous system and may cause
skin irritation. Care should be taken to avoid contact with vapors. Absolute ethanol is flammable and
should be kept away from open flame or sparks. Store In sealed containers in safety cabinet or cooler
lined with vermiculite.
10.3.5 Gasoline Is extremely volatile and flammable. Its vapors readily ignite on contact with heat, spark or
flame. Never attempt to refill the generator while h Is running. Always allow the generator to cool before
refilling. Keep gasoline out of direct sunlight to reduce volatilization and vapor release. Always wear
gloves and safety glasses when handling gasoline. Keep gasoline only in approved containers and store in
tightly closed container in safety cabinet or cooler lined with vermicullte.
10.4 QUALITY ASSURANCE/QUALITY CONTROL
10.4.1 The U.S. Environmental Protection Agency requires that all monitoring and measurement projects
have written and approved quality assurance project plans. This QA/QC plan is presented in the
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EMAP-SW-Streams, Fish Indicator, Field Methods. Section 10, Revision No. 2, March 1994, Page 9 of 14
Environmental Monitoring and Assessment Program Integrated Quality Assurance Project Plan for the
Surface Waters Resource Group (EPA600/X-91/080, Rev. 2.00).
10.4.1.1. Fish Sampling Quality Assurance Plan.
1. Sampling Design. Fish will be collected according to time and distance criteria. Results from the
1993 MAHA pilot project indicate a significant relationship between time sampled and distance sampled
and number of individuals collected. A significant relationship also exists between time and distance
sampled and number of species collected (this is subject to validation by museum). The length will be
determined by a multiple of 40 channel widths indexed at the x-site. Stream segment length should be
at least 150 m, but not more than 500 m. Collection time should be at least 45 min and not more than 3
hr per site. The length of channel surveyed will be used to define the stream segment for other
sampling activities. The EMAP research indicator team will continue its evaluation of reach length criteria
for larger wadeable streams.
2. Sampling and Analytical Methodologies. Fish collection protocols for electrofishing and seining are
given in Subsection 1.2.3 -1.2.4.
3. Quality Assurance Objectives
Precision-Precision of fish samples will be accomplished by careful application of methods and
thorough sampling of all available fish habitats.
Taxonomic Accuracv-Taxonomic accuracy will be achieved by careful adherence to protocols,
thorough sample preparation and analysis, and re-analysis of a subset of samples at a later date.
Organisms will be identified to the lowest possible taxonomic level, using standard references
and keys, and enumerated. Taxonomic identification wfll be performed only by trained
ichthyologists familiar with the fish species of the region. Verification of taxonomic identification
will be performed on voucher material by professional ichthyologists at the National Museum of
Natural History, Smithsonian Institution, Washington, D.C. or other regional museums and
academic institutions. Taxonomic nomenclature wfll follow Common and Scientific Names of
Fishes from the United States and Canada, American Fisheries Society Special Publication 20.
Completeness-Valid fish data are required from 90% of the sites visited from which fish were
collected.
Representativeness-Fish samples will be collected from all available habitats along the entire
length of stream segment.
4. Quality Control Procedures • •
Field Operations-All personnel involved in fish sampling are trained in the selection of sample
locations and in the use and maintenance of both seines and electrofishing gear. The minimum
qualifications for fish team leader are a BS in fisheries with 3-5 yr experience in fishes in region.
Preferred qualifications are MS in fisheries or zoology with specialization in fishes of region.
JLaboratorv Operations-Quality control of laboratory processing of fish samples wfll be achieved
by proper selection of taxonomic techniques and references, and preparation, documentation,
and archiving of reference specimens.
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EMAP-SW-Streams. Fish Indicator, Reid Methods, Section 10. Revision No. 2, March 1994, Page 10 of 14
information Manaaement-Fish samples returned to the laboratory are logged In and their
processing tracked until the samples are completed. All data sheets are Inspected for
completeness, accuracy, and legibility before proceeding to the next sample. Raw data sheets
are retained in a file, and the data is entered Into an ASCII fOe.
10.5 REFERENCES
Lagler, K.R. 1956. Freshwater fishery biology. 2nd. Edition. WDllarn C. Brown Co., Dubuque,
IA.
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EMAP-SW-Streams, Fish Indicator, Field Methods, Section 10, Revision No. 2, March 1994, Page 11 of 14
TABLE 10.1. FIELD PROTOCOLS FOR FISH COLLECTION.
1. Site Selection
a. Determine channel width.
b Survey sample reach.
c. Determine if reach requires block nets.
d. If conductivity is below 10 uS or ff flow, depth or turbidity make ft unsafe to electrofish, crew
may elect to use seine only or not sample. THIS IS A SAFETY DECISION.
e. In case of emergency, determine location of means of easy egress from stream.
2. Electrofishing
a. Set unit to 300 VA and pulsed DC. Select initial voltage setting. Start generator, set timer,
and depress switch to begin fishing.
b. Fish In an upstream direction, parallel to the current Adjust voltage and waveform output
according to sampling effectiveness and mortality fish specimens.
c. With switch depressed, sweep electrodes from side to side in the water. Sample available
cut-bank and snag habitat as well as riffles and pools.
d. Netters follow operator and net fish. Deposit fish in buckets. Block with seines in riffles,
pools and snags.
e. Continue for 40 channel widths. Record total time spent collecting and shocking time on
data sheets.
f. Identify and release any threatened and endangered species.
g. identify and measure (SL, TL, BD) sport fish and very large specimens, record external
anomalies, and release ur.;
h. Identify other specimens. Determine number of individuals in species, measure largest and
smallest Individuals, and voucher as described in Voucher Protocol.
i. Retain a subsample of target species for Fish Tissue Contaminants analysis.
3. Seining will be used in conjunction with electrofishing and in sites where stream is too deep for
electrofishing to be conducted safely.
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EMAP-SW-Streams, Rsh Indicator, Reid Methods, Section 10, Revision No. 2. March 1994, Page 12 of 14
TABLE 10.2. PROCEDURE FOR VOUCHERING RSH SPECIES AND UNIDENTIFIED SPECIMENS.
Category 1. Large species, difficult to identify, OR uncommon In region. Preserve 1-2 adults plus 2-5
YOY and juveniles. Document with photograph.
American Eel White Sucker Shads
Sturgeon Longnose Sucker Bullhead catfish
Paddlefish Hogsucker Channel catfish
Gars Qufllback Esocids
Bowfin Carpsuckers Morono spp.
Mooneye and Goldeye Maxostoma spp. Drum
Carp Buffalo fishes Salmonlds
Walleye and Sauger Micropterus spp. Grapples
Category 2. Small fish OR difficult to identify species. Preserve up to 25 adults, juveniles and YOY per
site.
Lampreys Troutperch Sculpins Madtoms
Cyprinids Chubsucker Sunflsh Sticklebacks
Dartors Topminnows . SOversldes Mudminnows
a. Place subsample of 25 individuals of total catch into a kfll jar containing 10% buffered
formalin solution.
b. When specimens are dead, transfer to perforated ziplock bag containing a waterproof label
with site ID, barcode, date, crew ID and species name.
c. Repeat process for all species identified.
d. If fewer than 25 individuals are collected, preserve entire sample.
e. Place all ziplock bags in labelled voucher jar.
Category 3. State and federal species listed as protected. Photograph and release. Voucher if
specimens have died.
NEW YORK VIRGINIA (An SC) Paddlefish
Mooneye
Mooneye Spotfin chub Grass pickerel
Round whitefish Yellowfln madtom Redside dace
Silver chub Orangefln madtom Satinfin shiner
Black redhorse (SC) Roanoke logperch Popeye shiner
Longear sunfish Mountain madtom New River shiner
Eastern sand darter Kanawha minnow
Bluebreast darter WEST VIRGINIA (All SC) Cheat minnow
Longhead darter Longnose sucker
Gilt darter Mountain brook lamprey Crystal darter
Gilt darter Shovelnose sturgeon Shield darter
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EMAP-SW-Streams, Fish Indicator, Field Methods. Section 10, Revision No. 2, March 1994. Page 13 of 14
TABLE 10.2. PROCEDURE FOR VOUCHERING FISH SPECIES AND UNIDENTIFIED SPECIMENS
(CONTINUED)
Category 3. State and federal species listed as protected. Photograph and release. Voucher If
specimens have died. (Continued)
WEST VIRGINIA (ALL SC) PENNSYLVANIA MARYLAND
(CONTINUED)
Slimy sculpin Eastern sand darter Maryland darter
Potomac sculpin Striped bass
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EMAP-SW-Streams, Fish Indicator, Reid Methods, Section 10, Revision No. 2, March 1994, Page 14 of 14
TABLE 10.3. EXTERNAL ANOMALIES
Categories and Codes:
NONE (Code » N). No anomalies present
Deformities (D) are skeletal anomalies of the head, spine, and'body shape.
Eroded fins (E) appear as reductions of fin surface area or hemorrhage of fin rays.
Lesions (I) are open sores or exposed tissue.
Tumors CO are areas of irregular cell growth which are firm and cannot be easily broken open
when pinched. (Masses caused by parasites usually can.)
Fungus (F) may appear as filamentous or fuzzy" growth on the fins or body.
Black soot disease (B) is caused by a parasite and appears as small black cysts on the fins and
body.
•ich' (I) is caused by a protozoan infestation and appears as white spots on the fins and body.
Anchor worm (A) is a parasitic infection characterized by a worm embedded in the flesh of the
fish.
Leeches (W) are annelid worms which have anterior and posterior suckers. They may attach
anywhere on the body.
Exopthalmia (X) or "popeye disease' is an anomaly seen as the bulging of the eye.
Missing (M) eyes, blindness or deterioration of the lens should also be noted.
Other (0) anomalies or parasites not specified.
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EMAP-SW-Streams, Fish Tissue Contaminants Indicator-Field, Section 11, Revision No. 2, Page 1 of 6
SECTION 11
FISH TISSUE CONTAMINANTS INDICATOR
11.1 INTRODUCTION
11.1.1 When coupled with a study design such as EMAP, the fish tissue contaminants indicator which
measures bioaccumulation of toxic chemicals, can be used to estimate regional risks of consumption to
predators of fish, either wildlife or human and to track how this risk changes with time In a regjon. It is
also meant to be used in conjunction with the other diagnostic indicators (physical habitat, water
chemistry, land L'sc. population density, other records of relevant anthropogenic stresses) and response
indicators (fish, macroinvertebrates, periphyton) to diagnose whether the probable cause of stream
degradation, when K is shown by the response indicators to occur, is water quality or physical habitat
11.1.2 The various studies that have been done on fish tissue contaminants have focused on different
pans of the fish: whole fish, fillets, livers. EMAP-SW will focus on whole fish because of its emphasis on
the ecological health of the whole stream (as opposed to a focus on human health concerns). Whole
fish are a good ecological Indicator and a better indicator of risk to piscivorous wildlife than fillets. It is
hoped to also be able to say something about risks to human health by analyzing whole fish. Whole fish
also present fewer logistical problems for field crews (no gutting required in the field) and the analytical
lab (no filleting necessary).
11.1.3 For the fish contaminants indicator in the 1994 pilot of EMAP-SW STREAMS, an attempt will be
made to collect two fish samples at as many sites as possible. One sample, of Primary Target
Species, will be of stream fish whose adults are small (dace, chub, scuipins, stonerollers, shiners, and
darters). The second sample, where available, of a Secondary Target Species, will be of a species
whose adults are of larger size (suckers, bass, trout, sunfish, carp). In addition to being more
ubiquitous than the larger fish (and therefore more likely to be present In sufficient numbers to
composite), small fish have other advantages over large fish. Most importantly, It may be possible to get
a more representative sample of the contaminant load in that stream section (although it would be at a
lower expected level of bioaccumulation) by compositing say, in the range of 20 to 200 small fish
individuals than by compositing 3 to 5 large fish. Small fish may be a more appropriate Indicator for
assessing ecological risk, as they might be expected to be prey for a larger number of fish-eating
animals (the majority of which wBI be piscivorous birds and small mammals). Small fish should afford
savings in time and money, being easier to composite in the analytical lab, and offering potential savings
in field processing. The major advantage that larger fish could potentially offer, whether predators
(piscivores) or bottom feeders, is a higher level of bioaccumulation and thus greater sensitivity to detect
contaminants. The relative bioaccumulation of contaminants by large and small stream fish is not
known, thus the reason for having Primary and Secondary Target Species in this study.
11.1.4 Therefore one of the goals of the pDot studies, is to examine if small fish in the same stream
section are bioaccumulating significantly relative to the larger fish. If they are, their other advantages
would warrant targeting them for collection in further studies. Some group of these species with small
adult individuals, perhaps representing a functional feeding group would be logical candidates for target
species in a national streams survey. It is also possible to get relative bioaccumulation data from this
kind of study, which may be of use In extrapolating from levels found in small fish to what might be
found in a species with large adults (which are more appropriate for human risk assessment) if
relationships between some prevalent species could be established.
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EMAP-SW-Streams, Rsh Tissue Contaminants Indicator-Reid, Section 11, Revision No. 2. Page 2 of 6
11.1.5 In trying to answer these questions, the field crews' efforts to apply the protocol for sampling,
handling and shipping in a consistent manner are critical. The diligence of the field crews in following
the protocols is especially important In a status and trends study such as EMAP-SW where it is critical to
get a standard sample from each site so that there is confidence that differences seen over time and
between sites represents variations in the ecosystems and not differences in sampling and handling
between the crews. Suggestions from field crew members on how the protocol can be improved are
welcorrod and will be incorporated to improve them, but protocols should be followed as written until
official changes are made. If any questions arise for which the answers are not contained In the field
guide, crews are encouraged to call the Indicator lead, Roger Yeardley, at (513) 569-7093. If It Is
necessary to leave a message on the voicemafl at this number, be sure and leave a phone number or
fax number through which a reply can be made. If necessary, a reply to a crew's question can be
incorporated Into the voicemafl greeting and the crew can call back to the above number that night or
the next day and get the answer, if message Is received and satisfactory, leave a second message
acknowledging this so that the voicemail greeting can be changed.
11.2 SELECTING RSH TISSUE SPECIMENS
11.2.1 If possible, obtain one sample each, of the desired weight or number (see below) of similarly
sized* individuals, from the Primary and Secondary target species lists (2 composite samples total).
To judge if the proper amount of a target species is present In the fish catch, weight wOl be used for
primary target species and number of individuals of sufficient size wfll be used for secondary target
species.
I. PRIMARY TARGET SPECIES
Small adult fish
(in priority order)
WEIGHT
1) Blacknose Dace 50** • 400 g
2) Another Dace species 50** • 400 g
3) Creek Chub or Fallfisn 50** - 400 g
4) Slimy Sculpin/Mottied ScuJpin 50** • 400 g
5) Stoneroller 50**-400 g
6) A Darter species 50** - 400 g
7) A Shiner species 50** - 400 g
A) Choose the highest priority target species from the above list, that has at least enough individuals
to attain the minimum weight (50 g). Get as much weight of fish as possible within the desired weight
range (50-400 g). Use scale provided to determine weight With dean hands, place the fish on a fresh
piece of aluminum fol (dull side towards fish) before placing In weighing container.
B) If less than the desired weight of anv primary target species Is collected, send Individuals of a
small nontarget species if 50 g or more are available.
* - The general rule-of-thumb for similar size is that the smallest Individual In the sample should be at
least 75% of the length of the largest Individual. This rule applies to both primary and secondary target
species. Crews just need to keep this criterion in mind whBe selecting the final sample. Any obviously
small or large individuals should not be kept if there is a sufficient sample to return or discard them. If
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EMAP-SW-Streams, Fish Tissue Contaminants Indicator-Field, Section 11, Revision No. 2, Page 3 of 6
there is a conflict between criteria, getting a sufficient sample is a higher priority than getting similar-
sized individuals
*• - This weight represents the minimum amount of tissue needed for laboratory analysis. Crews
should not settle for the minimum amount(weight) If more fish are present, but instead send as many
fish as possible up to the 400 g weight goal.
II. SECONDARY TARGET SPECIES
Collect and save a sample of secondary target species if such a sample of desired number of individuals
of desired size is available. Collect similar sized individuals if enough are present
Larger adult fish DESIRED DESIRED
(in priority order) SIZE NUMBER
1) White sucker ~ 2120 mm 5
2) Hogsucker s120 mm 5
3) A Bass species 2120 mm 5
4) A Trout species 2120 mm 5
5) A Sunfish species 2120 mm 5
6) Carp 2120 mm 5
A) If fewer than the desired number of secondary target species individuals Of desired Size are
collected, add smaller individuals of the same species, if available, to achieve the desired number (5).
B) If fewer than 5 fish of any size are available, you may send as few as 3 fish that are at least at or
near the minimum desired size (120 mm).
C) If an acceptable secondary target species sample (by the above criteria) is not available send only
the primary target species sample. If neither a primary or secondary species sample that meets
these criteria is available, use your best judgement in sending some type of fish sample.
11.3 PROCESSING TISSUE SPECIMENS
1. Keep hands, work surfaces, and wrapping materials dean and free of potential contaminants
(mud, fuel, formalin, sun screen, insect repellent, etc.)
2 Measure total weight of individuals for primary target species (to the nearest 5g) and count the
total number of individuals. Make sure that the scale is tared and the teflon weighing beaker (to
be used ONLY for weighing fish) has been rinsed with deionized or stream water. Measure the
total length fJL) of each secondary target species individual. Record all of this information
on the Fish Tissue Sample Tracking Form.
3. Write the bar code number(s)*** on the Fish Tissue Sample Tracking Form. Record if the
target species chosen was found throughout the measured EMAP stream reach (check-yes or
no), with an explanation or some further details if the answer is no. Make sure that the form is
filled out completely.
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EMAP-SW-Streams, Rsh Tissue Contaminants Indicator-Reid, Section 11, Revision No. 2, Page 4 of 6
4. Wrap fish in aluminum foQ. The primary target fish sample may be wrapped as a group.
Secondary target fish should be wrapped individually. The dull side of the aluminum foil should
be in contact with the fish. Once wrapped, place each sample in a ziplock or garbage bag.
5. Expel excess air and seal the bag(s). Wrap dear tape around the bag(s) to seal and make a
surface for each sample label.
6. Complete bar-coded fish tissue label(s). Make sure the numoer(s) is/are the same as the one(s)
on the Tracking Form, Apply It/them to the tape surface(s). Cover the label(s) with dear.
waterproof tape. As labels may sometimes fall off, there should always be a label on the inner
bag.
7. Place labeled ziplock or garbage bag(s) into second plastic bag(s) and seal and label second
bag(s) (I.e.. repeat steps 5 & 6).
8. Place double-bagged sample(s) in portable freezer untfl shipment Keep samples frozen untl
shipment
••• - If both primary and secondary target species are collected, the two samples should be wrapped
and bagged separately, with separate bar codes and labels, but only one Sample Tracking Form.
11.4 SHIPPING TISSUE SPECIMENS
11.4.1 Crews collecting fish tissue samples wfll have a portable freezer for freezing and storage of
samples until shipment Samples should be shipped weekley. Crews may want to have them picked up
at the motel in the morning, or ship in the evening and Indude that day's samples. Samples held by
crews over the weekend must be kept frozen.
11.4.2 Ship samples on toe or dry ice. When using ice, double bag the Ice and tape the last bag shut
to prevent contamination of samples by melting ice. Place toe or dry tea In a cooler with a week's worth
of samples (fish tissue, chlorophyll.periphyton.sediment). Place the week's field forms as well as the
computer-generated shipping forms for these indicators in a ziplock bag and Indude with the shipment
(tape to the underside of cooler lid). Tape coolers shut Ship by Federal Express next day service. If
shipping on dry Ice, attach a fled-out DRY ICE warning label to the shipping container and enter the
necessary information pertaining to dry ice in sections 3 & 4 on the FedEx Alrbffl, when filling out the
FedEx Airbill.
Address shipment to: .
U.S. EPA
Attn: EMAP
3411 Church St
Cincinnati, OH 45244
(513) 569-7095
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EMAP-SW-Streams, Fish Tissue Contaminants Indicator-Reid, Section 11, Revision No. 2, Page 5 of 6
11.5 TARGET ANALYTES
11.5.1 For details of how the fish are processed by the analytical laboratory to determine tissue
concentrations of EMAP-SW's target analytes, refer to the '1994 Pilot Stream Laboratory Methods
Manual For Streams."
ANALYTE (CAS NUMBER)
Aldrin (309-00-2)
Aluminum (7429-90-5)
Arsenic (7440-38-2)
Cadmium (7440-43-9)
Chlordane-cis (5103-71-9)
Chlordane-trans (5103-74-2)
Chromium (7440-47-3)
Copper (7440-50-8)
2,4'-DDD (53-19-0)
4,4'-DDD (72-54-8)
2.4'-DDE (3424-82-6)
4,4'-DDE (72-55-9)
2.4'-DDT (789-02-6)
4,4'-DDT (50-29-3)
Dieldrin (60-57-1)
Endosulfan-l (959-98-8)
Endosulfan-ll (33213-65-9)
Endrin (72-20-8)
Heptachlor (76-44-8)
Heptachlor Epoxide (1024-57-3)
Hexachlorobenzene (118-74-1)
Hexachlorocydohexane [Gamma-BHC/Lindane) (58-89-9)
Iron (7439-89-6)
Lead (7439-92-1)
Mercury (7439-97-6)
Mirex (2385-85-5)
Nickel (7440-02-0)
trans-Nonachlor (3765-80-5)
cis-Nonachlor (5103-73-1)
Qxychlordane (27304-13-8)
PCB Congeners
2.4-Dichlorobiphenyl, #8 (34883-43-7)
2,2',5-Trichlorobiphenyl, #18 (37680-65-2)
2.4.4'-Tricnlorobiphenyl, #28 (7012-37-5)
2,2',5,5'-Tetrachloroblphenyl, #52 (35693-99-3)
2,2',3,5'-Tetrachlorobiphenyl, #44 (41464-39-5)
2,3',4,4'-TetracWorobiphenyl, #66 (32598-10-0)
2,2',4,5,5>-PentacWorobiphenyl, #101 (37680-73-2)
2,3'.4,41,5-Pentachlorobiphenyl, #118 (31508-00-6)
W.M'.S.S'-Hexachlorobiphenyt. #153 (35065-27-1)
2,3,3',4,4'-PentacNorobipheny1, #1U5 (32598-14-4)
-------
EMAP-SW-Streams, Rsh Tissue Contaminants Indicator-Reid, Section 11, Revision No. 2, Page 6 of 6
PCB Congeners (Continued)
2,2',3.4,41,5-HexachJorobipheny1, #138 (35065-28-2)
2.2>.3.4>,5.5'.6-Heptachlorobiphenyl, #187 (52663-68-0)
2.2',3.3',4,4'-Hexachlorobipheny1, #128 (38380-07-3)
2.2>,3,4,41,5.51-Heptachlorobipheny1, #180 (35065-23-3)
2.2'.3.3',4,4'.5-Heptachlorobipheny1t #170 (35065-30-6)
2.21.3,3'.4t4',5,6-Octachlorobiphenyl, #195 (52663-78-2)
2.2>,3,3',4,4>.5.5',6-Nonachloroblphenyi, #206 (40186-72-9)
Decachlorobiphenyt, #209 (2051-24-3)
3,3'.4.4' Tetrachlorobiphenyl, #77 (32598-13-3)
3,3',4,4',5 Pentachlorobiphenyl, #126 (?)
S.S'.M'.S.S' HwcMoroblphenyt, #169 (32774-16-6)
Selenium (7782-49-2)
Silver (7440-22-4)
Tin (7440-31-5)
Zinc (7440-66-6)
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EMAP-SW-Stream, Sediment Toxicity Samples-Field, Section 12, Revision No. 2, March 1994, Page 1 of 1
SECTION 12
SAMPLE COLLECTION AND SHIPMENT FOR SEDIMENT TOXICITY SAMPLES
12.1 SEDIMENT COLLECTION AND SHIPMENT
12.1.1 Use the sediment left over from the Sediment Metabolism Indicator, Section 8. Benthic (Sediment)
Metabolism: Field Methods.
12.1.2 Mix sediment well with a stainless steel or plastic mixing spoon.
12.1.3 Fill a 1 gallon ziplock bag with at least 3 L of sediment.
12.1.4 Close bag, making sure seal is secure.
12.1.5 Fill out I.D. label; place label and sediment inside another ziplock bag. Seal.
12.1.6 Place these bags inside the third and final ziplock bag and seal.
12.1.7 Hold sediments samples on ice (do not freeze!) for laboratory analysis.
12.1.8 Ship samples to: Mark Smith/Lori Herrin/Ann Kneipp
TAI, C/0 U.S. EPA
3411 Church St.
Cincinnati, OH 45244
Phone (513) 569-7091
FAX (513) 569-7081
12.2 SEDIMENT TESTING METHODS
12.2.1 For further information on the testing of these sediments, see the '1994 EMAP-SW Pilot Laboratory
Methods Manual For Streams,' Section 8, Sediment Toxicity Test Methods.
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EMAP-SW-Streams, Safety and Health, Field Meth., Section 13, Revision No. 2, March 1994, Page 1 of 5
SECTION 13
SAFETY AND HEALTH
13.1 INTRODUCTION
13.1.1 Individuals working in the field and/or In the laboratory must become thoroughly familiar with all
EMAP safety and health requirements and procedures. This Section is found In both the EMAP 1994, Pilot
Field Operations Manual and the Laboratory Methods Manual. Specific safety and health requirements may
be found in the various indicator sections of the above mentioned documents.
13.1.2 Personnel safety and health are of the highest priority for all Investigative activities and must be
emphasized in safety and health plans for field, laboratory, and materials handling operations. Preventive
safety measures and emergency actions must be emphasized. The Safety and Health Section must be
available to all personnel and should be included in the field training.
13.1.3 Collection and analysis of samples (e.g., benthic Invertebrates, fish, periphyton, sediment) can involve
significant risks to personal safety and health (drowning, electrical shock, pathogens, etc.). While safety is
often not considered an integral part of field sampling routines, personnel must be aware of unsafe working
conditions, hazards connected with the operation of sampling gear, boats, and other risks (Berry et a!.,
1983). Management should assign health and safety responsibilities and establish a program for training
in safety, accident reporting, and medical and first aid treatment The laboratory safety document and
standard operating procedures (SOPs) containing necessary and specific safety precautions should be
available to all persons involved in fish sample collecting and processing. Reid and laboratory safety
requirements for biomonitoring laboratories are found also In USEPA (1986) and Ohio EPA (1990).
13.1.4 A planned line of communication for safety and emergency situations is essential. Reid personnel
should have a daily check-In procedure for safety. An emergency communications plan should include
contacts for police, ambulance, fire departments, search and rescue personnel. All field personnel need to
be fully aware of all lines of communication, and the communication strategy should be included in all
indicator sections of the field training and operations manual.
13.2 GENERAL PRECAUTIONS
13.2.1 Good housekeeping practice should be followed both in the field and in the laboratory. These
practices should be aimed at protecting the staff from physical injury, preventing or reducing exposure to
hazardous or toxic substances, avoiding interferences with laboratory operations, and producing valid data.
13.2.2 Field personnel and sampling crews must have mandatory training in Red Cross first aid,
cardiopulmonary resuscitation (CPR), boating and water safety, field survey safety (weather conditions,
personal safety, and vehicle safety), presurvey safety requirements (equipment design, equipment
maintenance, reconnaissance of survey area), and electrofishing safety (American Red Cross, 1979; National
Institute For Occupational Safety and Health, 1981; U.S. Coast Guard, 1987; Ohio EPA, 1990). It is the
responsibility of the group safety officer or field sampling leader to ensure that the necessary safety courses
are taken by all field personnel and that all safety policies and procedures are followed. Persons using
sampling devices should become familiar with the hazards involved and establish appropriate safety
practices prior to using them.
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EMAP-SW-Streams, Safety and Health, Reid Meth., Section 13, Revision No. 2, March 1994, Page 2 of 5
13.2.3 Operation of fish sampling devices involves potential hazards that must be addressed by the
individuals using the equipment Hectrofishing equipment should be operated carefully, and all safety
procedures must be followed. Electrofishing should always be done with at least three individuals. Persons
using electrofishers should become familiar with the hazards involved and establish appropriate safety
practices prior to using them (Reynolds, 1983; Ohio EPA, 1990). Nott: Individuals involved in
electro-fishing must be trained by a person experienced in this method or by attending a certified
electroftshing training course.
13.2.4 Reid personnel should be able to swim. Chest waders made of rubberized or neoprene material and
suitable footwear must always be worn with a belt to prevent them from filling with water In case of a fall.
The use of a life jacket Is advisable at dangerous wading stations if one is not a strong swimmer because
of the possibility of sliding into deep water.
13.2.5 Many hazards lie out of sight in the bottoms of lakes, rivers and streams. Broken glass or sharp
pieces of metal embedded In the substrate can cause serious Injury if care is not exercised when walking
or working with the hands In such environments. Infectious agents and toxic substances that can be
absorbed through the skin or inhaled may also be present In the water or sediment
13.2.6 Personnel must consider and parepare for hazards associated with the operation of motor vehicles,
boats, winches, tools, and other Incidental equipment Boat operators should be familiar with U.S. Coast
Guard rules and regulations for safe boating contained in a pamplet, Tederai Requirements for
Redcreatlonai Boats,' available from your local U.S. Coast Guard Director or Auxiliary or State Boating
AOfficiaJ (U.S. Coast Guard. 1987).
13.2.6 Prior to a sampling trip, personnel should determine that all necessary equipment is in safe working
condition and that the operators are property trained to use the equipment
13.2.7 Safety equipment and first aid supplies must be available in the laboratory and in the field at ail times.
All boats with motors must have fire extinguishers, boat horns, cushions, and flares or communication
devices.
13.2.8 Proper field dothing should be worn to prevent hypothermia, heat exhaustion, sunstroke, drowning,
or other dangers.
13.2.9 The following safety elements should be included in the field training of field personnel:
1. Clothing and safety gear
2. Safety log for field personnel
a. itinerary (vehicle used and Its description, time of departure, travel route, estimated
time of return)
b. medical and personal information (allergies, personal health conditions)
c. personal contacts (family, telephone numbers, etc.)
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EMAP-SW-Streams, Safety and Health, Field Meth., Section 13, Revision No. 2, March 1994, Page 3 of 5
3. Emergency action
a. first aid/CPR
b. communication
c. search and rescue
4. Emergency services required near base sites
5. Safety plan schedules
13.3 SAFETY EQUIPMENT AND FACILITIES
13.3.1 Necessary and appropriate safety apparel such as waders, lab coats, gloves, safety glasses, etc.
must be available and used in accordance with the EMPA-SW for streams field operations and methods
indicators and in the processing of all laboratory samples.
13.3.2 Bright colored caps (e.g., orange) must be available and worn in ail field operations and sample
collecting activities.
13.3.3 First aid kits, fire extinguishers, and blankets must be readily available in the field, and first aid kits,
fire estinguishers, safety showers, and emergency spill kits must be available in the laboratory at all times.
13.3.4 A properly installed and operating hood must be provided in the laboratory for use when working
with carcinogenic chemicals (e.g., formaldehyde, formalin) that may produce dangerous fumes (see
Subsection 13.4.6).
13.3.5 Communication equipment and posted emergency numbers must be available to field personnel and
those working in mobile labs in remote areas for use in case of an emergency.
13.3.6 Facilities and supplies must be available for cleaning of exposed body parts that may have been
contaminated by pollutants in the water. Soap and an adequate supply of dean water or ethyl alcohol, or
equivalent, should be suitable for this purpose.
13.4 FIELD AND LABORATORY OPERATIONS
13.4.1 Two persons (three to four persons for electrofishing) must be present during all sample collection
activities, and no one should be left alone while in the field.
13.4.2 Staff training in basic first aid and cardio-pulmonary resuscitation is strongly recommended.
13.4.3 All surface waters should be considered potential health hazards due to toxic substances or
pathogens and exposure to them should be minimized as much as possible. Exposed body parts should
be cleaned immediately after contact with these waters.
13.4.4 All electrical equipment must bear the approval of Underwriters Laboratories and must be properly
grounded to protect against electric shock.
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EMAP-SW-Streams, Safety and Health, Reid Meth., Section 13, Revision No. 2, March 1994, Page 4 of 5
13.4.5 Heavy gloves should be used when hands are used to agitate the substrate during collection of kick
net or square-foot type samples and when turning over rocks during hand picking.
13.4.6 During sample or specimen processing, persons must become famBiar with the health hazards when
using fixing and/or preserving chemical agents. Note: when using formaldehyde or formalin in the field,
care must be taken because it is highly allergenic, toxic, and dangerous to human health (carcinogenic) if
utilized improperly (see Subsection 13.3.3).
13.4.7 Persons working In areas where poisonous snakes may be encountered must check with the local
Drug and Poison Control Center for recommendations on what should be done In case of a bite from a
poisonous snake. If local advice Is not available and medical assistance is more than an hour away, carry
a snake bite kit and be familiar with its use. Any person allergic to bee stings or other insect bites must take
proper precautions and have any needed medications handy.
13.4.8 Personnel participating in field activities on a regular or infrequent basis should be In sound
physical condition and have a physical exam annually or In accordance with Regional or State Safety
requirements.
13.4.9 All field personnel should be familiar with the symptoms of hypothermia and know what to do In case
symptoms occur. Hypothermia can kill a person at temperatures much above freezing (up to 10°C or 50°F)
if he or she Is exposed to wind or becomes wet
13.5 DISEASE PREVENTION
13.5.1 Unknown pollutants and pathogens in surface waters and sediments should be considered potential
health hazards and exposure to them kept to a minimum.
13.5.2 Personnel who may be exposed to water known or suspected to contain human or animal wastes
that carry causative agents or pathogens must be immunized against tetanus, hepatitis, typhoid fever, and
polio. Biological wastes can also be a threat In the form of viruses, bacteria, rickettsia, fungi, or parasites.
Field personnel should also protect themselves against the bite of deer or wood ticks because of the
potential risk of acquiring pathogens that cause Rocky Mountain spotted fever and Lyme disease,
13.6 CHEMICAL WASTES
13.6.1 Chemical wastes can cause various hazards due to flammabllity, explosibillty, toxicity, causticity, or
chemical reactivity. All chemical wastes must be discarded according to standardized health and hazards
procedures (National Institute for Occupational safety and Health. 1981; USEPA, 1986).
13.7 REFERENCES
American Red Cross. 1979. Standard first aid and personal safety. American National Red Cross. 269
pp.
Berry, C.R. Jr., W.T. Helm, and J.M. Neuhold. 1983. Safety in fishery field work. In: Nielsen, LA. and D.L
Johnson (eds.). Fisheries Techniques, American Fisheries Society, Bethesda, MD. pp. 43-60.
National Institute for occupational Safety and Health. 1981. Occupational health guidelines for chemical
hazards. NIOSH/OSHA Pub. No. 81-123. (Two Volumes). U.S. Government Printing Office.
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EMAP-SW-Streams, Safety and Health, Reid Meth., Section 13, Revision No. 2, March 1994, Page 5 of 5
Ohio EPA. 1990. Ohio EPA Fish evaluation group safety manual. Ohio Environmental Protection
Agency, Ecological Assessment Section, Division of Water Quality Planning and Assessment,
Columbus, OH.
Reynolds, J.B. 1983. Electrofishing. In: LA. Nielsen and D.L Johnson (eds.). Fisheries Techniques.
Amer. Fish. Soc., Bethesda. MD. pp. 147-163.
U.S. Coast Guard. 1987. Federal requirements for recreational boats. U.S. Department of
Transportation, United States Coast Guard, Washington, DC.
USEPA. 1986. Occupational health and safety manual. Office of Planning and Management, U.S.
Environmental Protection Agency, Washington, DC.
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EMAP-SW Streams, Info Management, Field Methods, Section 14, Rev. No. 3, April 1995, page 1 of 26
SECTION 14
INFORMATION MANAGEMENT PROCEDURES
14.1 INTRODUCTION
14.1.1 Data are collected from each site using field data recorders, sample collection and tracking, and
receipt of analytical data from laboratories. The first two methods are used at the site and are essential to
ensure that laboratory results will be available and accurately linked to each site. Without proper data and
information collection methods, data can be lost and may result in revisiting the site.
Therefore, you are a major link in making sure the Mid-Appalachian Stream Pilot is complete. The
Information Management (IM) personnel have made every effort to make all aspects of the sample
shipment and tracking system easy and as painless as possible.
14.2 Field Data Entry Methods.
The primary method of data collection will be using Portable Data Recorders (PDR). This is a
small hand held computer that has a full QWERTY keyboard, is air and water tight, will withstand being
dropped into the water and has a viable battery life to be used for field data entry. The secondary method
is paper backup (see figures 14-5 thru 14-15) for the PDR.
In both methods, each field has a representative significant figure shown and in most cases with
have the proper unit of measure. If the measuring device used is in a different unit of measure and this is
not stated somewhere, the results of the data may be unpredictable. These units and significant figures are
important to follow. Just one digit longer or a different unit of measure may result in a value being
represented incorrectly and changing the total analysis of the sampling. In all cases, please be concise in
any entries and comment on anything notable in the comment sections provided.
14.2.1 PDR data entry
The field data recorders are a simple computing device. Each of the prior years field data entry
forms have been converted from paper to electronic format and are available for field use.
The programs are organized in three different segments, general field data collection, fish collection and
physical habitat. In all cases, the general field data collection program must be entered first to establish the
sampling site. Even if the site was not sampled nor visited for any reason, enter the information on the
PDR. Once this has been done, the rest of the programs will contain the information. Most of the fields can
be changed except certain ones to protect the integrity of the connection within the data (like Stream ID,
Date of collection, Visit Number).
The programs were written to be patterned off of the 1994 Streams Pilot field forms. Each section
within the program(s) is as close as possible to the field data sheet that it represents. The same data is
being collected and will be organized in the same manner as the prior year's data. A users guide is
available that has examples of each screen and a full explanation as to the entry (see Information
Management Handbook, PDR user Instructions).
This type of data entry does not allow you full view of the data as the forms did. You need to look
at each screen and make sure that you have at least entered that section and then check it again for
completeness. You can change the data as many times as needed until the data is transferred from the
PDR to the laptop. And even then, you do have one last opportunity to modify the data on the laptop. After
the data leaves the laptop, you no longer can change it
The data is transferred from the PDR to the laptop nightly. It is essential to the sample tracking
(STARS) system. Table 14.2 has a brief description on the process, refer to the Information Management
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EMAP-SW Streams, Info Management, Field Methods, Section 14, Rev. No. 3, April 1995, page 2 of 26
Handbook for full instructions. After you have completed the sample tracking information, the data will
need to be transferred from the laptop to the ERL-Corvallis VAX computer. This process is fully automated
and can run on it's own. You first need to make sure that the connection is made from your computer to
ERL-Corvallis. After that, the two (2) computers will take it from there.
If the data fails to transfer to ERL-Corvallis or you have no way to connect your computer to a
phone, then please be sure to do a backup from the Main Menu on the laptop. This will save the data on
something else other than the computer. Then, mail the diskette at the of the week as described in table
14.2 . .
Recharge the PDR, the PDR batter and the laptop computer each night This is essential to ensure
that there will be enough power for the equipment to be used the next day. Please refer to the Information
Management Handbook for full instruction. If at any time the PDR is not available, be it failure, power
failure, or not available, use the field data sheets that are provided (see the next section, 14.22).
14.2.2 Paper data entry
The forms are printed on specially treated paper, Rite in the Rain, and are designed for the type of
work that you will be doing. They have actually been dropped in the water and have retained all entries. A
pen has been selected to ensure that the entries will not smear and/or rub off this specially treated paper.
This pen (SANFORD Sharpie) will be issued and used by all team members. Also, a soft lead pencil (NO.
2 or less) will be provided. If the form gets wet, only the pencil will write on the form. DO NOT USE ANY
OTHER TYPE PEN OR PENCIL
Depending on the type of site (EMAP or TIME), the number and type of forms needed will vary.
Section 14.5, table 14.1 A-B illustrates the type and number of forms needed for the different sites. Check
this table to see what set of forms you will need to have for your team and get a weeks supply plus a few
days each week. The forms will be supplied at the beginning of field season. If more forms of any type are
needed during the season, contact your field coordinator or IM and some will be mailed to you.
Now, on to the importance of the forms. Each form is essential to the project They provide
information on each indicator with either field measurements and/or sample collection and tracking. The
entries must be legible and within the guidelines provided on each form. Illegible information is equivalent
to NO information. If any additional information is deemed necessary for the indicator, provide this in the
comment section on the specific form of indicator concentration. Review the forms at the end of the day at
the site. This will allow you time to complete something you may have forgotten. The lead on each team
will do a final check of the forms and enter their initials at the top of each page to indicate a check was
made.
At the beginning of each week, prepare a FED-EX envelope to put each days set of forms In.
Keep the envelope in your vehicles. Keep collecting the forms for each site until the end of the week.
When you get ready to send the weeks samples that go to Newtown, again check the forms for
completeness and place the envelope on the top of all the samples so that the ice does not saturate it See
section 1413.3 on sample shipping for any further details.
Each Indicator section in the 1995 Pilot Operations and Methods Manual has information regarding
completing each field data form. The specific entries and values for the forms will be covered in field
training. Any other questions can be directed to IM and/or the indicator lead for clarification.
14.3 Field Sample Processing
The collection of samples is just as important and in some cases the most important part of the
particular indicator the sample is associated to. Proper handling, preparation and documentation of any
sample is integral to a successful stream visit Taking time to property fill out the labels, making sure that
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EMAP-SW Streams, Info Management. Field Methods, Section 14, Rev. No. 3, April 1995, page 3 of 26
the samples are collected as instructed by the protocols, and processing the samples for shipment by
tracking them will ensure safe arrival to the laboratories for analysis. This section will assist you on the
process to accomplish this.
14.3.1 Sample Labels and Preparation.
Each sample collected will have a label to identify it (see figure 14-16). Fill in any known
information (e.g., stream name, stream id, date, etc.) on each label before departing from the vehicle so
long as the stream has been determined samples will be collected. Pre-labe! and prepackage Cubitainers,
syringes, and sample containers into sample kits prior to departure. Store an extra supply kit of sampling
supplies (field data forms, sample labels, sample containers, etc.) in the vehicles. Inventory these extra
supplies daily. Sample checklists are provided in tables 14-1A-1B
14.3.2 Sample Tracking and Reporting.
The sample tracking, shipping and reporting process will be completed back at the base site
(hotel/motel/etc.). Follow the instructions for hotel/motel connection and data downloading (see the
Information Management Book for further instructions).
Sample tracking is initiated at this time by using the notebook computer, bar code reader and the
Information Management Handbook that will be provided to each crew. Table 14-2 is a brief version of this
process. Any further instructions will be provided in the Information Management Handbook.
14.3.3 Sample Shipment
First and foremost, follow the indicator protocols on proper sample collection and storage. Then
make sure all labels are completely filled out, legible and taped with clear plastic tape to prevent the label
from smearing and/or getting wet. Make sure all the samples and labels are properly prepared. Just like
the field data forms, labels are an important part of this project. If the samples are labelled incorrectly or
illegibly, the site may have to be re-sampled and/or the data thrown out!
There are three (3) different sample shipping/destination locations that will be used. Table 14-3
lists the destinations for each sample type, the shipping times, and the preservatives required. The water
chemistry is the only one that will be shipped on the day of collection. The others are either shipped or
transported once a week. In the case of samples going to the supply depots, a designated crew member
will be transporting the sample(s) there (see table 14-4 for depot locations).
14.3.3.1 Newtown shipments
Samples of fish tissue, periphyton chlorophyll, uiomass, APA, ID, sediment tubes, and sediment
toxicity bags remain with the crews until you return to EPA-Newtown. Keep the samples that need to be
kept frozen (see table 14-3) in the provided freezer. If more samples are collected than the freezer can
hold, ship samples in a cooler by FEDEX. Follow the cooler preparation as stated below, process the
samples using STARS, complete .the pre-printed FEDEX airbill and call FED-EX with the location of the
cooler for pick up. Be sure to include the weeks worth of field data sheets.
Line each shipping cooler with some kind of paper (use the provided absorbent paper or paper
towels, brown paper shopping bags or even pizza boxes work real well!). Place a garbage bag in each
cooler. Prepare several bags of ice by writing on the bags the word "ICE" in large print. Then fill each bag
with fresh ice and seal the bags. Place at least one or two bag(s) on the bottom of the cooler inside the
garbage bag. Take the samples out of the freezer, Check all sample labels and make sure the information
is correct, complete and legible. Place them on top of the ice, and then place more bagged ice on top. If
the samples and the ice do not fit. get another cooler and prepare it the same way the first one was. Seal
the garbage bag with either tieing it in a knot or using a twist tie (if available).
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EMAP-SW Streams. Info Management. Field Methods. Section 14, Rev. No. 3, April 1995, page 4 of 26
If there are field data sheets and you are making a mid-week FEDEX shipment, prepare the weeks
worth of field data sheets by placing them in the FEDEX envelope and sealing it Then place the envelope
on the top of the sealed garbage bag. For all shipments, place the packing slip in a clear plastic bag and
enclose it in the cooler before sealing. Seal the cooler with clear tape and place the completed FED-EX
airbill in the attached plastic sleeve. Call FEDEX with the drop off location. Call Newtown (see table 14-3)
to expect a Saturday shipment (for Friday shipments only).
14.3.3.2 University of Maine shipments
Water samples (cubi's/syringes) are shipped to University of Maine the day after collection. Each
cooler will have a bag of sealed ice, the samples, packing slip and a label tapped to the inside of the cooler
top with one of the re-supply depots' address for cooler return.
Line the cooler with a garbage bag. Prepare several bags of ice by writing on the bags the word
"ICE" in large print. Place the sealed bags of ice in the garbage bag. Check all sample labels and make
sure the information is correct, complete and legible. Place the water sample(s) on the ice, add more bags
of ice on top of the samples, seal the garbage bag by knot, twist tie or tape, place the packing slip in a
plastic bag on the garbage, and then attach the supply depot return card to the top of the cooler. Seal the
cooler with clear tape and place the completed FEDEX airbill in the attached plastic sleeve. Call FEDEX
with the drop off location. If the shipment date is on a Friday, call the contact person (see table 14-3) or
leave a message that a Saturday delivery is coming.
14.3.3.3 Depot shipments
Biological specimens (fish and macroinvertebrae) will be taken to a depot (see table 14-4 for
locations) by a designated crew member. Due to the length of time the samples are being stored, take a
few extra moments to be sure the sample is properly prepared for storage. Reverify the fish voucher is
preserved correctly, each perforated Ziploc bag contains a water proof label and the jar has the jar label
taped on it. The macroinvertebrae should have the waterproof label in the jar and the bar coded label
outside of the jar and taped. Be sure the packing slip of all samples taken to the specific depot This
packing slip is a tracking mechanism that will follow the sample(s) at all times.
14.4 Supplies
Depending on your team, resupplies can be acquired at your designated supply depot (see table
14-4). Coolers that were sent to the University of Maine will be sent back to one of the resupply depots.
Each team will be supplied with address cards and will be instructed at field training as to how to prepare
the coolers for return. This way you "can get more coolers and other supplies at each location. Again be -
sure to collect one weeks supply plus some extras of each item to carry you over until the next week.
14.5 Forms, Labels and Checklists.
Figures 14.1 -14.15 are the field data forms that will be used throughout the field season. Rgure
14.16 illustrates the sample labels for sample collection. Table 14-1A-B are checklists for Information
Management and stream visit needs for each site. The table itself declares which type of site it Is to be
used for. Table 14-2 is a brief description on how to run the STARS sample tracking and reporting
program. Table 14-3 provides the shipping addresses and a brief synopsis of sample shipping. Table 14-4
is a list of each states' supply depot, contact and address.
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EMAP-SW Streams, Info Management, Field Methods, Section 14, Rev. No. 3, April 1995, page 5 of 26
TABLE 14-1A STREAM VISIT CHECKLIST - EMAP, REFERENCE AND REVISIT SITES
Stream Verification Form
Stream Assessment Form
Rapid Habitat Assessment Form: Glide/Pool
Rapid Habitat Assessment Form: Riffle/Run
Channel/Riparian Cross-Section Form
Fish Collection and Sample Tracking Form
Field Measurement Form
Sample Collection Form
Information Management Book
Stream Information Packet
Sampling Permit
EMAP Pamphlet
Field Operations and Method Manual
Composite Benthos Labels - Bar Coded
Benthos Identification Label Sheet - Non Bar Coded
Fish Voucher Label Sheet - Bar Coded
Fish Voucher Label Sheet - Non Bar Coded
Fish Tissue Labels - Bar Coded
Periphyton Labels (APA, BIOMASS, CHLA, ID) - Bar Coded
Sediment Metabolism Labels - Bar Coded
Sediment Toxicity Label - Bar Coded
Water Chemistry Labels - Bar Coded
Clear Tape Gun - Loaded
Clear Tape Pad
Cheat Sheets
Computer Equipment (see Information Management Book)
Clipboard/Sharpie/pencil
Shipping Airbills
1
1
1
1
11
1
1
1
1
1
1
10
1
2
1
2
1
2
8
5
1
5
1
2
1 set
1set
1 each
2
-------
EMAP-SW Streams, Info Management, Field Methods, Section 14, Rev. No. 3, AprO 1995, page 6 of 26
TABLE 14-1B STREAM VISIT CHECKUST -TIME SITES
Stream Verification Form
Stream Assessment Form
Rapid Habitat Assessment Form: Glide/Pool
Rapid Habitat Assessment Form: Riffle/Run
Fish Collection and Sample Tracking Form
Field Measurement Form
Sample Collection Form
Information Management Book
Stream Information Packet
Sampling Permit
EMAP Pamphlet
Field Operations and Methods Manual
Composite Benthos Labels - Bar Coded
Benthos Identification Label Sheet - Non Bar Coded
Periphyton Labels (APA, BIOMASS, CHLA. ID) - Bar Coded
Water Chemistry Labels - Bar Coded
Clear Tape Gun • Loaded
\
Clear Tape Pad
Cheat Sheets
Computer Equipment (see Information Management Book)
Clipboard/Sharpie/pencil
Shipping Airbill
1
1
1
1
1
1
1
1
1
1
10
1
2
1
2
5
1
2
1 set
1set
1 each
1
-------
EMAP-SW Streams, Info Management. Field Methods, Section 14. Rev. No. 3. April 1995, page 7 of 26
TABLE 14-2 SAMPLE AND DATA TRACKING PROCEDURE
1. Upon return from the sampling site, connect the PDR and computer to the provided power strip,
plug the power strip into a wall outlet and turn all equipment on. Connect the PDR to the.
computer. Select the PDR downloading process from the menus on the PDR and the computer
(See the Information Handbook for detail instructions).
2. To begin entering sample tracking data, select the STARS menu option from the laptop menu
system. The sample tracking screen will be displayed.
2. Using the bar code scanner, scan the bar code of the corresponding sample.
3a. (PDRs used) verify the information is correct and press F4 to save the record. Repeat the process
for all samples.
3b. (No PDRs used) enter the STREAM ID, DATE, CREW ID. SAMPLE CLASS and location sample
collected. Verify the information is correct and press F4 to save the record. Repeat the process for
all samples.
4. After all data have been entered for the day, press ESC to return to STARS main menu. Select
option 2 to generate shipping forms. A menu will appear. Select to either produce a new shipping
form or reprint a previous. A list of the sample tracking numbers available for shipping will appear.
Highlight the sample(s) to be shipped in a single cooler and press SPACEBAR to select it.
5. After all samples have been selected to be printed, press ESC to start printing. Scan in the FEDEX
airbill number. Check printer to ensure that shipping and sample tracking form(s) are being
printed. A shipping form should be printed for each cooler shipped. The program will ask if the
printing was successful. If not, select reprint. The program will then ask, if there are more coolers
to be shipped. Answer Y(es) to continue and follow steps 4-5 or enter N(o) to return to STARS
main menu and exit the program.
6. Samples are ready to be shipped to the respective laboratories/locations. The shipping form(s)
should be placed in a Ziploc bag(s) and taped to the inside of the cooler top. Follow the cooler
sample packing process as described in section 14 of the fields method manual.
7. If this is one of the days to prepare a backup disk (Tuesday or Friday), place a diskette in drive A of
the computer and select the Backup selection on the Main Menu of the laptop. Drop the sealed
diskette packet into any US mail outlet on Friday of each week.
9. Last process is to select the option on the Main Menu to transfer data from the laptop to the
Corvallis vax system. Follow the instructions provided in the Information Management Handbook.
This is automatic once connection has been made.
10. Recharge all equipment by first turning the computer and PDR off. Remove the battery from the
PDR, connect the PDR and the extra battery into the power strip using the provided AC adapters.
Leave the computer plugged in along with the PDR and let them charge over night.
-------
EMAP-SW Streams, Info Management, Field Methods. Section 14, Rev. No. 3, April 1995, page 8 of 26
TABLE 14-3 SHIPPING DESTINATIONS, TIMES AND PRESERVATIVES
Media
Composite Benthos (jar)
Fish Vouchers Oar)
Fish Tissue
Periphyton - APA
Periphyton - BIOMASS
Periphyton - CHLA
Periphyton - ID
Sediment Metabolism
Sediment Toxicity
Water Chemistry (Cubitainer,
syringes)
Preservative
Ethanol Solution
Formalin
Freezer/ Ice
Freezer/ Ice
Freezer/ Ice
Freezer/ Ice
Freezer/ Ice
Freezer/ Ice
Freezer/ Ice
Ice in Ziplocs
Shipping Times*
Upon arrival to the
depot center.
Upon arrival to the
depot center.
Weekly to Newtown"
Weekly to Newtown"
Weekly to Newtown"
Weekly to Newtown"
Weekly to Newtown"
Weekly to Newtown"
Weekly to Newtown"
Day of collection
Destination
Depot supply center.1
Depot supply center.1
US EPA -Newtown2
US EPA -Newtown2
US EPA - Newtown2
US EPA -Newtown2
US EPA -Newtown2
US EPA -Newtown2
US EPA - Newtown2
University of Maine1
• It may be necessary to ship items the following day if sampling activities pass the time for courier pickup of
dropoff. Make sure samples requiring ice remain frozen and in darkness until shipment
" there may be a mid-week shipment if the freezers become full. Otherwise, transport the samples to
Newtown.
1. Depot supply center.
2. US EPA-Newtown
3411 Church Street
Cincinnati, OH 45244
(513)569-7075
ATTN:EMAP-JimLazorchak
3. Sawyer Environmental Research Center
2nd Floor
University of Maine
Orono, ME 04469
(207) 581-3287
ATTN: Therese Anderson
-------
EMAP-SW Streams, Info Management, Field Methods, Section 14, Rev. No. 3, April 1995, page 9 of 26
TABLE 14-4 SUPPLY DEPOTS
DEPOT
CLEARFIELD
ELKINS
St. ALBINS
WHEELING
WILKES-
BARRE
CONTACT
Wayne Wynick
Bruce Evans
Janice Smithson
Jim Green
Ed Kupsky
ACCESS
M-F
8AM-5PM
M-F
7am-5pm
Call Ahead
Call Ahead
M-F
7am-5pm
PHONE #
814-765-
3741
304-637-
0245
304-759-
0701
304-234-
0243
717-826-
2553
ADDRESS
PA DER Bureau of Forestry
Moshannon State Forest,
RD1
Penfield.PA 15849
WVDEP
Ward Road
Elkins, WV 25241
WVDep
694 Wmfield Rd.
St.Albans, WV25177
US EPA - Wheeling Office
303 Methodist Bldg.
Wheeling, WV 26003
PA DER Water Management
Program
2 Public Square
Wilkes-Barre, PA 18711
-------
EMAP-SW Streams, Information Management, Reid Methods, Section 14,Rev. No. 2, page 11 of 26
R«vww*d by (initial):
- -.P.'.' -V *•»'•
•*-/*-! . -i .*" -
WATOl OARTTYiD CUM O
VBUALASSESSMBTT:
RIHAUJTY:
O OMU*"
O*.
COMB
M uacus:
OVERALL BOTIC KTEOWTY O tmmtm*
|mtM«Z.»4l
STREAM ASSESSMENT FORM - 1
Figure 14.1. Stream Assessment Form -1.
-------
EMAP-SW Streams. Information Management. Reid Methods, Section H,Rev. No. 2. page 12 of 26
(MtW):
STREAM NAME:
DATE OF VISIT:
/ / 94 vttfTft
113
STREAM ID:
TEAM 10 (dntol: 1 2348678
SAMP* ?^fU?
a Itogutar • Rowing
o bitflnnittMii • OHT
a Dry - No W«ttr Anywhw*
a Alt»nd - Strum PI Mint bM not M on HMO
a Otiwr (•zpWn In
O NO
on WAI
MMUO/POMD)
ONOTW/
> 1 U M» OW NMf OF MAC*
OA<
ai
HfORMATTOM
MAW. *«O UCTQ4 GBIOM.
Itov. 3/1U»4 Umwvt.MI
Stream Verification Form- 1
Figure 14.2. Stream Verification Form • 1.
-------
EMAP-SW Streams, Information Management, Field Methods, Section 14,Rev. No. 2, page 13 of 26
by (WtW):
Stravn Vwifieation Form- 2
Rgure 14i Stream Verification Form - 2 (Continued).
-------
EMAP-SW Streams, Information Management, ReW Methods. Section HRev. No. 2, page 14 of 26
STREAM NAME:
DATE OF VISIT:
TEAM ID (drctol: 12348678
M*. 3/1WM (tMM1.S4l
FMdMt
n Fomv SutBini • 1
Rgure 14.3. Reid Measurement Form-Streams -1.
-------
EMAP-SW-Streams, Rapid Habitat Stream Assessment Forms, Reid, Sec. 15, Rev. No. 2, Page 5 of 5
15.2.3.1 The goal of EMAP-SW is to assess three major ecologicaJ values with respect to streams:
Trophic State, Fishability, and Biotic Integrity. To this end, any comments field crews can make in
assessing these values, based on their field experience, would be most valuable. Comments on these
values should be written in this section. The keywords on the left side of this section are there to
stimulate thought and are not comprehensive, nor do they all have to be addressed. In this section, we
are looking for an assessment of these values based on the field visit
Trophic State - Is defined as 'rate or amount of aloae & macrophvtes produced or present In a stream*.
Listing any observed potential nutrient sources to the stream (e.g. septic tanks, agricultural runoff). Give
your visual impression of trophic status as either low productivity (little/no blomass In streamwater),
moderate productivity (intermediate amounts of biomass in streamwater), or high productivity (large
amounts of biomass in streamwater).
Fishability - is defined as "a fish assemblage containing fish that are catchable. desirable, and safe to
consume bv wildlife and humans'. Write down any observations about fishabflity; Impressions of fish
habitat, conversations with locals, presence of fish, fishermen.
Biotic Integrity - is defined as the ability to support and maintain a balanced, integrated, adaptive
community with a biological diversity, composition, and functional organization comparable to natural
streams of the region'. Record your overall impression of the "health" of the biota in the stream. The
presence of higher order consumers (fish eating birds, mammals) is an indication of a healthy food web
and could be noted here. Similarly, the lack of an organism that you expected to see would be an
important observation. Also, any input on possible causes of impairment would be appropriate.
Water Body Character - is defined as the physical habitat Integrity of the water body, lamely a
function of riparian and littoral habitat structure, volume change, trash, turbidity, slicks, scums, color, and
odor*. We're attempting (pilot) to define water body character through two attributes, degree of human
development, and aesthetics. We'd like you to rate each of these attributes on a scale of 1 to 5. For
development, give the stream a "5" if It is pristine, with no signs of any human development A1* would
indicate a stream is totally developed,.for example the entire stream is lined with houses, or the riparian
zone has been removed. For aesthetics, base your decision on any factor about the stream that bothers
you (trash, algal growth, weed abundance, overcrowding). Circle the number that best describes your
opinion on how suitable the stream water is for recreation and aesthetic enjoyment today:
5. Beautiful, could not be any nicer.
4. Very minor aesthetic problems; excellent for swimming, boating, enjoyment.
3. Enjoyment impaired.
2. Level of enjoyment substantially reduced.
1. Enjoyment nearly impossible.
Add any comments you feel might aid data interpretation in the Comments Section.
-------
EMAP-SW-Streams, Rapid Habitat Stream Assessment Forms, Field, Sec. 15. Rev. No. 2. Page 3 of 5
entering a stream from runoff, controls erosion, and provides stream habitat and nutrient input into the
stream. A relatively undisturbed riparian zone reflects a healthy stream system; narrow, far less useful
riparian zones occur when roads, parking lots, fields, lawns, bare son, rocks, or buildings are near the
stream bank. The presence of "old fields' (i.e., a previously developed field allowed to convert to natural
conditions) will rate higher than fields in continuous or periodic use. Paths and walkways in an
otherwise undisturbed riparian zone may be judged to be inconsequential to destruction of the riparian
zone.
15.1.4 Pool/Glide Prevalence Form
15.1.4.1 Instream Cover for Fish (see Subsection 15.1.3.1)
15.1.4.2 Epifaunal Substrate for Macroinvertebrates, In muddy bottom streams are mostly on
submerged logs or snags, and aquatic vegetation (also see Subsection 15.1.3.2)
15.1.4.3 Pool Substrate Characterization evaluates the type and condition of bottom substrates found in
pools. Firmer sediment types (e.g., gravel, sand) and rooted aquatic plants support a wider variety of
organisms than a pool substrate dominated by mud or bedrock and no plants. In addition, a stream
that has a uniform substrate in its pools will support far fewer types of organisms than a stream that has
a variety of substrate types.
15.1.4.4 Pool Variability rates the overall mixture of pool types found In streams, according to size and
depth. The four basic types of pools are large-shallow, large-deep, small-shallow, and small-deep. A
stream with many pool types wDI support a wide variety of aquatic species. Rivers whh low sinuosity
(few bends) and monotonous pool characteristics do not have sufficient quantities and types of habitat
to support a diverse aquatic community. As a general guideline: consider a pool deep is ft is greater
than 1 m deep, and large if ft length, width, or oblique dimension is greater than half the stream width.
15.1.4.5 Channel Alteration (see Subsection 15.1.3.5)
15.1.4.6 Sediment Deposition (see Subsection 15.1.3.6)
15.1.4.7 Channel Sinuosity evaluates the meandering or relative frequency of bends of the stream.
Streams that meander provide a variety of habitats for aquatic organisms, whereas straight stream
segments are characterized by monotonous habitats that are prone to flooding. A high degree of
sinuosity creates a variety of pools and reduces the energy from surges when the stream flow fluctuates.
The absorption of this energy by bends protects the stream from excessive erosion and flooding. In
•oxbow* streams of coastal areas and deltas, meanders are highly exaggerated and transient Natural
conditions are shifting channels and bends. Alteration of these streams is usually In the form of flow
regulation and dlversloa
15.1.4.8 Channel Flow Status determines the percent of the channel that is filled with water. The flow
status will change as the channel enlarges or as flow decreases as a result of dams and other
obstructions, diversions for Irrigation, or drought The water will not cover as much of the streambed,
thus decreasing the amount of living areas for aquatic organisms. In muddy bottom streams, the
decrease in water level wfli expose logs and snags, thus reducing the areas with good habitat
15.1.4.9 Condition of Banks (see Subsectlon15.1.3.9).
15.1.4.10 Bank Vegetative Protection (see Subsection 15.1.3.10).
-------
EMAP-SW-Streams, Rapid Habitat Stream Assessment Forms, Reid, Sec. 15, Rev. No. 2. Page 4 of 5
15.1.4.11 Grazing/Disruptive Pressure (see Subsection 15.1.3.11).
15.1.4.12 Riparian Zone Vegetative Width (see Subsection 15.1.3.12).
15.2 STREAM ASSESSMENT FORM
15.2.1 The objective of the stream assessment form is to record field crew observations of
catchment/stream characteristics useful for future data interpretation, ecological value assessment,
development of associations, and verification of stressor data. Observations and impressions of field
crews are extremely valuable. Thus, it is important that these observations about stream characteristics
be recorded for future data interpretation and validation. This form is designed as a template for
recording pertinent field observations. It is by no means comprehensive and any additional observations
should be recorded In the Comments sectloa It Is meant to be completed at the end of the sampling,
taking into account all observations the sampling crew has made while on site. The form consists of two
major sections: Site Activities/Disturbances, and Environmental Value Assessment (see Section 14,
Information Management for forms).
15.2.2 Site Activities/Disturbances
15.2.2.1 Record any of the following stressors that were observed either while on the stream or
driving/walking through the stream catchment For activfties/stressors that you observe, rate them as to
abundance or Influence as Low, Medium, or High by putting an L, M, or H In the line next to the listed
stressor. Leave the line blank for any stressor not observed. The distinction between low, medium, and
high wfll be subjective. For example. If there are 2-3 houses on a stream, I would mark the Houses line
with a V for low. If the stream Is in a suburban housing development, I'd rate it as high (H). Similarly,
a small patch of dear cut logging on a hfll overlooking the stream, would rate a low ranking. Logging
activity right on the stream shore, however, would get a high disturbance ranking. The form is broken
down into the following Major Categories:
Residential: Note the presence of any of the listed disturbances adjacent to or near the
stream.
Recreational: Record in this section the presence of organized public or private parks,
campgrounds, beaches or other recreation areas around the stream. If there
are signs of Informal areas of camping, swimming or boating around the stream
(e.g.. swimming hole), record them as 'primitive" parks, camping.
Agriculture: Note the presence of cropland, pasture, orchards, poultry, and/or livestock.
Industrial: Note any industrial activity (e.g., canning, chemical, pulp), commercial activity
(stores, businesses) or logging/mining activities around the stream or in the
catchment Describe in more detail in the Comments section.
Management Note any evidence of liming activity, water treatment, dredging or
channelization, flow control structures, etc.
Any oddities, or further elaboration should be recorded in the Comments section.
15.2.3 Environmental Value Assessment
-------
EMAP-SW-Streams, Rapid Habitat Stream Assessment Forms, Reid, Sec. 15, Rev. No. 2, Page i of 5
SECTION 15
RAPID HABITAT AND STREAM ASSESSMENT FORMS
15.1 RAPID PHYSICAL HABITAT ASSESSMENT FORM
15.1.1 The habitat assessment approach used In this protocol Is adapted from EPA's Rapid
I .assessment approach and refined from various applications across the country. The approach
f.ojses on Integrating information from specific parameters on the structure of the physical habitat The
field data sheets that summarize the major attributes of each parameter are shown in Section 14,
Information Management. Specific instruction and training are necessary for an adequate assessment of
habitat quality. For each of the parameters listed on the form, cirde the point score that best describes
the condition of the survey reach based on the described categories.
15.1.2 Different assessment forms are used for streams that are riffle/run prevalent versus those
that are pool/glide prevalent. After making the initial survey of the reach, classify the stream as either
riffle/run or pool/glide prevalent based on your visual Impression of the dominant habitat type. Choose
the prevalent habitat type based on which habitat occupies the majority of the reach length. Then fill out
the habitat assessment form corresponding to the chosen habitat type.
15.1.3. Riffle/Run Prevalence Form
15.1.3.1 Instream Cover for Fish includes the relative quantity and variety of natural structures in the
stream, such as fallen trees, logs, and branches, large rocks, and undercut banks, that are available for
refugia, feeding, or laying eggs. A wide variety of submerged structures in the stream provides the f.:. =
with a large number of niches, thus increasing the diversity.
15.1.3.2 Epifaunal Substrate for Macrolnvertebrates are essentially the amount of niche space or hard
substrates (rocks, snags) available for insects and snails. Numerous types of Insect larvae attach
themselves to rocks, logs, branches, or other submerged substrates. As with fish, the greater the variety
and number of available niches or attachments, the greater the variety of Insects in the stream. Rocky-
bottom areas are critical for maintaining a healthy variety of insects in most high gradient streams.
15.1.3.3 Embedded/iess refers to the extent to which rocks (gravel, cobble, and boulders) are covered
or sunken in to the sBt, sand, or mud of the stream bottom. Generally, as rocks become embedded, the
surface area available to macroinvertebrates and fish (shelter, spawning, and egg Incubation) Is
decreased. To estimate the percent of embeddedness, observe the amount of silt or finer sediments
overlying and surrounding the rocks. If kicking does not dislodge the rocks .or cobble, they may be
greatly embedded. It is useful to observe the extent of the dark area on their underside of a few rocks.
15.1.3.4 Varying Velocity and Wafer Depth examine the avaflabOlty of each of the four primary
current/depth combinations: (1) slow-deep, (2) slow-shallow, (3) fast-deep, and (4) fast-shallow. The
best streams in high gradient regions wOl have all four habitat types present The presence or availability
of these four habitats relates to the stream's ability to provide and maintain a stable aquatic environment
The general guidelines are 0.5 m depth to separate shallow from deep, and 0.3 m/sec to separate fast
from slow.
-------
EMAP-SW-Streams, Rapid Habitat Stream Assessment Forms, Reid. Sec. 15. Rev. No. 2, Page 2 of 5
15.1.3.5 Channel Alteration is basically a measure of large-scale changes in the shape of the stream
channel. Many streams in urban and agricultural areas have been straightened, deepened, or diverted
into concrete channels, often for flood control purposes. Such streams have far fewer natural habitats
for fish, macroinvertebrates. and plants than do naturally meandering streams.. Channel alteration is
present when the stream runs through a concrete channel; when artificial embankments, riprap, and
other forms of artificial bank stabilization or structures are present; when the stream is very straight for
significant distances; when dams and bridges are present, and when other such changes have occurred.
15.1.3.6 The Sediment Deposition parameter measures the amount of sediment that has accumulated
and the changes that have occurred to the stream bottom as a result of the deposition. Deposition
occurs from large-scale movement of sediment caused by watershed erosion. Sediment deposition may
cause the formation of islands, point bars (areas of increased deposition usually at the beginning of
meanders that Inuease in size as the channel is diverted toward the outer bank) or shoals, or result in
the filling of pools. Increased sedimentation also results in increase deposition. Usually this Is evident in
areas that are obstructed by natural or man-made debris and areas where the stream flow decreases,
such as bends. High levels of sediment deposition create an unstable an continually changing
environment that becomes unsuitable for many organisms.
15.1.3.7 Frequency of Riffles is a way to measure the sequence of riffles occurring in a stream. Riffles
are a source of high-quality habitat and diverse fauna, therefore, an Increased frequency of occurrence
greatly enhances the diversity of the stream community. For areas where riffles are uncommon, a
run/bend ratio can be used as a measure of sinuosity. A large degree of sinuosity provides for diverse
habitat and fauna, and the stream is better able to handle the high energy flows that result from storms
than are relatively straight streams.
15.1.3.8 Channel Flow Status is the degree to which the channel Is fined with water. The flow status will
change as the channel enlarges or as flow decreases as a result of dams and other obstructions,
diversions for Irrigation, or drought When water does not cover much of the streambed, the amount of
useable substrate for aquatic organisms is limited.
15.1.3.9 Condition of Banks measures whether the stream banks are eroded (or have the potential for
erosion). Steep banks are more likely to collapse and suffer form erosion than are gentry sloping banks
and are therefore considered to be unstable. Signs of erosion include crumbling, unvegetated banks,
exposed tree roots, and exposed sol
15.1.3.10 Bank Vegetative Protection measures the amount of the stream bank that is covered by
vegetation. The root systems of plants growing on stream banks help hold soB in place, thereby
reducing the amount of erosion that is likely to occur.. This parameter supplies Information on the ability
of the bank to resist erosion, as well as some additional information on the uptake of nutrients by the
plants, the control on instream scouring, and stream shading. Banks that have full, natural plant growth
are better for fish and macroinvertebrates than are banks without vegetative protection or those shored
up with concrete or riprap.
15.1.3.11 Grazing/Disruptive Pressure is a measure of disruptive changes to the riparian zone because
of grazing or human interference (e.g., mowing). In areas of high grazing pressure from livestock or
where residential and urban development activities disrupt the riparian zone, the growth of a natural
plant community is impeded. Residential developments, urban centers, golf courses, and rangeland are
the common causes of anthropogenic effects on the riparian zone.
15.1.3.12 The Riparian Vegetated Zone Width measures the width of natural vegetation from the edge of
the stream bank (riparian buffer zone). The riparian vegetative zone serves as a buffer zone to pollutants
-------
EMAP-SW Streams, information Management, Field Methods, Section HRev. No. 2, page 25 of 26
Rcvwwod by (Modi:
WFFLE/RUN-
HABITAT PARAMETER
CATEGORY
7.
O»R»*US
SCORE:
oMdod by Ow wtth •(
tmmoojuofc It*
7* *
•
by ««•«••*•» «f
7tt Ifc
v •• AM <*«w or
poor
by th» width ol «M
Pun 25.
1lto2t..
SCORE:
w«
of both tank* and •
(• U «• 71% •( «•
Vory M* mmm In ctwnnol.
•nd monry oroum u
SCORE: f
Mote
•0% otowfc* In raoert
..•Oio
100%ofbmkho*
10. kMKVHRATIWI
SCORE:
Don C0% •< «•
11.
12.
ZOW WBtM (UACT
SCORE:
12
it
Zen* wttdi to tatwoon •
•^•^ •• m~ tWo^^^ o^K«rfw^wio^
W •* Bl! nBR^MI VBvWOTl
••nno*
MArrO HAWTAT ASSCSSMBIT FORM: KmC/RUN- STREAMS • 2
Rgure 14.9. Rapid Habitat Assessment Form: Riffle/Run-Streams - 2 (Continued).
-------
EMAP-SW Streams, Information Management, Reid Methods, Section U, Rev. No. 2, page 26 of 28
FISH-JAR
STREAMED: _. S
DATE: / / 93
SAMPLE TYPE: UKKHOWM vouam
i nm JIB HUD
383898
nSH VOUCHER-BAG
STItfiAMID: S
DATE: ___ / _ / 93
PBHCODB;
[
Ł63838
FISH TISSUE
STREAM ID: S STREAM ID: S
DATE:_/_/04 DATE:_/_/94
HABITAT: RIFFLE POOL
100200
WATER CHEMISTRY
STREAM ID: S
OATE:__/_/04
PERtPHYTON - APA
10O2O1
PCfUPHYTON - BIOUASS
10020O
STREAM ID: S
DATE:__/_/Q4
HABITAT: RIFFLE POOL
PERIPHYTON . CHLA
1OQ201
PERIPHYTON - ID
OATEt _/__/••
HAMTAT: fOOL
MBSAMPLEVOUME:
EV
"*•
10O2OO
BARCODE NO.:.
1OO2O1
STREAM ID:
DATE:_/_/84
STATION:
S
SEDIMENT METABOLISM
STREAM ID: S
DATE:_ /_/04
SAMPLE TYPE: mm KM*
PERIPHYTON
BOMASS / D / CHLA / APA
tnCAMO: •
DATE: / /•»
MAHTAT: POOL
mmi:
ACTUAL VOUMB:
100201 100201
Figure 14.10. Examples of Indicator Sample Labels.
TOTAL VOLUME:
BARCOOCfto^.
-------
EMAP-SW Streams, Information Management. Field Methods, Section 14,Rev. No. 2, page 23 of 26
**••»»«d by (inredl:
HABITAT PARAMETER
•ngrh 3 M 4 *nM
ong*r Oun H k «u In •
V«V M* •«•( in chum)
I. CMAMO. Mm STATU*
u» to
•0% •< taMn taraMh tew
rta
It
12. ftMMMlVHITA
ZOM WBTMIUMT
KAPIO HAHTAT AttCSSMOtT POMI: OUOE/POOL- S1KEAMS • 2
Figure 14.8. Rapid Habitat Assessment Form: Glide/Pool-Streams - 2 (Continued).
-------
EMAP-SW Streams, Information Management, Reid Methods, Section HRev. No. 2. page 24 of 26
*•***•* br (MM):.
1 2 S 4 I • T t
KAMO MAHTAT AilBSMBUT FORM: MFREmUN- STMAMS • t
Rgure 14.9. Rapid Habitat Assessment Form: Riffle/Run-Streams -1.
-------
EMAP-SW Streams, Information Management, Reid Methods, Section 14,Rev. No. 2, page 21 of 26
-
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EMAP-SW Streams, Information Management, Reid Methods, Section HRev. No. 2, page 22 of 26
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EMAP-SW Streams, Information Management, Field Methods, Section 14,Rev. No. 2, page 19 of 26
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EMAP-SW Streams, Information Management, Field Methods, Section HRev. No. 2, page 17 of 26
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EMAP-SW Streams, Information Management, Field Methods, Section 14,Rev. No. 2, page 18 of 26
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EMAP-SW Streams, Information Management, Field Methods, Section HRev. No. 2, page 15 of 26
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EMAP-SW Streams, Information Management. Reid Methods, Section 14 Rev. No. 2, page 16 of 28
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DATE OP VISIT: / / 94
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