United States
Environmental Protection
Agency
Office of Research and
Development
Washington, DC 20460
EPA/620/R-06/002
February 2006
\
Great River Ecosystems
Field Operations Manual
•ft PSfe • i
Environmental Monitoring and
Assessment program
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EPA/620/R-06/002
February 2006
ENVIRONMENTAL MONITORING AND ASSESSMENT PROGRAM
GREAT RIVER ECOSYSTEMS (EMAP-GRE)
FIELD OPERATIONS MANUAL
Ted R. Angradi1 (editor), E. William Schweiger5, Brian H. Hill1, David W. Bolgrien1,
James M. Lazorchak2, Erich B. Emery3, Terri M. Jicha1, Jeff A. Thomas3,
Donald J. Klemm2, Spence A. Peterson4, David M. Walters2, Brent R. Johnson2,
and Mark Bagley2
1U.S. Environmental Protection Agency
Office of Research and Development
National Health and Environmental Effects Research Laboratory
Mid-Continent Ecology Division
Duluth, MN 55804
2U.S. Environmental Protection Agency
Office of Research and Development
National Exposure Research Laboratory
Ecological Research Division
Cincinnati, OH 45268
3Ohio River Valley Water Sanitation Commission
Cincinnati, OH 45228
4U.S. Environmental Protection Agency
Office of Research and Development
National Health and Environmental Effects Research Laboratory
Western Ecology Division
Corvallis, OR 97333
5National Park Service
1201 Oakridge Drive
Fort Collins, CO 80525
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NOTICE
The information in this document has been funded by the U. S. Environmental Protection
Agency. It has been subjected to review by the National Health and Environmental Effects
Research Laboratory and approved for publication. Approval does not signify that the contents
reflect the views of the Agency, nor does mention of trade names or commercial products
constitute endorsement or recommendation for use.
Suggested citation for this document:
Angradi, T.R. (editor). 2006. Environmental Monitoring and Assessment Program: Great River
Ecosystems, Field Operations Manual. EPA/620/R-06/002. U.S. Environmental
Protection Agency, Washington, D.C.
ACKNOWLEDGMENTS
The following people contributed to the development of this manual: Tony Olsen, David Peck,
Marlys Cappaert, Suzanne San Romani, Jana Seeliger, John Chick, Terry Dukerschein, Heidi
Langrehr, Yao Yin, Jeffery Jack, Anthony Aufdenkampe, Paul Bukaveckas, Stephen Porter,
John Sullivan, Barry Poulton, Suzanne Femmer, Deb Taylor, Robert Jacobson, Allan
Batterman, Frank McCormick, Mary Ann Starus, Mary Moffet, Corlis West, and all the EMAP-
GRE sampling crews who helped us test and refine the methods described herein.
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SECTION AUTHORS
Addresses for authors are provided in each section.
Section 1: Ted R. Angradi, David W. Bolgrien, Terri Jicha, E. William Schweiger, and Brian
H. Hill
Section 2: Ted R. Angradi
Section 3: Ted R. Angradi and Terri M. Jicha
Section 4: E. William Schweiger, Ted R. Angradi, and David W. Bolgrien
Section 5: Terri M. Jicha, Ted R. Angradi, and Brian H. Hill
Section 6: Ted R. Angradi and E. William Schweiger
Section 7: E. William Schweiger and Ted R. Angradi
Section 8: Erich B. Emery, Jeff A. Thomas, Mark Bagley, and Ted R. Angradi
Section 9: James M. Lazorchak, Erich B. Emery, David M. Walters, and Spence A.
Peterson
Section 10: Ted R. Angradi, Donald J. Klemm, Jim M. Lazorchak, and Brent R. Johnson
Section 11: Brian H. Hill and James M. Lazorchak
in
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IV
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TABLE OF CONTENTS
Section Page
NOTICE ii
ACKNOWLEDGEMENTS ii
SECTION AUTHORS iii
ACRONYMS, ABBREVIATIONS, AND MEASUREMENTS UNITS vi
SECTION 1 - INTRODUCTION 1
SECTION 2 - OVERVIEW OF FIELD OPERATIONS 17
SECTION 3 - BASE LOCATION ACTIVITIES 31
SECTION 4 - SITE VERIFICATION 59
SECTION 5 - WATER CHEMISTRY AND PLANKTON 89
SECTION 6 - AQUATIC VEGETATION 109
SECTION 7 - RIPARIAN HABITAT 123
SECTION 8 - FISH 149
SECTION 9 - FISH TISSUE CONTAMINANTS 177
SECTION 10 - BENTHIC MACROINVERTEBRATES 187
SECTION 11 - PERIPHYTON AND SEDIMENT 207
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ACRONYMS, ABBREVIATIONS AND MEASUREMENTS UNITS USED IN THIS DOCUMENT
Acronyms and abbreviations
AFDM ash-free dry mass
AFS American Fisheries Society
ALK alkalinity
AP aquatic plant
BPJ best professional judgment
CENR (White House) Committee on the Environment and Natural Resources
CHL chlorophyll
CPR cardio-pulmonary resuscitation
CFR Code of Federal Regulations
DBH diameter at breast height
DC direct current
DELT deformities, erosions, lesions, and tumors
DFS distance from shore
Dl de-ionized
DLG digital line graph
DNR Department of Natural Resources
DO dissolved oxygen
EERD Ecological Exposure Research Division
EMAP Environmental Monitoring and Assessment Program
EMAP-SW EMAP-Surface Waters Resource Group
EMAP-WP EMAP Western Pilot Study
EPA U.S. Environmental Protection Agency
GCM geochemical markers
GPS Global Positioning System
GIS Geographic Information System
GRE Great River Ecosystems
HOPE high density polyethylene
ID identification
IM information management
INHS Illinois Natural History Survey
IP invasive plant
VI
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LTRMP Long Term Resource Monitoring Program
LWD large woody debris
MCS main channel shoreline
MDC Missouri Department of Conservation
MED Mid-Continent Ecology Division
MSDS Materials Safety Data Sheet
NAWQA National Water-Quality Assessment Program
NHD National Hydrography Database
NERL National Exposure Research Laboratory
NHEERL National Health and Environmental Effects Research Laboratory
NIOSH National Institute for Occupational Safety and Health
NTU nephelometric turbidity units
ORD Office of Research and Development
OSHA Occupational Safety and Health Administration
PDF portable document format
PE polyethylene
PFD personal flotation device
QA quality assurance
QCCS quality control check sample
REMAP Regional Environmental Monitoring and Assessment Program
SAV submersed aquatic vegetation
SL standard length
SMSU Southwest Missouri State University
SOP standard operating procedure
TBD to be determined
TOC total organic carbon
TSS total suspended solids
TL total length
UMR Upper Missouri River
UN United Nations
UMESC Upper Midwest Environmental Sciences Center
USCG U.S. Coast Guard
USEPA U.S. Environmental Protection Agency
USDA U. S. Department of Agriculture
USGS U. S. Geological Survey
VHP very high frequency
VII
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Acronyms and abbreviations, continued
VSS volatile suspended solids
WAAS wide-area augmentation system
WCC water chemistry composite
WED Western Ecology Division
YOY young of the year
YSI Inc Yellow Springs Instruments, Incorporated
Measurement units
amps amperes
C degrees Celsius
cm centimeter
g gram
gal gallon
ha hectare
Hz Hertz
km kilometer
L liter
m meter
m2 square meters
mg/L milligram per liter
mL milliliter
mm millimeter
um micrometer
uS/cm microsiemens per centimeter
msec millisecond
ppm parts per million
psi pounds per square inch
V volts
VA volt-ampere
VIII
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Section 1
Introduction
Ted R. Angradi1, David W. Bolgrien1, Terri Jicha1, E. William Schweiger2, and Brian H. Hill1
This manual describes procedures for collecting samples and field measurements for
biotic assemblages and abiotic characteristics of the Great Rivers of the Central Basin of the
United States: the Missouri, Upper Mississippi, and Ohio Rivers. The purpose of this manual is
to document the field procedures to be used in the Environmental Monitoring and Assessment
Program for Great River Ecosystems (EMAP-GRE). In addition to the technical and logistic
aspects of field operations, this manual emphasizes health and safety considerations, data
quality assurance (QA), and information management (IM). Not included in this manual are
laboratory protocols, sample-design details, or protocols for data analysis or interpretation.
The procedures in this manual are based on a variety of sources, including previously
published EMAP manuals (e.g., Baker et al. 1997, Strobel and Heitmuller 2001, Peck et al.,
unpublished drafts), other EPA documents (Klemm et al. 1990, Barbour et al. 1999, Kaufmann
et al. 1999, Flotermersch et al. 2000), an unpublished Upper Missouri River EMAP Field
Operations Manual (Angradi et al. 2002), the unpublished findings of an EPA-sponsored
workshop on Great River indicators (Angradi and Hill 2003), USGS NAWQA (Moulton et al.
2002) and USGS LTRMP protocols (e.g., Yin et al. 2000), the scientific literature, and the
experience and professional judgement of section contributors.
This manual is designed to be a comprehensive set of required procedures and
checklists for EMAP-GRE field sampling. This manual is also an informal safety and QA
guidance document for field crews; it is a reference document for planning EMAP-GRE field
operations, including office-based reconnaissance activities; it is a training tool for EMAP-GRE
crews; and it serves as documentation of methods in support of EMAP-GRE publications and
products.
1 U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
2 National Park Service, 1201 Oakridge Drive, Fort Collins, CO 80525
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Earlier versions of this manual were used in the 2004 and 2005 field seasons. The
current version of the manual supports ongoing EMAP sampling for EMAP-GRE and other large
river sampling programs.
1.1 Overview of EMAP
EMAP is a long-term research program focused on developing indicators and unbiased
statistical designs for assessing the condition of aquatic ecosystems at a variety of spatial
scales (USEPA 2002). Ecosystems included in past EMAP efforts have included lakes,
estuaries, wetlands, and surface waters (wadeable and non-wadeable streams). The two main
goals of EMAP (USEPA 2002) are relevant to the selection of indicators and approaches for
assessment and to the field operations that generate the data upon which the assessments are
based:
Develop the science needed for a state-based statistical monitoring framework to
detect trends in condition of the Nation's aquatic ecosystems. EMAP indicators are
chosen for their usefulness in revealing the condition of aquatic ecosystems. Field
operations must be designed so that the samples and measurements are collected in a
manner that maximizes the reliability and sensitivity of the indicators. This includes the
equipment, procedures, and personnel used; the spatial arrangement of samples; the
timing of data collection and the habitats in which data are collected.
Transfer EMAP science and technology to the states, tribes, and EPA regions.
Indicators and methods developed for use by states and tribes must have attributes that
favor their adoption. These attributes include cost-efficiency, public acceptance, and
relevance to management. High quality assurance (QA) during field operations is also
central to this EMAP goal since only data sets with known performance characteristics
can be efficiently combined and shared (Barbour et al. 1999).
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1.2 Overview of EMAP-GRE
1.2.1 Objectives and scope
The objectives of EMAP-GRE are to 1) develop monitoring and assessment tools for the
Great Rivers of the Central Basin: the Upper Mississippi, Missouri, and Ohio Rivers; and 2) to
demonstrate those tools in a regional assessment of Great River ecosystem condition. Because
of the size and complexity of these river ecosystems, they represent a major assessment
challenge for states and tribes. In general, robust GRE monitoring and assessment tools
proven to be useful across the region are not yet available.
EMAP-GRE will address 3 general questions:
1. What proportion of the GREs of the Central Basin, expressed in river miles, are
in good, fair, and poor condition?
2. What is the extent of aquatic, floodplain, and riparian habitat in the GREs of the
Central Basin?
3. What is the relative importance of stressors (e.g., bank stabilization, excess
nutrients, metals, invasive species, riparian disturbance) in the GREs of the
Central Basin?
The resource population of interest for the EMAP-GRE assessment is selected habitats
(described below) of the Missouri River from Fort Peck Dam in Montana to the confluence with
the Mississippi River, the Mississippi River from Lower St. Anthony Falls in Minneapolis to the
confluence with the Ohio River, and the Ohio River from the confluence of the Monongahela
and Allegheny Rivers to the confluence with the Mississippi River (Figure 1-1). Fifteen states
are included within the scope of EMAP-GRE.
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EMAP-GRE Strata
*4^** Mississippi. Missouri and Ohio Rivers
-'\y--~ Missouri Rive
r Reserve r (not in design)
1
•"•— t. W
\^Jn
"
Figure 1-1. Geographic scope of EMAP-GRE. Mainstem Missouri River reservoirs are
excluded from EMAP-GRE.
1.2.2 The EMAP-GRE sample design
The details of the EMAP-GRE sample design are beyond the scope of this manual.
This manual describes activities that occur after field personnel have the "design file" in hand.
The design file is a spreadsheet containing the locations and support information of all
candidate sample sites for each river. The design file is discussed in Section 4 of this manual.
1.2.3 The index period
The index period is the period of the year in which sampling may occur. The index
period for EMAP-GRE is July 1 - September 30. This period corresponds to a season of high
biological activity, clement weather, and relatively stable flows.
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1.2.4 EMAP-GRE habitats
Habitats included in EMAP-GRE are relevant to the goals of EMAP-GRE either because
they 1) are specified in the Clean Water Act (i.e., water quality in the river channel), 2) are well
documented to be essential for a reliable assessment of river ecosystem condition (e.g.,
vegetation in the riparian zones), or 3) reveal aspects of ecological condition not captured by
sampling other aquatic habitats (e.g., littoral benthos). Off-channel habitat types, including
backwaters, tributary mouths, and floodplain lakes are not sampled in EMAP-GRE.
Three basic habitats are included in EMAP-GRE:
1. Main channel. This habitat includes the fluvial channel containing the most discharge.
Navigation pools on the Upper Mississippi River an Ohio Rivers are included. Lake
Pepin on the Upper Mississippi River is included. Mainstem reservoirs on the Upper
Missouri River are excluded.
2. Main-channel littoral zone. This habitat includes the main-channel margin to a
maximum depth of 6 m or a maximum distance of 30 m from the wetted perimeter
(whichever is closer to the shoreline). The depth criteria is defined by the effective
sampling range of the electrofishing gear.
4. Main-channel riparian zone. This habitat includes terrestrial habitat in a 30-m wide
zone adjacent to the main channel.
1.3 EMAP-GRE indicators
Indicators are measurements that characterize an ecosystem or one of its critical
physical, biological, or chemical components. Indicators vary in their cost, variability, sensitivity
to different stressors, and societal value (Jackson et al. 2000). Presumptive performance
relative to these criteria provides the rationale for including each indicator in the EMAP-GRE
assessment.
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In April, 2003, an EMAP-GRE Indicator Workshop was held in Minneapolis, MN (Angradi
and Hill 2003) to identify elements of GREs for which indicators should be developed and to
identify appropriate measurements and methods for each indicator. Experts on indicators, the
ecology of large rivers, and the EMAP approach participated in the workshop. The advantages
and shortcomings of a large number of potential Great River indicators were considered. The
indicators and methods included in this manual reflect those and subsequent discussions. Due
to financial and other constraints, not all of the indicators identified in the workshop as
potentially useful for assessing GREs are currently included in EMAP-GRE.
EMAP-GRE includes four types of indicators: condition indicators, stressor indicators,
exposure indicators, and function indicators. Condition indicators are biotic or abiotic
characteristics of an ecosystem that reveal the condition of an ecosystem or habitat relative to
an environmental value or reference condition. Fish assemblage structure is an example of a
biotic condition indicator. Stressor indicators characterize or quantify anthropogenic effects on
the condition of ecosystems. River nutrient concentration is an example of a stressor indicator.
Exposure indicators quantify exposure of the biota at one or more trophic levels to toxic
contaminants. Fish tissue contamination is an example of an exposure indicator. Function
indicators measure the magnitude or rate of an ecosystem process relevant to ecosystem
condition. Sediment enzyme activity is an example of a function indicator.
Not all indicators will be sampled or measured in all habitats. In some cases, the
indicator is only relevant to or present in a single habitat (e.g., riparian vegetation structure). In
other cases, logistic considerations render collection of an indicator more feasible in some
habitats than in others (e.g., benthic macroinvertebrates in littoral areas versus the main
channel). Some indicators are presumed to reliably characterize GRE condition when
measured in a particular habitat and are not measured elsewhere to reduce the overall effort
required. For example, fish are sampled in the main-channel littoral zone of the main channel
rather than in the thalweg or in backwaters.
1.3.1 Water chemistry and plankton (Section 5)
In EMAP-GRE, water chemistry data are primarily used to define reference conditions
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and to identify stressor gradients. Great River stressors associated with water chemistry may
include nutrient enrichment, inorganic contamination, anoxia, temperature stress, turbidity, and
suspended sediment. Water chemistry sampling includes a grab sample of water for laboratory
analysis and in-situ measurements, including dissolved oxygen, conductivity, pH, turbidity, total
suspended solids (TSS), and seston geochemistry.
Plankton include algae (phytoplankton) and microinvertebrates (zooplankton)
suspended in the water column. Plankton assemblages are potentially useful indicators of
environmental condition because they are important to the trophic structure of larger rivers, and
they are likely sensitive to a number of anthropogenic disturbances, including flow regulation,
habitat alteration, invasive species, and contamination by nutrients, metals, and herbicides.
1.3.2 Aquatic vegetation (Section 6)
Aquatic vegetation has multiple ecological functions in Great River ecosystems. Aquatic
plant communities generate dissolved oxygen, stabilize bed sediments, filter suspended
sediment, and immobilize nutrients and toxic substances. Plant parts are an important food
source for waterfowl and other wildlife. Submerged, floating, and emergent aquatic plants
provide substrate for invertebrates and habitat for fish. Submerged aquatic vegetation (SAV) is
sensitive to anthropogenic stressors, including excessive turbidity, sedimentation, flow
modification, and exotic herbivores. Relating SAV community structure, abundance, and
distribution to stressors can provide a biological basis for water quality criteria. For example,
understanding the influence of turbidity on SAV beds could lead to development of light-related
water quality criteria for Great Rivers (UMRCC 2003).
1.3.3 Riparian habitat (Section 7)
Interactions among aquatic and riparian ecosystem components are important in Great
River ecosystem functioning and condition. Riparian ecosystems contribute to and moderate
the flux of materials and energy between terrestrial and aquatic habitats within GREs. Riparian
and shoreline habitat characteristics (vegetation, shoreline stability, human disturbance)
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influence affect channel form, water velocity, substrate and other physical habitat attributes at
multiple spatial scales. Riparian measurements are also important in EMAP-GRE because they
provide robust abiotic indicators of human disturbance at the site scale that are useful for
identifying reference sites.
1.3.4 Fish (Section 8)
EMAP-GRE fish sampling methods are designed to collect all but the rarest fish in near-
shore littoral habitats at each site. The sample collected is assumed to accurately represent the
proportional abundance of the targeted assemblage at the site. Benthic species that inhabit the
thalweg or other deep main channel habitats may not be adequately sampled by electrofishing.
Fish sample data include species composition, size distribution, and the occurrence of
anomalies on individual fish. Other measures of assemblage structure and function can be
calculated from the data and combined into indices of condition useful for assessing the
condition of Great Rivers (Emery et al. 2003, Simon and Emery 1995).
1.3.5 Fish tissue (Section 9)
Fish tissue contaminants are an indicator of bioaccumulation of persistent toxic
substances in the environment, and can be used to estimate exposure to contaminants
associated with fish consumption for higher trophic levels. EMAP-GRE will focus on whole fish
rather than on fillets because of its emphasis on the health of the ecosystem. Although whole-
fish contamination is primarily an indicator of contaminant exposure to piscivorous wildlife,
whole fish data are still relevant for estimating human exposure to contaminants through fish
consumption.
1.3.6 Benthic macroinvertebrates (Section 10)
Benthic macroinvertebrates inhabit river bed sediments or adhere to hard substrates.
Macroinvertebrates have several advantages as condition indicators (Barbour et al. 1999,
Klemm et al. 1990). Macroinvertebrates are ubiquitous throughout all GRE aquatic habitats and
8
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are relatively easy to collect in large numbers in most habitats. In some situations of multiple
stressors, macroinvertebrate assemblages have diagnostic power to identify stressors. Benthic
macroinvertebrates generally live one or two years, and they usually recolonize substrates
relatively rapidly after disturbance. Therefore, they are likely most useful for detecting stressors
at temporal scales of several months up a year or two. Mobility partly determines the spatial
scale at which organisms respond to stress. Macroinvertebrates are relatively sessile and
substrate specific, and are very sensitive to sedimentation, sediment contamination, and habitat
alterations (e.g., rip rap). EMAP-GRE includes two types of macroinvertebrate sampling:
shoreline (littoral) kick sampling, and snag surface sampling from a boat.
Two types of condition indicators can be developed based on macroinvertebrate
assemblages: multimetric additive indices of condition, and multivariate approaches which use
predictive modeling to assess condition. Multimetric indices combine various ecological
attributes of the benthic assemblage into an index that reflects the condition of the assemblage
and is compared to a reference value for assessment. Examples of the multimetric approach
include Kerans and Karr (1994), Barbour et al. (1996), and Klemm et al. (2003). In the
multivariate approach, abiotic data associated with each sample are used to predict which
organisms should be in the sample based on a model developed from reference sites.
Examples of this approach include Wright (1995) and Reynoldson et al. (1995).
1.3.7 Periphyton and sediment (Section 11)
Periphyton include algae, fungi, bacteria, protozoa, and associated organic matter on
the surface of aquatic substrata. Periphyton is a useful indicator of environmental condition
because periphyton is easy to collect and it responds rapidly to a number of anthropogenic
disturbances, including habitat alteration and contamination by nutrients, metals, herbicides,
hydrocarbons, and acids (Pan et al. 1996, Hill et al. 2003). Indicators based on periphyton may
be based on assemblage species composition, cell density, ash-free dry mass, chlorophyll
concentration, and enzyme activity. As for macroinvertebrates, a index can be developed which
combines multiple ecological characteristics of periphyton into an additive index useful for
bioassessment (Hill etal. 2003).
Benthic organisms are in intimate contact with river sediments and are influenced by the
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physical and chemical properties of sediment. Sediment characteristics serve as exposure
indicators for benthos, fish, and other wildlife (e.g., sediment toxicity) and as functional
indicators of key ecosystem processes (e.g., nutrient dynamics as revealed by sediment
enzyme activity) (Sinsabaugh and Foreman 2001, Hill et al. 2002).
1.4 Quality assurance/quality control
EMAP-GRE has a rigorous QA/QC program that includes all aspects of the project,
including field operations, lab analysis, and information management. Generic field-operations-
related QA/QC considerations are outlined in Table 1-1. More detailed QA/QC considerations
are included in each section of this manual.
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Table 1 -1. Generic QA activities for EMAP-GRE field operations.
Category
Standardization
Calibration
Considerations
Training All crews will be thoroughly and consistently trained for assigned field tasks,
safety, and project QA procedures. Initial training is supplemented by
annual "booster" training.
Crews will receive standardized training based on this manual.
Standardized field forms and labels will be used by all crews. Field
instruments, sampling equipment, and supplies will be specified or supplied.
Calibration of field instruments will be integrated with field operations.
Objectivity
Field operations are designed to minimize unnecessary subjectivity in
measurements. To the extent possible, rules will be provided for site
verification and other field decisions.
Communication Regular communication between field crews and coordinating EPA
personnel forestalls problems, misinterpretations, and supply shortages and
promotes inter-river and among-crew uniformity. Field-season QA audits
and post-field-season debriefings improve QA.
Documentation Non-standard or unusual situations or conditions are documented with data
quality flags and notes on the field forms and by communication with EPA
scientists and IM personnel.
Information Web-based sample tracking and 100% data proofing are fully integrated
management into the program
1.5 Safety
Safety is paramount. Operating on large rivers is inherently hazardous and involves
significant risks to crew health and safety. Obtaining data is always less important than
maintaining the safety of the crew. Specific safety practices and precautions related to field
procedures and logistics are integrated into this manual. These practices do not supercede or
replace the safety practices or guidelines of other agencies whose personnel are conducting
EMAP field work. Refer to Flotermersch et al. (2001) for additional guidance on operational
safety when working on rivers.
1.6 Training
Training sessions were conducted prior to the first EMAP-GRE field season. The
training included a full day of classroom instruction and a day of on-the-river demonstration of
field operations. At a minimum, all crew leaders receive the complete training. Crew members
11
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not present at the EPA training were trained in the field. Prior to subsequent field seasons, a
"booster" training is held which reinforces the initial training, trains new crews leaders, and
emphasizes any changes in procedures that may have occurred.
1.7 EPA support of field operations
EMAP-GRE field sampling will typically not be conducted by EPA personnel, but the
EPA principal investigators will be available during the field season for consultation with field
crews on sample design, logistics, methods, and other issues that arise during the field work.
Table 1-2 includes contact information for project principal investigators and the EMAP data
manager.
Table 1-2. Contact information for EPA-GRE principal investigators.
David Bolgrien
(218)529-5216
fax: (218) 529-5003
bolgrien.dave@epa.gov
Lead role: Project administration,
project public relations, site verification
Ted Angradi
(218) 529-5243 (office)
(720)480-7321 (mobile)
fax: (218) 529-5003
angradi.theodore@epa.gov
Lead role: Field Operations Manual, logistics
Marlys Cappaert
(541) 754-4467
fax: (541) 754-4338
cappaert.marlys@epa.gov
Lead role: EMAP data manager,
sample tracking
Terri Jicha
(218) 529-5153
fax: (218) 529-5003
jicha.terri@epa.gov
Lead role: Information management, water
sampling, sample tracking
1.8 Literature cited
Angradi, T.R., and B.H. Hill (editors). 2003. Environmental Indicators for Great River
Ecosystems. A Report from the EMAP-GRE Indicator Workshop, April 29-30, 2003,
Minneapolis, MN.
12
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Angradi, T.R., E.W. Schweiger, and D.W. Bolgrien. 2002. Upper Missouri River Pilot Project
Field Operations Manual for River Sampling. Environmental Monitoring and
Assessment Program. Mid-Continent Ecology Division, National Health and
Environmental Effects Research Laboratory, Office of Research and Development, U.S.
Environmental Protection Agency, Duluth, MN. 41p.
Baker, J.R., D.V. Peck, and D.W. Sutton (editors). 1997. Environmental Monitoring and
Assessment Program Surface Waters Field Operations Manual for Lakes. EPA/620/R-
97/001. U.S. Environmental Protection Agency, Washington, DC.
Barbour, M.T., J. Gerritsen, G.E. Griffith, R. Frydenborg, E. McCarron, J.S. White, and M.L.
Bastian. 1996. A framework for biological criteria for Florida streams using benthic
macroinvertebrates. Journal of the North American Benthological Society 19: 185-211.
Barbour, M.T., J. Gerritsen, B.D. Snyder, and J.B. Stribling. 1999. Rapid bioassessment
protocols for use in streams and wadeable rivers: periphyton, benthic
macroinvertebrates and fish, Second Edition. EPA/841/B/99/002. U.S. Environmental
Protection Agency; Office of Water: Washington, DC.
Http://epa.gov/OWOW/monitoring/techmon.html
Emery, E.B., T.P. Simon, F.H. McCormick, P.L. Angermeir, J.E. DeShon, C.O. Yoder, R.E.
Sanders, W.D. Pearson, G.D. Hickman, R.J. Reash, and J.A. Thomas. 2003.
Development of a multimetric index of assessing the biological condition of the Ohio
River. Transactions of the American Fisheries Society 132:791-808.
Flotermersch, J.F., S.M. Cormier, and B.C. Autrey. 2001. Logistics of ecological sampling on
large rivers. EPA/600/R-00/109. U. S. Environmental Protection Agency, Washington,
DC.
\\\\, B.H., AT. Herlihy, and P.R. Kaufmann. 2002. Dehydrogenase activity in sediments from
Appalachian Mountain, Piedmont and Coastal Plains streams of the eastern U.S.A.
Freshwater Biology 47:185-194.
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Hill, B.H., AT. Herlihy, P.R. Kaufmann, M.A. Vander Borgh, and S.J. DeCelles. 2003.
Assessment of streams of the eastern United States using a periphyton index of biotic
integrity. Ecological Indicators 2:325-338.
Jackson, L.E., J.C. Kurtz, and W.S. Fisher, 2000. Evaluation guidelines for ecological
indicators. EPA/620/R-99/005.U.S. Environmental Protection Agency, Washington, DC.
Kaufmann, P.R., P. Levine, E.G. Robinson, C. Seeliger, and D.V. Peck. 1999. Quantifying
physical habitat in wadeable streams. EPA/620/R-99/003. U.S Environmental Protection
Agency, Washington, DC.
Kerans B.L. and J.R. Karr. 1994. A benthic index of biotic integrity (B-IBI) for rivers of the
Tennessee Valley. Ecology 4: 768-785.
Klemm, D.J., P.A. Lewis, F. Fulk, and 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, Washington, DC.
Klemm D.J., K.A. Blocksonm, F.A. Fulk, AT. Herlihy, R.M. Hughes, P.R. Kaufmann, D.V. Peck,
J.L. Stoddard, W.T. Thoeny, M.B. Griffith, and W.S. Davis WS. 2003. Development and
evaluation of a macroinvertebrate biotic integrity index (MBII) for regionally assessing
Mid-Atlantic highlands streams. Environmental Management 31: 656-669.
Moulton, S.R.II, J.G. Kennon, R.M, Goldstein, and J.A. Hambrook. 2002. Revised protocols for
sampling algal, invertebrate and fish communities as part of the National Water-Quality
Assessment Program. Open-File Report 02-150, U.S. Geological Survey, Reston, VA.
Pan, Y., R.J. Stevenson, B.H. Hill, AT. Herlihy, and G.B. Collins. 1996. Using diatoms as
indicators of ecological conditions in lotic systems: A regional assessment. Journal of
the North American Benthological Society 15:481-495.
14
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Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and
Monaco, P. A. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Non-Wadeable
Rivers and Streams, U.S. Environmental Protection Agency, Washington, DC.
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Ringold, P. L, Magee, T. and
Cappaert, M. R. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams,
U.S. Environmental Protection Agency, Office of Research and Development,
Washington, DC.
Reynoldson, T.B., R.C. Bailey, K.E. Day, and R.H. Morris. 1995. Biological guidelines for
freshwater sediment based on benthic assessment of sediment using a multivariate
approach for predicting biological state. Australian Journal of Ecology 20:198-219
Simon, T.P. and E.B. Emery. 1995. Modification and assessment of an index of biotic integrity
to quantify water resource quality in Great Rivers. Regulated Rivers: Research and
Management 11:283-298.
Sinsabaugh, R.L. and C.M. Foreman. 2001. Activity profiles of bacterioplankton in a eutrophic
river. Freshwater Biology 46:1239-1249.
Strobel, C.J. and T. Heitmuller. 2001. National Coastal Assessment Field Operations Manual.
EPA/620/R-01/003. 71 p. U.S. Environmental Protection Agency, Washington, DC.
Upper Mississippi River Conservation Committee, Water Quality Technical Section (UMRCC).
2003. Proposed water quality criteria necessary to sustain submersed aquatic
vegetation in the Upper Mississippi River. Rock Island, IL. 6p.
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USEPA. 2002. Research Strategy of the Environmental Monitoring and Assessment Program.
U.S. Environmental Protection Agency, Washington, DC. EPA 620/R-02/002, July 2002.
Wright, J.F. 1995. Development and use of a system for predicting the macroinvertebrate fauna
in flowing waters. Australian Journal of Ecology 20:181-197.
Yin, Y., J.S. Winkelman, and H.A. Langrehr. 2000. Long-term resource monitoring program
procedures: aquatic vegetation monitoring. U.S. Geological Survey, Upper Midwest
Environmental Sciences Center, 2630 Fanta Reed Road, La Crosse, Wl 54603.
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Section 2
Overview of Field Operations
Ted Angradi1
This section describes the daily operational scenario for EMAP-GRE field activities.
Included are field-crew configuration and responsibilities, a discussion of boat operations, a
flow chart of daily operations, guidelines for recording data, and general safety considerations.
2.1 Crew configuration and responsibilities
EMAP-GRE field operations require at least two crews: a three-person fish-sampling
crew and a three-or four-person river-sampling crew. In some cases the same crew may
perform both functions. These crew sizes are only recommendations; alternative crew
configurations are acceptable. Crew responsibilities are outlined in Table 2-1. Many logistical
aspects of field operations such as site verification, lodging, transportation, and sample tracking
and shipping may overlap between crews. In the field, each crew is supervised by a crew
leader, who is responsible for daily operational planning, data quality, and safety.
Whenever possible, the crews should coordinate their activities. By visiting the same
site on the same day, crews can share equipment, coordinate laying out the reach, and provide
mutual support in case of a breakdown or safety emergency. River and fish-sampling at a site
should be completed within five days of each other whenever possible.
2.2 Boat operations
2.2.1 Operational logistics
Each crew requires a boat for sampling. Care must be taken to maintain the boats in
good order. Consult Flotermersch et al. (2001) for guidance on the logistics of boat operations
for river sampling.
1 U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
17
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2.2.2 Navigation
The boat trip from the ramp to the sample site may be many miles, and may involve
potential hazards. All boats should be equipped with a high-quality dash-mounted GPS/sonar
unit with preloaded basemaps. Site location (latitude, longitude) data from the design file
should also be loaded into the GPS units as waypoints. As part of pre-visit activities (described
in more detail in Sections 3 and 4), crews should plan their route to make sure they use the
closest suitable ramp, and that they are aware of any hazards, including locks, rapids, and
shoals.
Table 2-1 . Outline of the responsibilities of each field crew.
Crew Habitats Sampling responsibilities
Fish sampling Near-shore littoral Fish assemblages
Substratum
Fish cover
Fish tissue for contaminants
Fish tissue for DMA
River sampling Main channel Water chemistry
Phytoplankton
Macrozooplankton
Microzooplankton
LWD
Near-shore littoral LWD
Benthic macroinvertebrates
Snag macroinvertebrates
Sediment
Periphyton
Aquatic vegetation
Riparian zone Bank characteristics
Riparian vegetation structure
Human disturbance
Invasive plants
Section of manual
8
8
8
9
8
5
5
5
5
10
10
10
10
11
11
6
7
7
7
7
2.2.3 Boating safety
Boating on large rivers presents multiple safety hazards. The river must always be
treated with respect to avoid situations that threaten the health and safety of crews. Table 2-2
18
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lists safety recommendations related to general boat operations. Crews should also receive
safe-boating training. Other safety consideration related to sampling are described in
subsequent chapters.
2.3 Flow of daily operations
This section outlines a proposed general flow of daily operations for the fish and river
sampling crews (Figure 2-1). The two different crews may operate independently or together
depending on the schedule of operations. The details of each activity are provided in
subsequent chapters. At any particular site, circumstances may require that the order of the
activities be altered.
2.3.1 River-sampling crew
After completing base location activities (Section 3) and navigating to the sample site,
the crew leader evaluates whether the site is safe to sample under the existing conditions
(sampleability may be apparent at the boat ramp). See Section 4.6 for more guidance on
sampleability. If the site is safely sampleable, the river-sampling crew collects depth- and
width-integrated water and plankton samples and makes water quality measurements (Section
5). The crew then locates and flags two 500-m shoreline transects (if they have not already
been flagged by the fish-sampling crew; site layout is described in Section 4). Two crew
members collect a composite littoral macroinvertebrate sample (Section 10), a composite
sediment sample (Section 11), and a composite periphyton sample (Section 11) at each of 11
stations along the primary 500-m shoreline transect. While this is occurring, the other crew
member collects bank and riparian habitat data (Section 7) and aquatic vegetation data
(Section 6), Following this, the crew returns to the boat, collects a macroinvertebrate sample
from the surface of a snag in the channel, and collects LWD abundance data (Section 10).
Before departing the site, the river-sampling crew performs a subjective overall site assessment
(Section 7).
19
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crew
Location
activities
crew
Coiled littoral
benthos samples
(1 or 2 people)
Collect riparian
habitat data
(1 or 2 people)
location
activities
Figure 2-1. Flow chart of field activities for EMAP-GRE river- and fish-sampling crews
(proposed). Each crew consists of at least three people. Site verification and the
location and flagging of the MCS transects is done by the first crew to arrive at
the site. The river-sampling crew may split up for the littoral sampling. One or
two people can collect benthos, periphyton, and sediment samples, while the
other crew member(s) collects riparian habitat and aquatic vegetation data. Pre-
and post-visit activities are described in Section 3; site verification and reach
layout is described in Section 4; sampling is described in Sections 5-11.
2.3.2 Fish-sampling crew
After completing base location activities (Section 3) and navigating to the sample site,
the crew leader evaluates whether the site is sampleable under the existing conditions
(sampleability may be apparent at the boat ramp). The fish-sampling crew locates and flags two
500-m near-shore transects to be electrofished (if they have not already been flagged). The
20
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crew electro-fishes each 500-m transect (Section 8). Captured fish are identified, measured,
weighed, and released. A subsample of captured fish are retained as taxonomic vouchers and
for fish tissue analysis (Section 9). The crew then collects substrate-size and fish cover data
along each transect (Section 8).
2.3.2 River teams
The fish-sampling and river-sampling crew for each section of river together comprise a
river team. Each team is assigned a three letter code that is used in certain field forms to aid
with information management and sample tracking:
Team code
LCM
BVM
OLM
HSM
GRM
ORM
MRU
MRM
MRL
OHR
River
Mississippi
Mississippi
Mississippi
Mississippi
Mississippi
Mississippi
Missouri
Missouri
Missouri
Ohio River
Name (headquarters location)
Lake City (Lake City, MN)
Bellvue (Bellvue, IA)
Onalaska (La Crosse, Wl)
Havana (Havana, IL)
Great Rivers (Brighton, IL)
Open River (Jackson, MO)
Upper Missouri River (Bismarck, ND)
Middle Missouri River (Lincoln, NE)
Lower Missouri River in Missouri and Kansas (Rolla, MO)
Ohio River (Cincinnati, OH)
21
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Table 2-2. Safety considerations related to general boat operations.
All field personnel should receive U.S. Coast Guard-approved safe boating and water safety
training.
Be conservative navigating the river. Grounding on sandbars, snags, or wing dams is extremely
dangerous and damaging. Be aware that the safest and deepest approach to sites in
secondary channels, backwaters, or among sandbars is usually from downriver.
Silver carp (Hypophthalmichthys molitrix) >10 kg can leap >2 m out of the water. Several
people have been seriously injured in carp collisions. Silver carp are present in the lower
reaches of all three rivers. Be alert for leaping fish while running the river and during
electrofishing.
Trust the GPS unit for back-navigating shoals and at night. All crew mem bers should know how
to use the "man-overboard" feature on the GPS.
Secure and stow dangerous objects forward when running. They become unguided missiles
when the boat stops suddenly.
Always double check the anchor set when landing on shore. Be aware that river stage may
change dramatically in a short time and plan accordingly. Always anchor from the bow eye
when in any current.
Make sure the boat is equipped with enough suitable PFDs for everyone aboard, a throwable
device, a fire extinguisher, and other required safety items.
A large fluke anchor with a section of chain and a line buoy is recommend for anchoring in
current in sand-bottom reaches. At least a 7:1 ratio of anchor line length:depth is
recommended for anchoring in current.
Carry a cell phone, tools, first aid supplies, engine oil, sun block, and insect repellent on board.
Carry sufficient line to tow another boat back to the ramp.
Check the VHP weather channel frequently for storm alerts. Thunder or lightning means get off
the river.
Perform a regular inspection of boat tie-downs, trailer connections, winch parts and cable, tires,
bearings, and lights.
When it is safe to do so, render immediate assistance to other boats or boaters in distress.
Insure that a safe situation exists before returning to field work.
In clear water, polarized sunglasses greatly improve the ability of the boat driver to see
underwater hazards.
22
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2.4 Guidelines for recording data and information
Following the guidelines for filling out field forms and sample labels (Table 2-3) is
essential for quality assurance. Errors or sloppiness in recording data can result in data being
lost from the program. Forms are designed to be optically scanned into electronic files.
Photocopying alters the dimensions of forms so that they are not readable using the scanning
system. Therefore, only original forms and labels provided by EPA can be used. Originals
should be photocopied as soon as possible after field work.
23
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Table 2-3. Guidelines for recording field data and other information (adapted from
Peck et al., unpublished draft)
Recording data on field forms
Use forms preprinted on water-resistant paper supplied by EPA.
Header information on each field form links the data. Make sure headers are filled in completely on both
sides of each form.
Never mark on or around the corner blocks or ID box on field forms. These preprinted markings are
necessary for the optical scanning software; obscuring them will affect performance.
Write legibly using a soft pencil or pen so information can be read by the optical scanner. Erase
mistakes completely and write in the correct value whenever possible.
If a value must be lined out, write the correct value adjacent to the line-out so the data entry operator can
find it.
USE ALL CAPITALS WHEN ENTERING TEXT ON THE FIELD FORMS. Clearly distinguish letters from
number (e.g., 0 vs. O, 2 vs Z, 7 vs T). Do not put lines through 7's, O's, or Z's. Do not use slashes.
WRITE IN ALL CAPITALS IN THE COMMENTS SECTION. Be concise but avoid abbreviations or
shorthand notation. Attach additional sheets if necessary.
Always record the full AFS common name for fish.
Record data and information so that all the entries are obvious. Enter data completely in every field that
is used. Follow the "comb" guidelines - print each number or letter in the individual space provided.
Keep letters and numerals from overlapping.
When values need to be circled, use these proportions: QJ
Record data to the number of decimal places provided on the forms.
If the measurement is zero, enter a zero. Blanks will be interpreted as missing data (which should be
flagged).
If the field calls for meters, enter the value in meters. Do not use different units and a notation to that
effect. If the number is negative, enter the value and flag it as a negative value in the comments.
Do not enter longitudes as negative values (as in the dossier). Use 2 spaces left of the decimal place for
longitudes <100 degrees west; use 3 spaces left of the decimal place for longitudes >100 degrees west.
Record information on each line even if it has to be recorded repeatedly (e.g., fish species names). Do
not use a vertical line to indicate repeated entries.
Make a copy of completed forms and return the original field forms to the data manager (Corvallis).
Retain the copy in a master file for the site. Keep forms in order (they are numbered) and do not staple
them together. Three-ring binders are a good option for the form copies.
Continued
24
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Table 2-3. Guidelines for recording field data and other information, continued.
Data Flags
Use only defined flag codes from the list below and record them on the data form in the
appropriate field. If data are collected for which there is no space on the form, choose a flag
and record the data in the comment section.
Flag Comment
F1, F2, Fn Miscellaneous comments assigned by field crew; use only once per form
K Sample not collected or lost; no measurement made
U Suspect sample, measurement, or observation; sample collected using a
nonstandard procedure
Q Unacceptable QC check associated with measurement
Sample Labels and Tracking
Use adhesive labels with preprinted sample ID numbers for each type of sample.
Record information on labels using a fine-point permanent marker (e.g., Sharpie). Cover labels
with clear tape.
Record sample ID numbers from the label on field forms and on sample tracking forms.
Reconcile sample ID numbers on samples and tracking forms before shipping samples.
Include a copy of the sample tracking form with each sample shipment (the original is part of
data form packet for the site that is mailed to the information manager).
25
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2.5 Collecting permits
All states require collecting permits for fish sampling. Federal permits may also be
required. Some states require permits for collecting plants or macroinvertebrates. Obtaining
collecting permits and filing collecting reports is the responsibility of the field crews.
Copies of the permits should be carried on boats when sampling. Crews should closely follow
the specifications of the permit(s). These specifications may include destruction of certain
species if captured (e.g., Asian carp), notification of the permitting agency prior to field
sampling, and submission of an annual report listing the fish collected and their disposition.
Consult Walsh and Meador (1998) for a summary of state permitting agencies and their
reporting requirements.
2.6 General safety considerations for field operations
Field work on Great Rivers is inherently hazardous and involves significant risks to crew
safety and health. Safety considerations for boat operations are given in Table 2-2. Additional
general safety considerations are presented in Table 2-4. Safety considerations associated
with specific sampling activities and gear are presented in subsequent chapters. Additional
resources include the American Red Cross and Handal (1992), Ohio EPA (1990), USCG
(1987), and USEPA (1986). Web sites with useful safety information include
www.cdc.gov/niosh (occupational safety), www.nws.noaa.gov/safety (weather safety),
www.uscgboating.org (boating safety), and www.firstaidguide.net (includes insect bite
information).
Personnel on EMAP-GRE field crews should be in sound physical condition, be able to
swim, and have a physical exam annually or in accordance with their agency policy. Crew
members with "MedicAlert" health conditions (e.g., severe allergies, diabetes,
susceptibility to seizures) should make crew leaders and other crew members aware of
their condition, the symptoms, and the actions required in a health emergency.
Water and sediment samples handled in the field should be considered potential health
hazards due to toxic substances or pathogens. Personnel must be familiar with health hazards
associated with using chemical fixing or preserving agents. Material Safety Data Sheets for all
chemicals used in field operations must be available to personnel. Chemical wastes can cause
26
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various hazards due to flammability, explosiveness, toxicity, causticity, or chemical reactivity. All
chemical wastes must be discarded according to standardized health and hazards procedures
(NIOSH 1981, USEPA 1986).
During the course of field activities, crews may observe apparent violations of
environmental regulations, may discover improperly disposed hazardous materials, or may
observe or cause an accidental spill or release of hazardous materials. In such cases, it is
important that the proper actions be taken and that field personnel do not become exposed to
harmful substances. The following guidelines apply (Peck et al., unpublished drafts):
First and foremost during any environmental incident, it is extremely important to
protect the health and safety of all personnel. Take any necessary steps to avoid
injury or exposure to hazardous materials. If you have been trained to take action
such as cleaning up a minor field spill during fueling of a boat, do so. However,
always err on the side of personal safety.
• Field personnel should never disturb or retrieve improperly disposed hazardous
materials from the field and bring them back to the facility for disposal. This
action may worsen the impact to the area of the incident, incur liability, cause
personal injury, and waste time and money. However, field personnel should not
ignore environmental incidents. There is a requirement to notify the proper
authorities of any incident of this type.
• For most environmental incidents, the following emergency telephone numbers
should be provided to field crews as part of a Health and Safety Plan: state and
tribal Departments of Environmental Quality or Protection, United States Coast
Guard and the United States Environmental Protection Agency Regional Office.
In the event of a major environmental incident, the National Response
Center and Terrorist Hotline should be contacted at 1-800-424-8802.
2.7 Equipment and Supplies for EMAP-GRE sampling
Table 2-5 is a checklist of the generic equipment and supplies needed for all for EMAP-
GRE field operations. Each section of the manual also has a checklist of specific equipment
and supplies needed for sampling.
27
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Table 2-4. General safety guidelines for field operations.
Crews should receive adequate training including first aid, CPR, vehicle safety, boating and
water safety, electrofishing safety, and laboratory safety.
Crews should carry cell phones to maintain reliable communications, and should carry contact
information for local police, ambulance, fire and rescue departments, and should program
important numbers into the cell phones.
All crew members should know the location of the truck keys.
Serious health problems may be associated with working in polluted waters. Exposure to river
water and sediments should be minimized, especially near effluent discharge points. Use
gloves if necessary.
All electrical equipment must bear the approval seal of Underwriters Laboratories and must be
properly grounded.
Use appropriate protective equipment (e.g., gloves, safety eyewear) when handling and using
hazardous chemicals.
Be aware of risks posed by and first aid for poisonous snake bites, bee stings, ticks, and
poisonous plants. Plan for potential allergic reactions.
Be aware of hypothermia and heat exhaustion risks, symptoms, and first aid.
Use extreme care walking on ramps and shorelines, especially on riprap.
• All crew members should be aware of any "MedicAlert" conditions among crew members.
28
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Table 2-5. Generic equipment and supply checklist for all EMAP-GRE field operations.
Basic boat safety equipment not listed. Quantities are per boat.
Qty
1
1
1
6
6
2
1 box
1
1
1
1 copy
1 set
1
1
1 or 2
1
Item
River crew. Boat with a winch and GPS/sonar unit loaded with appropriate
base maps
Fish crew. Boat wired for electrofishinq with GPS/sonar unit loaded with
appropriate base maps
Hand-held WAAS-enabled GPS unit with extra batteries
Soft pencils for filling in field forms
Fine- and medium-point permanent markers for labeling
Form-holder clip-boards
Clear tape strips for covering labels
Multi-tool
Scissors for cutting labels
First-aid kit
EMAP-GRE Field Operations Manual
State and federal collecting permits
Health and Safety Plan with emergency contact information
Cell phone
Coolers with ice in sealed bags
Portable freezer (optional)
2.8 Literature cited
American Red Cross and K. A. Handal. 1992. The American Red Cross First Aid and Safety
Handbook. Little Brown and Company, Boston. 321 p.
NIOSH. 1981. Occupational Health Guidelines for Chemical Hazards (2 Volumes). National
Institute for Occupational Safety and Health, NIOSH/OSHA Publication No. 81-123. U.S.
Government Printing Office, Washington, DC.
29
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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.
Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and
Monaco, P. A. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Non-Wadeable
Rivers and Streams, U.S. Environmental Protection Agency, Washington, DC.
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Ringold, P. L, Magee, T. and
Cappaert, M. R. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams,
U.S. Environmental Protection Agency, Office of Research and Development,
Washington, DC.
U.S. Coast Guard. 1987. Federal Requirements of Recreation 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.
Walsh, S.J. and M.R. Meador. 1998. Guidelines for quality assurance and quality control offish
taxonomic data collected as part of the National Water-Quality Assessment Program: U.
S. Geological Survey Water-Resources Investigations Report 98-4239, 33p.
Http://water.usgs.gov/nawqa/protocols/WRI98-4239/index.html
30
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Section 3
Base-Location Activities
Ted R. Angradi1 and Terri M. Jicha1
Field crews conduct a number of activities at "base locations" before and after visiting
each river site. Base locations will usually be either the crews' temporary lodging facility or a
state or federal facility. Base-location activities are usually conducted on the same day as the
sampling visit.
3.1 Pre-visit base-location activities
Pre-visit activities include confirming suitability and location of launching facilities,
inspecting equipment, calibrating instruments, and assembling supplies and sample containers.
Procedures and guidelines for these activities are described in this section.
Pre-visit
QBIV feeders
« sie
* site
» GPS
*
*
pnwtous sles
•
•
*
Wwr ing visifJJ>
•
*
* fonts
• Fai to
• pick up
dews
*
•deon and
»
*
»
Figure 3-1. EMAP-GRE base-location activities. Fish- and river-sampling crew activities
combined.
1 U.S. States Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
31
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3.1.1 Confirming site location and status
The crew leaders will be provided with a "site dossier" containing location and access
information for each site. The crew leaders should confirm that the nearest launch facilities are
adequate and determine if there are any hazards or other special circumstances for the site.
Additional reconnaissance of the site may be necessary. The site dossier and site verification
are described in detail in Section 4.
3.1.2 Water quality meters
Five water quality parameters must be measured as part of EMAP-GRE field sampling:
temperature, dissolved oxygen (DO), conductivity, pH, and turbidity (turbidity may be measured
at the base location). No preferred manufacturer or model of meter is specified. Any reliable
meter that can be properly calibrated and adapted to the depth-integrated sampling protocol
(Section 5) is acceptable. A combination of meters (e.g., a DO/conductivity/temperature meter
and a pH meter) or a single multi-parameter sensor may be used. Specific calibration and
maintenance procedures and instructions for preparing calibration standards are not included in
this manual and are the responsibility of the field crew.
3.1.3 Meter calibration
At the beginning of the sample season, the accuracy of the DO meter should be tested
using a modified Winkler titration kit. Before each calibration attempt, inspect the membranes. If
bubbles are present, or if the membrane is discolored or torn, replace the membrane according
to manufacturer instructions. The DO meter should be calibrated daily either at the base
location or at the sample site. Record all calibration details in a calibration log book including
(as applicable) meter model and serial number, date, calibration standards used, elevation, and
meter maintenance performed. Each entry in the log should be signed. The calibration log will
be inspected during field QA audits and will become part of the EMAP-GRE data record.
Conductivity meters should be re-calibrated at least weekly. Follow instructions in the
instrument's operations manual and use a conductivity standard that is appropriate for the
expected range. Calibration with a 1000 uS/cm standard is usually satisfactory. Record
32
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calibration results in the calibration log book.
pH meters should be calibrated at least weekly at the base location using two standards,
pH 7 and pH 10. Record calibration results in the calibration log book.
Turbidity meters are usually factory calibrated for a wide range of turbidity values, but
should be calibrated at the beginning of the field season and about every two or three weeks
thereafter using a standard in the range of expected values. Record calibration results in the log
book.
Calibration of global positioning system (GPS) receivers should be done according to
the manufacturers specifications for initialization, after replacing batteries, or if a new reference
point is needed. Whenever possible, older units should be upgraded to modern units with built
in or up-loadable base maps. Crews that use multiple GPS receivers should check them
against each other and replace units that produce outlier coordinates.
3.1.4 Preparation of equipment and supplies
To ensure that all activities at a site can be conducted efficiently, field crews should
check all equipment and supplies before leaving the base location. Sample containers and
labels should be prepared ahead of time to the extent possible. Crews should inventory
equipment and supplies prior to departure using the checklists appropriate for their
responsibilities. Meters, probes, cameras, rangefinders, and other sensitive gear should be
packed to avoid shock, exposure, and other damage.
Prepare stock preservative solutions as described in Table 3.1. Regulations pertaining
to formalin and ethanol are in the Code of Federal Regulations. These requirements should be
summarized for all hazardous material being used for the project and an MSDS file should be
available to field personnel. Transport and store formalin and ethanol solutions in clearly-
labeled, non-breakable containers within secondary containment and outside vehicle cabs or
other unventilated spaces.
33
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Refuel vehicles and conduct maintenance and repairs the night before sampling, if
possible. Inspect vehicles every morning before departure. Check lights, boat tie downs, trailer
connections, and tire pressure. Make sure the spare tire for the trailer is in good condition.
Grease trailer hubs and jet drives (if applicable) frequently.
34
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Table 3-1. Stock preservative solutions and instructions for their preparation. All
stock solutions should be stored in clearly-labeled non-breakable containers.
Labels should include container contents, date of preparation, and the initials of
the preparer.
Solution
100% borax-buffered
formalin3
(pH 7-8)
100% carbonate-buffered
formalin (pH 10)b
12% buffered formalin-
sugar solution0
(pH 7-8)
10% borax-buffered
formalin
95% benzene-free
ethanol
85% benzene-free
ethanol
75% benzene-free
ethanol
10% carbonate-buffered
formalin (pH10)
Concentrated rose
bengal solution
1% bleach solution'
(optional)
Clorox Formula 409®
degreaser9
(optional)
Use
Preservative for
phytoplankton and
periphyton; stock
solution for fish
Stock solution for
macroinvertebrate
preservative
Preservative for
zooplankton
Preservative for fishd
Stock solution for fish
preservation (2005
DMA sites)8
Field preservation of
fish (2005 DMA sites)
Lab preservation of
fish (2005 DMA sites)
Preservative for
macroinvertebrates
Stain added to
macroinvertebrate
samples
Used to
decontaminate field
gear
Used to
decontaminate field
gear
Recipe
Add 20 g borax (hydrated sodium borate:
Na2B4O7-10H2O) detergent (20 Mule Team®) per
L 100% formalin (37% formaldehyde). Test pH
with paper.
Add 35 g Na2CO3 (also called "washing soda")
per L 100% formalin (37% formaldehyde). Test
pH with paper.
Add 600 ml_ 100% formalin, 5 tablespoons borax
and 400 g table sugar (sucrose) to 4.4 L tap
water (makes 5 L). Test pH with paper.
Add 1 part 100% borax-buffered formalin to 9
parts tap water.
Full strength agriculture-derived.
Add 9 parts 95% ethanol to 1 part tap water.
Add 8 parts 95% ethanol to 2 parts tap water.
Add 1 part 100% carbonate buffered formalin to
9 parts tap water. Test pH with paper.
Add 1 teaspoon rose bengal powder to 1 L of
10% carbonate-buffered formalin stock solution.
Add 1 part bleach to 99 parts water in a plastic
spray bottle.
Full strength or 50% dilution.
Formalin is a potential human carcinogen. Formalin should be handled only in well-ventilated areas while
wearing chemical-resistant gloves and approved eye protection.
High pH solution required to preserve mollusk shells (Merritt et al. 1997).
Modified from Haney and Hall (1975); this solution should be kept on ice in the boat.
Unbuffered 10% formalin may be used to preserve fish.
Ethanol allows preserved fish to be sampled for DMA analysis.
From Moulton et al. (2002); solution used to reduce risk of translocation of living organisms.
Kills operculate snails (e.g., New Zealand mud snails).
35
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3.2 Post-visit base-location activities
Upon reaching the base location or mobile laboratory after sampling, subsamples of
river water are filtered for chlorophyll a, total suspended solids (TSS), and geochemical
markers, and turbidity is determined (described in Section 5). The crew leader reviews data
forms and sample labels for accuracy, completeness, and legibility; attempts to fill in (and flag -
see Section 2.4) missing information as accurately as possible; and initials the data forms. Data
files from digital cameras should be recorded, downloaded, and backed up as soon as possible.
The other crew members should inspect and clean sampling equipment and boats as needed,
check the inventory of supplies, preserve and store samples, and prepare unpreserved samples
for shipment the next morning. Equipment maintenance tasks are listed in Table 3-2.
Invasive plants, fish, and invertebrates are potentially present at every EMAP-GRE
sample site. Drain bilges and live wells on site and inspect all sampling equipment, including
nets and boats, and remove any plant or animal material to prevent transporting nuisance
species between sites. Decontaminate gear (Table 3-2) when there is a risk of organism
translocation.
Crews should use discretion and common courtesy when using public facilities like river access
parking areas for sample processing. Do not litter or dispose of sediment, fish, or preservative
inappropriately. The curious public should be treated with respect and patience.
3.2.1 Sample packing, shipping, and tracking
An important aspect of program QA is sample handling between the field and the lab.
Inconsistent practices compromise QAand complicate information management. Crews should
follow the packing and shipping guidelines as closely as possible (Table 3-3 to 3-5) and should
ship fresh, unpreserved samples (water, sediment, aquatic plant voucher specimens) as soon
as possible after collection. Sample types, sample codes, shipping destinations, and which
tracking form to use for each sample type are summarized in Table 3-6.
Samples that must be frozen (chlorophyll filters and fish tissue) may be shipped in
36
-------
weekly batches but should be shipped as soon as possible if a freezer is not available. Samples
preserved by drying (TSS filters, geochemical markers) can be shipped as a batch at the end of
the field season. Formalin- or ethanol-preserved samples can be shipped in batches at the end
of the season (4% formalin) or retained for later transport to the laboratory (10% formalin, 75%
ethanol).
All samples, whether they are to be shipped immediately or retained and shipped later
are recorded on laboratory-specific sample tracking forms (Figure 3-2 to 3-9) which are copied
and included with each shipment. These forms also serve as a chain-of-custody between field
crews and the laboratory contact (the sample tracking responsibilities of the laboratory contact
are described in Section 3.3). Original sample tracking forms are faxed to the EMAP data
manager in Corvallis (541-754-4637) and then are added to the form package for the site which
will eventually be shipped to the data manager. Customer copies of courier airbills should be
saved in case of loss by the courier. Sample tracking is described in more detail in Table 3-7.
3.2.2 Samples not preserved with formalin
Samples not preserved with formalin include water, chlorophyll, TSS and geochemistry
filters, sediment, fish tissue samples for DMA analysis, and aquatic plant voucher specimens.
Each sample should be entered as a separate line on the appropriate sample tracking form.
Guidelines for packing and shipping these samples are given in Table 3-3. Use ice
substitute packs whenever possible to avoid leakage due to melting ice. When shipping with
ice, use block ice when available. Ice and ice substitute packs should be sealed in plastic bags
labeled "ice." Seal cooler lids with tape to prevent leakage in transit. A completed return airbill
(including billing account information) should be enclosed in each cooler so coolers do not
accumulate at laboratories.
Ship water chemistry and sediment samples as soon as possible after collection to
meet holding time requirements. Water and sediment samples that cannot be shipped
immediately should be refrigerated at 4°C. Samples collected on Friday and held over the
weekend should be shipped "same-day delivery" on Monday. Follow cooler packing instructions
37
-------
in Table 3-3. Be sure that at least half the final weight of the packed cooler is ice. Frozen
samples (fish tissue, chlorophyll filters) can be shipped in small batches comprising no more
than a week's worth of samples. However, if a freezer is not available, ship the chlorophyll and
fish tissue samples as soon as possible. Be sure labels are protected and the samples are well
sealed. Double-bag ice so that meltwater does not contaminate samples.
Table 3-2. Equipment care after each site visit.
Prior to departing the river, drain all water from bilges and live wells.
At the ramp or at the base location, inspect boats, trailers, and other gear for plant fragments or
animal remains and remove them. If appropriate, decontaminate gear with a 1 % bleach
solution to reduce risk of organism translocation.
Rinse water quality apparatus several times with distilled water after use.
Rinse the periphyton sampling equipment with tap water after use.
Rinse conductivity probes with deionized water and store moist.
Drain water sampling hoses.
Check fish dip nets, sieves, plankton, and benthos nets for holes and repair or replace.
Store nets dry to prevent mildew.
Inventory equipment and supplies and relay needs to field coordinator.
Examine DO meter membrane for cracks, wrinkles, or bubbles; replace if necessary.
Charge or replace all batteries as needed.
• Refuel boats, trucks, and generators.
38
-------
3.2.3 Samples preserved with formalin or ethanol
Formalin or ethanol-preserved samples include macroinvertebrates and fish specimens
preserved in 10% borax- or carbonate-buffered formalin, periphyton and plankton samples
preserved in 4% borax-buffered formalin, and 2005 fish specimens for DMA analysis preserved
in 85% or 75% ethanol. Samples preserved in ethanol or 10% formalin must be stored and
transported in secondary containers (tubs or coolers) and stabilized to prevent spillage.
For each sample, enter the sample ID from the label, the sample type, number of
containers, and any comments on the appropriate sample tracking form. For forms that include
only retained samples, leave the date sent and airbill number fields blank and note in the
comments which samples are being retained. When these samples are to be shipped or
transported to a lab, fill in the missing information and re-fax the completed forms to Corvallis.
Samples preserved in ethanol or 10% formalin should be retained by the crews until they can
be conveniently transported to the appropriate laboratory. Leave copies of the completed
sample tracking forms with the samples where they are being stored prior to delivery to the lab.
Tables 3-4 and 3-5 provide guidance for formalin- and ethanol-preserved samples that need to
be shipped by commercial carrier.
3.2.4 Inter-laboratory sample shipments
Some sample types are created in the laboratory by subsampling the original sample
(Table 3-6). There are three types of inter-laboratory samples: unpreserved sediment samples
shipped from NERL to MED, and preserved water samples for TOC and dissolved metals
shipped from UMESC to MED. All the sample packing, shipping, and tracking guidelines for
samples shipped from the field apply to inter-lab shipments. The sample tracking form and label
for inter-laboratory shipments is shown in Figures 3-2 and 3-3.
3.3 Sample tracking responsibilities of laboratory contacts
Sample tracking forms faxed from the field to the EMAP data manager will be forwarded
via email to the appropriate laboratory contact in the form of *.tiff image files. This email should
39
-------
arrive in advance of the shipped samples and serves to notify the laboratory contact that the
samples are in route or are being retained by the crew for later shipment or hand delivery.
When the samples arrive at the laboratory, the laboratory contact should check the contents of
the shipment against the tracking forms included with the shipment and forwarded by the data
manager, and assign condition codes to each sample. If there are problems with the shipment
(e.g., the sample is warm or mislabeled), the laboratory contact should communicate with the
field crew to prevent future problems.
As soon as possible after the samples are checked in, the laboratory contact must enter
the sample information into the EMAP Surface Water Information Management (SWIM) sample
tracking web page at https://emapsw.cor.epa.gov/tracking/labsamples.php3. When sample
information is entered in SWIM, the EMAP data manager is informed that samples have been
received at the laboratory. Inter-laboratory samples are treated the same as samples shipped
from the field. To access the SWIM sample tracking web page, the IP address from the lab's
computer(s) must be supplied to the EMAP data manager (Section 3.4). A username and
password will be provided to each laboratory contact.
3.4 Information management
A copy should be made of each set of completed and reviewed field forms. The
originals are arranged in numeric order (do not staple forms together) and mailed or shipped to
the EMAP data manager in Corvallis approximately monthly. The copies should be retained by
the field crew. Crews should maintain a master file for each site including the site dossier
(described in Section 4), copies of all field forms, sample tracking forms, air bills, data disks,
and any other documents that pertain to the site. Mail forms to:
Marlys Cappaert
EMAP Data Manager
Computer Sciences Corporation, c/o U.S. EPA, NHEERL/WED
200 S.W. 35th Street
Corvallis, OR 97333
(541)754-4467
40
-------
3.5 Equipment and supplies
A checklist of equipment and supplies required for base-location activities is presented
in 3-8. Generic supplies required for all EMAP-GRE field sampling are listed in Table 2-5.
41
-------
Table 3-3. General guidelines for packing and shipping unpreserved samples
(adapted from Peck et al., unpublished drafts).
Sample type
(container)
Guidelines
Samples requiring refrigeration (4° C)
Water chemistry
(4-L cubitainer and
500-mL bottle)
Sediment
(PE bag)
Plant voucher
specimens
(plastic bag)
Ship on day of collection or within 24 h by overnight courier.
Use fresh block ice in labeled ("ice") plastic bags.
Final weight of cooler should be at least half ice.
Line coolerwith a plastic bag.
Cover sample labels with clear tape to prevent label loss.
Confirm that sample ID data on the sample tracking form and field
forms agree.
Enclose a copy of the completed sample tracking forms and a pre-
paid return airbill in each cooler in a self-sealing plastic bag.
Minimize the volume of air space in the packed cooler.
Samples requiring freezing (-20° C) within 24 h of collection
Fish tissue
(plastic bags)
Chlorophyll
(filter holders)
If samples cannot be kept frozen in field, ship on day of collection or
within 24 h by overnight courier.
Frozen fish tissue samples should be shipped as soon as they are
frozen solid.
Frozen chlorophyll samples should be shipped in weekly batches.
Use fresh block ice in labeled ("ice") plastic bags.
Final weight of cooler should be at least half ice.
Cover sample labels with clear tape to prevent label loss.
Package samples so that meltwater does not contaminate them.
Confirm that sample ID data on the sample tracking form and field
forms agree.
Enclose a copy of the sample tracking form(s) in a self-sealing
plastic bag.
Minimize the volume of air space in the packed cooler.
Samples requiring drying (30 -50° C) for 24 h
Geochemical markers
(filter holder)
TSS
(filter holder)
If no oven is available, place in an air-conditioned room for 24-48 h
with lids ajar; cover with a sheet of foil to exclude dust.
Cover sample labels with clear tape to prevent label loss.
Confirm that sample ID data on the sample tracking form and field
form agree.
Enclose a copy of the sample tracking form in a self-sealing plastic
bag.
Dried filters should be shipped as a batch at the end of the season in
the tray boxes they came in (if available).
42
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Table 3-4. General guidelines for storing and shipping formalin-preserved samples
(adapted from Peck et al., unpublished drafts). In most cases, formalin-
preserved samples will not be shipped, but will be carried by truck to the
laboratory.
Sample type
(container)
Periphyton
(500 ml_ bottle)3
Macrozooplankton
(250 ml_ bottle)3
Microzooplankton
(250 ml_ bottle)3
Phytoplankton
(2 L bottle)3
Fish specimens
(jars of various sizes)
Macroinvertebrate
samples
(250-mL and 500-mL jars)
Final preservative
strength of
sample
4% borax-
buffered formalin
10% borax-
buffered formalin
10% carbonate-
buffered formalin
Guidelines
Labels or tags placed inside sample jars
must be of water-resistant paper.
The adhesive label on the outside of the
container should be completely covered with
clear tape.
Confirm that sample ID data on the sample
tracking forms and field forms agree.
Store samples in secondary containers at
base location.
Enclose copies of the sample tracking forms
in a self-sealing plastic bag with stored
samples.
Packaging and Shipping Guidance for samples preserved in formalin (IATA instructions 914, no limit)
Inside packaging
Outside packaging
Absorbent material
Screw top plastic bucket (20 L)with ratcheted lid is recommended (UN
spec 1 H2). Line container with plastic bag meeting IPS specifications.
Not required. Stabilize contents with packing peanuts.
Labeling
HOPE bottles with leakproof screw-top cap that meets UN spec IP2. Seal
cap with a strip of plastic tape. Fill jar to shoulder to provide headspace.
Outside package marked with UN shipping name and ID no.:
"Environmentally hazardous substance, liquid, n.o.s. (Formalin < 5%),
UN3082", and a "Class 9 Miscellaneous" label, and at least two package
orientation labels should be affixed to the container.
Shipping forms
Include packing list with each container. Note total quantity of formalin in
liters and the gross container weight in pounds. Prepare Shipper Manifest
prior to shipment.
Preserved periphyton and plankton samples may be shipped as unpreserved samples because formalin
concentration is low (<5%).
43
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Table 3-5. General guidelines for storing and shipping ethanol-preserved samples
(adapted from Peck et al., unpublished drafts). In most cases, ethanol-preserved
samples will not be shipped, but will be carried by truck to the laboratory.
Sample type
(container)
Final preservative
strength of
sample
Guidelines
2005 fish tissue DMA
specimens
(jars of various sizes)
75-85% ethanol
Labels or tags placed inside sample jars
must be of water-resistant paper.
The adhesive label on the outside of the
container should be completely covered with
clear tape.
Confirm that sample ID data on the sample
tracking forms and field forms agree.
Store samples in secondary containers at
base location.
Enclose copies of the sample tracking forms
in a self-sealing plastic bag with stored
samples.
Packaging and Shipping Guidance for samples preserved in formalin (IATA instructions 307, 60-L limit)
Inside packaging
HOPE bottles with leakproof screw-top cap that meets UN spec IP2. Seal
cap with a strip of plastic tape. Fill jar to shoulder to provide head space.
Outside packaging
Screw top plastic bucket (20 L) with ratcheted lid is recommended (UN
spec 1H2). Line container with plastic bag meeting IPS specifications.
Each pail can hold no more than 5.0 L total liquid.
Absorbent material
Sufficient volume of absorbent material (vermiculite, UN A100) Absorbent
sheets or equivalent) to absorb contents of all inner packaging. Stabilize
contents with packing peanuts.
Labeling
Outside package marked with UN shipping name and ID no.: "Alcohol,
flammable, toxic, n.o.s. (Denatured alcohol), UN1986, and a "Class 3
flammable" label, and a "Class 6 Toxic" label, and at least two package
orientation labels should be affixed to the container.
Shipping forms
Include packing list with each container. Note total quantity of formalin in
liters and the gross container weight in pounds. Prepare Shipper Manifest
prior to shipment. There is a 60-L limit for a single shipment.
44
-------
Table 3-6. Summary of sample type, sample codes amd shipping destination. Each sample destination (lab) has a unique
sample tracking form (Figures 3-2 to 3-9). Preserved samples are usually retained and shipped in batches or
transported to the lab at the end of the season. Shipped samples are not preserved and are shipped as soon as
possible after collection. Fish voucher specimens are retained by the crew for later laboratory identification.
Sample type
Phytoplan kton composite
Benthos kick sam pie composite
Benthos snag sample
Periphyton composite
Sedim ent chemistry su bsam pie
Total organic carbon
Dissolved metals
Small fish tissue
Large fish tissue
Sedim ent grab sam pie
Fin tissue DMA
W ater ch em istry com pos ite
Alkalinity grab
Aq uatic p lant vo uch ers
Geochemical markers filter
TSS filter pair 1
TSS filter pair 2
Chlorophyll filter
Macrozooplankton (63-pm mesh)
Microzooplankton (20-pm mesh)
Macrozooplankton (63-pm mesh)
Microzooplankton (20-pm mesh)
Macrozooplankton (63-pm mesh)
Microzooplankton (20-pm mesh)
Composite fish vouchers (field)
Fish species vouchers
Type code
pp
BK
BS
PA
SC
TC
DM
ST
LT
SG
DS
we
AL
AP
GF
SS1
SS2
CF
BZ
LZ
BZ
LZ
BZ
LZ
CV
SV
Shipped immediately or preserved
Retained
Retained
Retained
Retained
Shipped
Shipped
Shipped
Shipped
Shipped
Shipped
Shipped
Shipped
Shipped
Shipped
Retained
Retained
Retained
Retained
Retained
Retained
Retained
Retained
Retained
Retained
Retained
Retained
Sample destination
MED
MED
MED
MED
MED
MED
MED
NERL
NERL
NERL
NERL
UMESC
UMESC
UMESC
Stroud
Stroud
Stroud
Un iv Louisville
Un iv Louisville
Un iv Louisville
INHS
INHS
SMSU
SMSU
MED
MED
Comments
Carried to lab or shipped at end of season.
Carried to lab.
Carried to lab.
Carried to lab or shipped at end of season.
Inter-lab sample. Shipped fresh from UMESC to MED.
Inter-lab sample. Shipped in weekly batches from UMESC to MED.
Inter-lab sample. Shipped in weekly batches from NERL to MED.
Shipped frozen. May be shipped in weekly batches.
Shipped frozen. May be shipped in weekly batches.
Shipped fresh.
Shipped frozen. Shipped weeklywith ST and LT samples
Shipped fresh.
Shipped fresh.
Shipped fresh.
Shipped dried. Ship at end of season.
Shipped dried. Ship at end of season.
Shipped dried. Ship at end of season.
Shipped frozen weekly.
Samples from Ohio Riveronly. Ship at end of season.
Samples from Ohio River only. Ship at end of season.
Samples from Mississippi River only. Ship at end of season.
Samples from Mississippi River only. Ship at end of season.
Samples from Missouri River only. Ship at end of season.
Samples from Missouri River only. Ship at end of season.
Processed in lab of fish-sampling crew; tracked on MED form.
Carried to MED from crews' labs; tracked on MED form.
45
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Table 3-7. Sample tracking procedures.
1. Com plete the laboratory-specific sample tracking forms for the site (if it is sampled) including
a|[ samples collected regardless of whether they are to be immediately shipped to a lab or are
to be preserved and retained. Fill in the site ID, sample date, visit number, crew leader, and
team (see Section 2.3.2)
2. Fill in the sample ID for each sample and circle the sample type that applies. Enter the number
of containers holding the sample and (if applicable) whether the samples are to be shipped
immediately or retained by the crew and shipped as a batch or retained until the end of the
season. The condition code is filled in at the lab on receipt of the shipment.
3. Fill in the date that samples are to be shipped and the airbill number. If the form only includes
samples that are to be retained and not shipped immediately, do not fill in the date sent or the
airbill Number. Include a copy of the sample tracking form(s) in each container shipped to each
lab. Place forms in a self sealing plastic bag in the container. If the samples are delivered to a
FEDEX location directly after sampling before the form can be copied), a hand made copy of
the sample tracking form should be made and faxed in as soon as possible
4. Fax all the completed sample tracking forms (including inter-lab transfers and retained
samples) and the field verification form to the EMAP data manager (contact information in
Table 1 -2). The field verification form must be faxed in after every site visit, even if the site is
not sampled for some reason. The faxed sample tracking forms will be forward via email as
*.tiff files to the appropriate lab contacts so they will have a prior notification of sample
shipments.
5. Make copies of the sample tracking forms so a copy can accompany each shipment to the lab
and can be associated with every preserved and retained sample.
6. Ship the samples via overnight carrier. Friday samples should be shipped Monday morning for
same-day delivery. Retain the customer copies of the airbills in the master file for the site. The
laboratory will contact the field crews if the shipment does not arrive as expected, or if the
shipment does not match the sample tracking form.
7. When the retained samples are to be shipped or carried to the lab, fill in the Date Sent and
Airbill Number (if the information is missing) on the original sample tracking form, and re-fax
the form to Corvallis.
8. Return the original sample tracking form to the form packet for the site.
46
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Table 3-8. Checklist of supplies for base-location activities.
Qty
Item
Before departure to site
1
1
Site dossier
Instrument calibration supplies
Packing and shipping supplies (per
2
1-2
1-2
1
1-2
variable
1 roll
site)
1 -gallon self-sealing plastic bags for tracking forms
30-gallon plastic garbage bags to line coolers for shipping
Coolers for shipping samples
Containers suitable to transport and/or ship preserved
samples
Shipping air bills and sleeves
Block ice or ice substitute
Clear tape to seal coolers
3.6 Literature cited
Haney, J.F. and D.J. Hall. 1973. Sugar-coated Daphnia: a preservation technique for
Cladocera. Limnology and Oceanography 18:331-333.
Merritt, G.D, V.C. Rogers, and D.V. Peck. 1997. Base site activities. In J.R. Baker et al.
(editors) Environmental Monitoring and Assessment Program - Surface Waters, Field
Operations Manual for Lakes. U.S. Environmental Protection Agency, Corvallis, OR,
EPA/620/R-97/001.
Moulton, S.R. II, J.G. Kennon, R.M, Goldstein, and J.A. Hambrook. 2002. Revised protocols for
sampling algal, invertebrate and fish communities as part of the National Water-Quality
Assessment Program. Open-File Report 02-150, U.S. Geological Survey, Reston, VA.
Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and
Monaco, P. A. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Non-Wadeable
47
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Rivers and Streams, U.S. Environmental Protection Agency, Washington, DC.
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Ringold, P. L, Magee, T. and
Cappaert, M. R. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams,
U.S. Environmental Protection Agency, Office of Research and Development,
Washington, DC.
48
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Draft
TEAM:
EMAP-GRE TRACKING - MED
SITE ID: QRW04449- DATE COLLECTED:
1200
ANNUAL VISIT NUMBER: D 1 D 2 CREW LEADER:
AIRBILL NUMBER:
DATE SHIPPED:
/ 2 0 0
Sam pis H>
Sample Typ®
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK iS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
PP BK BS PA CV SV
*ol
€«i*iir«rs
Cowl,
e«ta
tNppwV
ftfMntd
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
Comments
Sample Types
PP = Phytoplankton
BK = Benthos kick
BS = Benthos snag
PA = Perlphyten
CV = Composite fish voucher
SV = Fish species voucher
Condition Codes
C = Cracked jar
F s Frozen
L = Leaking
ML = Mssin§ label
NP = Not preserved
W = Warm
OK =SimpIe OK
Filled in at lab
Chain of Custody
Date Received:
; /
Received by:
Filled in at lab
Contact Information
Tracking
f^arlys Cappaert
p) 541-754-4467
Lab
Terri M Jicha
US Environmental Protection Ageney- MEO
6S01 Congden Btvd
Pyluth, MM 55804
p) 218-529-5153
f) 218-529-4003
jiohsi.f enKH&pa ,gov
FAX TO
Tracking; i41-7§4-4637
Figure 3-2. Sample tracking form for samples shipped or carried to the Mid-Continent
Ecology Division (MED) Lab.
49
-------
• ^3—;
Draft
TEAM:
AIRBILL NUMBER:
Sample ID
EMAP-GRE TRACKING -NERL PAGE: QF •
^H
SITE ID: GRW04449- DATE COLLETED; / / 2 i 0
ANNUAL VISIT NUMBER: D 1 D 2
Sample Typ®
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
DS ST LT
Sample Types
DS = ONA
ST = Small fish tissue
LT = Large fish tissue
SG = Sediment grab
•
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
SG
Containers
Cond.
Code
CREW LEADER:
Shipped/
Retained
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
DATE SHIPPED: / / 2 0 0
Common name, photo file, or comments
Condition Codes
c
Ml
NF
W
C*
Fill
= Crocked Jar
Frosen
Leaking
- Missing lab
= Not presar
= Warm
sd in at
•el
lab
Chain of Custody Contact Information
Date Received: Tracking
, Msrlys Cappaert
Received by:
Lab
MmrM Smith
Sobran Environmental
National Exposure Research i-atooratof^
2i W, Martin Lutlwr (ting Dr
Filled in at lab
f) 513.569-7554
FAX TO Tracking; 541-754-4637 3
•
Figure 3-3. Sample tracking form for fish tissue and sediment samples shipped to the EPA
National Exposure Research Laboratory (NERL). Write the fish species collected
under "comments."
50
-------
1
1
• •LJPLJH
_ ^_ g_
^•••v
Draft S
TEAM:
AIRBILL NUMBER:
Sample V®
EMAP-GRE TRACKING - UMESC ^
TEID: 6RW0444S- DATE COLLECTED: / / 2 0 0
ANNUAL VISIT NUMBER: D 1 D 2
Sample Type
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
WC AL AP
*AII samples to be shipped ASAP
Sample Types
V¥C = Water chemistry
AL = Alkalinity
AP ~ Aquatic plants
I
#of
CREW LEADER;
DATE SHIPPED: / / 2 0 0
Condition Codes Chain of Custody Contact Information
C = Cracked jar
F s Frozen
L ~ Leaking
ML = Mlsslnfl label
NP = Not preserved
W = Warm
OK =Satnpl* OK
Filled in at lab
Date Received; Tracking
, , Wartys Cappaert
Received toy:
Lab
Shiriev Yuan. Chemist
U.S.G.S., Upper Midwest Environmental
Sciences Center
2830 Fanta !R@@d Road
Filled in at lab
tf| 608-783-6086
Xi&ol l_Vuan@usgs< g ov
FAX TO _
Tracking: 541-754-4837 4- I
1
Figure 3-4. Sample tracking form for water samples and aquatic plant specimens shipped or
carried to the Upper Midwest Environmental Sciences Center (UMESC).
51
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EMAP-6RE TRACKING - STROUD
Draft
TEAM:
SITE ID: GRW04449-
DATE COLLECTED:
/ 2 0 0
ANNUAL VISIT NUMBER: D 1 D 2 CREW LEADER:
AIRBILL NUMBER:
DATE SHIPPED:
1200
Simple IP
Staple TVpt
GF SS1 SS2
GF S31 SS2
GF SS1 SS2
GF SS1 SS2
GF SS1 SS2
GF SS1 SS2
QF SS1 SS2
6F SS1 SS2
GF SS1 SS2
GF SS1 SS2
GF SS1 SS2
GF SS1 SS2
GF SS1 SS2
GF SS1 SS2
SF SS1 SS2
*o»
eontalrwsr*
C«iet
Cods
comments
' Retain until end of field season. Be sure sample is dry.
Sample Types
GF = GeocHemical markers filter
SS1 = Total suspended solids filter pair 1
SS2 = Total suspended soli* filter pair 2
Condition Codes
C = Cracked jar
F = Frozen
L = Leaking
ML = Missing label
HP = Not preserved (wet)
W = Warm
OK =Sample OK
Chain of Custody
Date Received:
Received by:
Filled in at lab
Contact Information
Tracking
Marlys Cappaert
p) 541-754-446?
Lab
Mark Monk
Geochemistry Department
Stroyd Water Research Center
§70 Spenesr Road
Avondale, PA 19311
PJ810-5SI-21 53x27$
mmonh@straudcerrter.org
FAX TO
Tracking; 541-754-4637
Figure 3-5. Sample tracking form for filters shipped to Stroud Water Research Center.
52
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Draff
TEAM:
EMAP-GRE TRACKING - University of Louisville
SITE ID: GRW04449- DATE COLLECTED; / / 2 0 0
ANNUAL VISIT NUMBER: D 1 D 2 CREW LEADER:
AIRBILL NUMBER:
DATE SHIPPED:
/ 2 0 0
ID
Ssmpl® Type
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
CF BZ LZ
• of
containers
Cemci
Cade
Mpptty
Setalnfef
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
Cerarasnls
' Ship weekly
Sam pie Types
Condition Codes
Chain of Custody
Contact Information
CF = Chlorophyll inter
B2. = MaerozooplanMon {63 um)
LZ = Mta"G2Qoptankton |20 um|
C = Cracked jar
F = Frozen
L = Leaking
ML ss Hissing label
HP = Not preserved
W = Warm
OK =SampI» OK
Filled in at lab
Date Received:
Received by:
Filled in at lab
Tracking
Marlys Cappaert
p) S41-754-44«7
Lab
PEIchard Sehultz
Erwironmerrtal Analysis Laboratory
Stpartftwnt of Biology
139 Life Sciences
University of Louisville
Loutsviite KY
402§2 EXCEPT Fed EX uses 40208
p) 502-852-2543
f| 502-852-0725
reschu01@gwtssJouisviIle.edu
FAX TO
Tracking: §41-754-4637
Figure 3-6. Sample tracking form for all chlorophyll filters and Ohio River zooplankton
samples shipped or carried to the University of Louisville.
53
-------
EMAP-GRE TRACKING - IHJnOIS Nat. HJSt.
Drat
TEAM:
SITE ID: GRW04449-
DATE COLLECTED:
/ 2 0 0
ANNUAL VISIT NUMBER: D 1 D 2 CREW LEADER:
AIRBILL NUMBER:
OATE SHIPPED:
/ 2 0 0
Sample «3
sample Type
BZ LZ
12 LZ
BZ LZ
12 LZ
BZ LZ
1Z LZ
B2 L2
BZ LZ
B2 L2
BZ LZ
1Z LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
#ot
feartlstosFS
Send,
CM*
Comments
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
S R
* Shipped In batches
Sample Types
Bl s Maerozoopiankton (63 umj
LZ = Werozoopiankton |20 urn}
Condition Codes
C a Cracked Jar
F = Frozen
L = Leaking
ML = Missing label
NP = Not preserved
W = Warm
OK =S»mple OK
Filled in at lab
Chain of Custody
Date Received:
Received by:
Filled In at lab
Contact information
Tracking
IWarlys Cappaert
p) 541 -754-4467
Lab
Eric Rateliff
Illinois Natural History Survey
Great Rivers Field Station
8450 Monteliir Ave,
Brighton, IL 62012
p)«18-M&M90
f)818-46S-S68i
ratclif@lnhs,uluc,eeiu
FAX TO
Tracking: 541-754-4637
Figure 3-7. Sample tracking form for Mississippi River zooplankton samples shipped or
carried to the Illinois Natural History Survey, Great Rivers Field Station.
54
-------
m h^^d iMAP-GRE TRACKING - SMSU fj^
SITE ID: QRW04449- DATE: / / 2 0 0
TIAM:
AIRBILL NUMBER:
1
Sample ID
1
VISIT NUMBER: D 1 D 2 C
Snwipli Type
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
BZ LZ
5 or
Cond.
Code
3 D 4 CREW LEADER:
DATE SENT: / / 2 0 0
a™.*
Sample Types Condition Codes Chain of Custody Contact Information
8Z - Macrozooplankton (63 um}
L2 - Microzoopia^ton. (20 urn)
c =
r -f
Ml =
NP =
W ~
Fills
bracked j
rozen
eaking
Missing
Not pres
Warm
Sample
dim
af Date Received. Tracking
Wlartys Cappaert
Sniaf,s Received by;
Lab
-^ Kim Medley or John Havel
Department of Biology
Southwest Missouri State University
901 S. National Av.
, , , — -,| _, - , , , Springfield, Missouri 65SQ4 OSA
t lab Filled in at lab Pj w-wfrwos
johnhawil@smsu.edu jHavel)
FAX TO
Tracking; S41 -754-4637 8
I
1
Figure 3-8. Sample tracking form for Missouri River zooplankton samples shipped or carried
to Southwest Missouri State University.
55
-------
AERSILL NUMBER:
EMAP-GRE TRACKING - INTERLAB TRANSFERS
DftTESEHT: / / 2 0 0
Set* iD
Date Sample Collected
MM;DD«YYYY
Visit
Sampie iD
S^^ ft"**™.
TC SC Dm
TC SC DM
TC SC DM
TC SG Dm
TG SC Dm
TC SC Dm
TC SC Dm
TC SC DM
TC SC DM
TC SC DM
TC SC DM
TC SC DM
TC SC DM
TC SC DM
TC SC DM
eonci.
Code
Comment*
FAX TO
Tracking: 541-754-463?
Figure 3-9. Sample tracking form for inter-lab sample transfers. This form is used when
shipping samples between laboratories. SC = sediment chemistry subsample
shipped from NERL to MED, TC = preserved total organic carbon water sample
shipped from UMESC to MED, DM = preserved dissolved metals water sample
shipped from UMESC to MED.
56
-------
INTER-LAB SAMPLE
TRANSFER
sc TC DM
GRWQ4449-
/ 200
ID.
Figure 3-10. Label for inter-lab sample transfers. This label is only used when shipping
samples between laboratories. SC = sediment chemistry subsample shipped
from NERL to MED, TC = preserved total organic carbon water sample shipped
from UMESC to MED, DM = preserved dissolved metals water sample shipped
from UMESC to MED. Labels for shipping samples from the field are depicted in
subsequent sections. Not actual size.
57
-------
Blank page
58
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Section 4
Site Verification
E. William Schweiger1, Ted R. Angradi2, and David W. Bolgrien2
Site verification is the process by which 1) available background information for potential
EMAP-GRE sample sites is evaluated in the office, and 2) sites are visited to determine if a site
can and should be sampled. It includes procedures for adjusting sampling locations at each site
to avoid safety hazards and to correct for errors in sample location from the design. Parts of
this section are adapted from Herlihy (2003).
4.1 The EMAP sample design
EMAP uses an unequal selection probability randomized design to select sample sites
(for details, see http://www.epa.gov/nheerl/arm/). EMAP-GRE sample locations were selected
from a river-centerline GIS data layer (or "sample-frame") developed from the National
Hydrography Database (NHD). Sites were stratified by river, creating three explicit and
independent sample designs for the Missouri, upper Mississippi, and Ohio Rivers. Subsets of
sites within each strata were partitioned into sections defined by state boundaries (Missouri and
upper Mississippi Rivers) or multi-state river reaches (Ohio River) (see Figure 1-1). Strata and
sections are important in structuring how sites are replaced (see Section 4.1.1) and during data
analysis.
The list of sites generated through the design process is stored in a database called the
design file. The design file includes geographic (e.g., state, river section, river bank to sample)
and other coding fields identifying the characteristics of each sample location. All sites are
referenced with a Site ID number. Each crew will receive a copy of the design file. Additional
data fields may be added to a crew's copy of the design file to assist in field operations;
however, the structural relationships of the cells in a design file should never be altered (to
prevent a Site ID becoming associated with incorrect attribute fields).
National Park Service, 1201 Oakridge Drive, Fort Collins, CO 80525
U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
59
-------
Table 4-1. EMAP-GRE sample sites and site visits by river and section for 2004 and
2005 sample seasons. The total number of site visits is a sum of all initial site
visits and revisits within each section. To calculate the number of sites (or site
visits) in a specific state, sum all values in rows that include the state. For
example, Nebraska has sites shared with Missouri (5), South Dakota (10), and
Iowa (31), resulting in a total of 46 Nebraska sites (or 56 site visits).
Section Strata Section name
(river) (reach or
river left/river right state)
1 Lower
2 Ohio Middle
3 Upper
4 Illinois/Missouri
5 Illinois/Iowa
Upper Mississippi .... . .,
6 Wisconsin/Iowa
7 Wisconsin/Minnesota
8 Minnesota/Minnesota
9 Missouri/Missouri
10 Missouri/Nebraska
11 South Dakota/Nebraska
.„ Missouri ., . ... .
12 Montana/Montana
13 North Dakota/North Dakota
14 Iowa/Nebraska
15 Missouri/Kansas
Primary
site N
36
27
27
30
23
12
22
7
19
5
10
19
22
31
30
Total number
of site visits
49
33
32
40
28
17
24
9
23
5
15
22
22
36
37
Total number
of site visits
114
118
160
4.1.1 Site types and site replacement
Each section has a unique set of primary and oversample sites. All primary sites in a
section will eventually be sampled if they are not classified as non-target or unsampleable. If a
primary site is classified as non-target or unsampleable during the site verification process it
may not be sampled later, even if it becomes sampleable. Oversample sites are used to replace
non-target or unsampleable primary sites. Oversample sites are used for site replacement in
the order in which they appear in the oversample list within a section in most cases. However,
because some sections have a small expected primary sample size, it is possible that
oversample replacement sites in these sections may be from another section.
60
-------
As site verification forms are returned to the data information manager, the list of sites to
sample within a section will be updated and returned to each crew.
4.1.2 Panels
Primary sites within each section are divided by year into subsets called panels. Primary
sites should be sampled within their designated panel. Oversample sites acquire the panel
designation of the primary site they replace.
4.1.3 Site revisits
A subset of sites will be revisited two or three times. Data from these revisits are used to
refine the population estimates generated from EMAP sampling (for more details go to
http://www.epa.gov/nheerl/arm/). The design file designates which sites are revisited within
each strata. Allocation of revisits is across each strata (rivers). Therefore, individual sections
within a strata are not guaranteed any revisit sites (e.g., the Missouri/Nebraska section of the
Missouri River, Table 4-1). The revisit schedule is a combination of a "4-visit" and "3-visit"
approach (Table 4-2) which optimizes the allocation of effort between site-specific estimates
and the spatial distribution of revisits. Within each strata, the first four sites in panel 1 are
revisited three times (for a total of four site visits). The next six panel 1 sites are revisited two
times (for a total of three site visits). The first three of these six sites receive their intra-annual
visit in 2004 and their inter-annual visit in 2005. The second three sites are revisited twice in
2005. The first visit to revisit sites should be early in the sampling season (ideally they should
be among the first sites visited). Intra-annual revisits should be among the last sites sampled,
maximizing the time interval between visits to a site. If an oversample site is used to replace a
primary site designated a revisit site, the oversample site acquires the revisit schedule of the
original primary site.
61
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Table 4-2. Revisit schedule for each river, "x" denotes a site visit.
Panel 1
primary
site order
1
2
3
4
5
6
7
8
9
10
2004
Original visit
(early in index period)
X
X
X
X
X
X
X
X
X
X
Intra-annual 1
(late in index period)
X
X
X
X
X
X
X
not revisited in 2004
2005
Inter-annual 1
(early in index period)
X
X
X
X
X
X
X
X
X
X
Inter/lntra-annual 2
(late in index
X
X
X
X
not revisited
X
X
X
period)
in 2005
4.2 The EMAP-GRE sample site
Sampling at EMAP-GRE sample sites is done at point locations, within plots, or along
transects associated with an "X-site" location (latitude and longitude) from the design file
(Figures 4-1 and 4-2). Sample stations and their default (pre-adjustment) locations are as
follows:
1. Three main-channel sample locations. Water samples and plankton are
collected in the thalweg and at point locations half the distance from the
thalweg to each shoreline along a cross-channel transect oriented
perpendicular to the river centerline. The thalweg is defined as the line
connecting the deepest points in the main channel.
2. Two 500 m main-channel shoreline (MCS) transects. Each MCS transect
begins at the intersection of the cross-channel transect and the river right or
river left (facing downriver) MCS as designated in the design file. The primary
MCS transect extends 500 m upriver from this point. A secondary 500 m
MCS transect extends down river from the starting point of the primary
transect (Figure 4-1). Fish assemblages and fish habitat are sampled in the
near-shore littoral zone along both MCS transects. In addition, large woody
62
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debris, bank, and channel morphology measurements are taken along the
primary MCS transect.
3. Sample stations and plots along the primary MCS transect. Eleven
locations, spaced 50 m apart along the primary MCS transect (Figure 4-2),
define sample locations for riparian measurements, macroinvertebrates,
sediment and periphyton, bank morphology, and aquatic vegetation. Not all
indicators are sampled at all stations.
4. Other sample locations. A single macroinvertebrate sample is collected
from a snag (large woody debris in the channel) near the X-site. A subjective
assessment is made of the reach adjacent to the X-site.
4.3 The site dossier
Site dossiers contain aerial imagery and other site attribute information used in site
verification and sampling activities. Dossiers will be provided to crews by EMAP Duluth staff. An
example is given in Figure 4-3. Dossiers include approximate (GIS-derived) locations for the X-
site, cross-channel transect, and main-channel sample locations, and show the approximate
linear extent of the primary and secondary 500-m MCS transects. These data are displayed on
aerial imagery and in tables. Point coordinates are given in decimal degrees to facilitate
importation into GPS units. Crews may use a different local coordinate projection for
navigation, but all data must be reported as decimal degrees in NAD83 datum.
Dossiers contain a typographic error: main-channel locations for water and plankton
sampling locations identified as point ID numbers 2 and 3 in the data table in each dossier are
described ("detail") as "1/3" and "2/3" points. These points are actually half the distance
between the thalweg and the main-channel shoreline, and their locations are not accurately
characterized by a fraction of channel width. Point identified in the dossier as "250 m" and "500
m" points (ID numbers 7 and 8) can be ignored. These locations refer to sample locations
relevant for bathymetry procedures that have been dropped from EMAP-GRE.
63
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IVhin
and
Primary
transect
sample
lociion
Secondary
traised
Figure 4-1. Idealized sampling site. In this view, the target shoreline is at river left and the
NHD centerline and thalweg are in the same location.
64
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— Bank morphology
Q MacroinwrtebratB, periphyton
sediment
@ Aquatic vegetation
Primary j
_ and /
I \
T
transect
^^^^r
r K D-[
j n
i ...D.^1
H n
G n!
F n
D C
C .,D
B n
* * L_J
m
iim
i
fm
LU
i
Figure 4-2. Detail of the 500-m long primary main channel shoreline (MCS) transect showing
a subset of samples/measurements collected. Shoreline stations are spaced 50
m apart along the river margin.
65
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EMAP-GRE Site Dossier
GRW04449-300
River Thalweg Shoreline Transect
Points Lines: Attribute Information
JVAME NED ELEV I SEfTlors ^V\IPIE BINT* ( LOWEST RR FR MIIE I * PI ^NTm ISTTS POOL/REACH
DL1 UL
-, S r
J. ill Ll
2 > Pt ml
f i -,iCh irel T-H '? \\ Irf
LI-I*. U lu d lui 'If S lit
Trainee v
2 jn <«(,
Kin ^ilc
Pnmin ^f m ! p=trf rn^il ( Tr iise t
Secmdai\ »mEo n^tcanM jTiaii^cd
IRrfN
•\
Tr
^
ID
HO
-i i
#!-
Hi
„( S
-1
*,, -
tt< •,
' "
1i '
DESIGN NO
10
1
i
I
'
i
n
i )
'j
LOW DD
I! -> 1
•It f> 4
-> > -)<4^
Jli^-i!
(1 I'i-iS
Jli i^l
uii^j--
«) 14"-.
Ml ^
LIT DD
11 IS '
41 4ifl"
41 4j2"
11 fsJJ
41 4S4C
41 4 "
41 4->su
41 4>11
41 4i7^
OR AZ
lH r-T
SB UIS1
:
-------
4.4 Site verification
Site verification includes three phases for determining whether a site can and should be
sampled (its "sampling status"): 1) office-based verification, 2) field reconnaissance, and 3)
field-based verification. Each verification phase has a unique (but similar) data form. The
sampling status of a site is determined primarily by two factors: 1) whether or not the X-site falls
within one of the Great Rivers of interest, and 2) whether the site can be safely sampled. Site
verification results in the assignment of a status code to each site (Table 4-3)
Office-based verification and field reconnaissance occur before the sample season. The
completed verification forms must be returned to the EMAP data manager (Table 1-2) at least
two weeks prior to the first day of sampling. If a site is determined to be target, but is not
sampleable when the crew arrives for sampling, the EMAP-GRE design contact (Table 1-2)
should be consulted for guidance.
Site verification (both office and field phases) includes an evaluation of the main-channel
sample locations (see Section 4.2) and the MCS transects for position errors and any possible
safety hazards. Both office and field verification procedures include rules for adjusting sampling
locations when errors in their position or a safety hazard exists. These procedures should be
followed as closely as possible to maintain the validity of the EMAP sample design.
Non-target sites (NTS) are sites that fall in an Upper Missouri River reservoir (navigation
pools are considered river for EMAP-GRE). These sites will not be sampled. This status may be
apparent from office-based review, but given fluctuations in reservoir elevations, field
verification for sites in reservoir delta areas may be necessary. Local definitions and
professional judgement should be used to define the boundary between the river and the
reservoir into which it flows.
Target, not sampleable sites (TNS) are target (non-reservoir) but are unsafe to sample
even after all allowable adjustments (described in Section 4.4.4) to sample locations are
attempted. Classifying a site as TNS requires the following conditions: 1) If a site cannot be
safely electrofished, it is assumed that littoral and riparian sampling is also unsafe and the site
is classified as TNS; or 2) If it is unsafe to operate on the cross-channel transect because it is
67
-------
too close to a safety hazard (e.g., falls, dam, lock) then the site is TNS. If only a portion of the
cross-channel transect is unsafe, the sampleability is up to the discretion of the crew leader(s).
If electrofishing can be safely conducted, but some littoral or riparian stations appear unsafe,
the site should not be classified as TNS and all data that can be safely collected should be
collected.
Target, wrong strata (TWS) sites are sites near confluence areas that fall in the wrong
river. Target, wrong section (TWE) sites are sites that are in the wrong river section (state or
reach; Table 4-1). These sites may be sampled if safe. However, in the analysis phase, data
generated from the site will be grouped with their correct strata or section. An oversample site
that is in the correct strata or section must also be sampled to meet the primary N requirements
for the original strata/section. Crew leaders must determine if they want to sample a TWS or
TWE site and a correct strata or section oversample replacement site. TWS and TWE sites
should be identifiable through office-based review.
Sites that are not classified as NTS, TNS, TWS, or TWE during office verification are
classified as target, office-only sites (TOO) and do not require a separate reconnaissance visit
and field reconnaissance form. The final status of TOO sites is determined by field-based
verification at the time of sampling. Sites that are not classifiable with office-based verification
alone and that appear to be at risk of being NTS or TNS should be classified as unknown
(UNK) by office verification. These sites will require a separate field reconnaissance.
68
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Table 4-3. Site status codes.
Code
Meaning
Description
Applicable phase
NTS
Non-target
Non-target because the site falls in a
Missouri River mainstem reservoir.
Office-based
Field reconnaissance
Field-based
Office-based
TNS
TWS
TWE
TOO
UNK
TSA
TSS
Target, not
sampleable
Target,
wrong strata
Target,
wrong section
Target,
office only
Unknown
Target, sampleable
Target, successfully
sampled
sampling not possible due to safety hazards.
See Section 4.4.
Target but belongs to another strata. See
Section 4.4.1.
Target but belongs to another state or reach
section. See Section 4.4.1.
Probably target. High degree of certainty
allows omission of field reconnaissance.
Unknown from office verification alone,
requires field reconnaissance.
Verified with a field reconnaissance visit.
Target and successfully sampled.
Field reconnaissance
Field-based
Office-based
Field reconnaissance
Field-based
Office-based
Field reconnaissance
Field-based
Office-based
Office-based
Field reconnaissance
Field sampling
4.4.1 Office-based site verification
Office-based site verification relies on information in the site dossier, the crews'
experience, and local knowledge. Crews will evaluate and return an office-based site
verification form (Figures 4-4 and 4-5) for all primary sites in each section and for a subset of
oversample sites (approximately 10% of primary site N within each river strata). Office-based
site verification may be omitted for revisits. Procedures are described in detail in Table 4-4.
Although the sample design process will often place the X-site on or near the thalweg (the
line connecting points of maximum discharge in the main channel), errors may occur in its
position such as when the X-site falls away from the thalweg, outside of the main channel, or on
an island or bar. Furthermore, the NHD centerline (used to locate the X-site) may not
necessarily be parallel to the azimuth of either the thalweg or the MCS, causing the azimuths
used to create the cross-channel transect to be incorrect.
69
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The shorelines used to locate the MCS in the dossiers may also be incorrect due to the
resolution of the source data. These situations can lead to incorrect placement in all or some of
the sample locations in the dossier. Generally, office-based adjustment of incorrect sample
locations should be done in cases when an error in X-site position, cross-channel transect
azimuth, or MCS delineation is obvious and the possible solution is apparent in the dossier
image. In these cases, sample station locations may be adjusted by hand (e.g., by drawing on a
copy of the dossier) or in a GIS (data layers will be provided to crews by EMAP Duluth staff
upon request). If adjusted by hand on the dossier, a flag and explanation of these changes
must be provided on the office-based site verification form and the copy of the corrected
dossier returned with the office-based site verification form. If new values are calculated by a
crew in a GIS, these should be reported on the office-based site verification form. Office-based
site verification forms do not need to be faxed to Corvallis. They can be mailed or FedExed in
with the other field forms.
70
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Table 4-4. Office-based site verification procedures.
1. Fill out the header information on the office-based site verification form (Figure 4-5) including
the site ID, the date the form was filled out, the team (see Section 2.3.2), the verifier, and the
closest river mile (from the dossier).
2. Transfer the coordinates for the X-site, river left/right sample locations, MCS intersections, and
cross-channel transect azimuth from the dossier. Do not record negative numbers for longitudes.
Round dossier azimuths to the nearest degree.
3. Using all available data, answer the questions about apparent frame errors or safety hazards.
Answers to these questions assist with remaining steps in office-based site verification.
4. Assign a site status code to the site from among the options in Table 4-3. Sites classified as UNK
during office-based verification require field reconnaissance prior to sampling. Use flag/comments
to explain status assignments if necessary. Hazards include (but are not limited to) falls, rapids,
high and low dams, locks, and barge fleeting areas. A site is considered target, not sampleable
(TNS) if it appears that electrofishing cannot be safely conducted or if the main-channel sample
locations on the cross-channel transect cannot be safely sampled after all allowable adjustments
have been made (see steps 6-11). If only a subset of littoral or riparian sampling is prevented by
unsafe conditions, the site is still considered sampleable.
5. If the site is classified as NTS or TNS, explain the reasons in the comments. In this case, site
verification is complete; otherwise go to step 6.
6. Following the rules in Section 4.4.4, determine if a lateral shift or longitudinal slide is required for
either the primary and/or secondary MCS transect to avoid a safety hazard. During office-based
review, this should be done only as the quality of the dossier data allows. Shifts or slides may only
be apparent after going through steps 8-12.
7. Longitudinal slides only apply to MCS transects. The X-site and cross-channel transect may not
be slid up- or downriver.
8. On the provided imagery, examine the X-site, the river right/left points, the cross-channel transect,
and the shoreline intersection points. River right and river left are defined looking downriver. If the
location of the NHD centerline at the X-site, and the GIS-derived azimuth of the cross-section
transect and the left and right banks of the MCS appear valid, the intersection of the channel
cross-sections with the left and right bank should be approximately at the correct position. A "valid"
cross-channel transect azimuth will appear to be approximately perpendicular to the general
orientation of the thalweg (deepest point in the main channel) and/or the MCS. A "valid" MCS will
be reasonably close in position and orientation to the main-channel shoreline on the imagery. See
Section 4.4.4 for further details.
Continued
71
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Table 4-4. Office-based site verification procedures, continued.
9. If any of the locations from step 8 are suspect, a new cross-channel transect azimuth and
associated sample station locations should be drawn in by hand on the dossier (and/or in a GIS).
Suspect situations include an X-site that falls on an in-channel terrestrial feature or outside of the
bankfull channel, or the X-site on the NHD centerline appears to not fall in the thalweg. In all of
these cases, mark on the dossier image an adjusted position of the X-site - the "thalweg sample
location" in the thalweg closest to the original X-site along the cross-channel transect. An
approximate thalweg location can often be detected in imagery provided in the dossier (e.g., the
thalweg is generally in the navigation channel which may be apparent and is usually close to
outside bend shorelines). Estimate by eye the azimuth of the thalweg or MCS. Draw by hand
(and/or in a GIS) a new cross-channel transect line perpendicular to the thalweg or MCS and
through the X-site (or the substitute X-site). Estimate (or calculate in a GIS) its new azimuth and
record on the form.
10. The MCS transects should be evaluated and adjusted if necessary. The MCS on the imagery can
often be determined by examining the lateral positions of the most obvious or prominent bank line.
Draw a new MCS on the dossier maps (and/or in a GIS). With the thalweg location as a reference,
extend, if necessary, the cross-channel transect laterally to the new MCS. Place a point where the
cross-channel transect intersects the MCS by hand and/or in a GIS.
11. Examine the position of the river-right and river-left sampling locations on the cross-channel
transect (Figure 4-1). If they appear to be in the wrong location or unsampleable (e.g., on land)
they should be adjusted by moving them to the closest suitable position along the cross-channel
transect. The reasons for any adjustments should be recorded in the comment or explanation
sections of the office-based site verification form.
12. If any of the above adjustments are made in a GIS, record the new sample site coordinates on the
office-based Site verification form. If adjustments were made by hand, note this on the form
and include the marked copy of the dossier in the mailing to the EMAP data manager.
13. Record the name and location information for the boat ramp nearest the X-site.
14. (Optional) Repeat step 11 for a secondary ramp in case the primary ramp is unavailable or
unsuitable.
15. Use all available resources (e.g., consulting local resource contacts, maps, aerial photos) to
identify and describe any safety hazards associated with accessing the site.
16. Record contact information for any sources used to complete the office-based site verification
form.
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4.4.2 Field reconnaissance
Sites classified as unknown (UNK) during office-based verification require a field
reconnaissance visit prior to sampling. For sites that receive a reconnaissance visit, a
completed field reconnaissance form (Figure 4-6) must be returned to the data manager. Table
4-5 presents field reconnaissance procedures in detail. The primary purpose of field
reconnaissance is to establish the status of the site. However, to do this effectively, sample
locations may need to be visited, and the crew may elect to locate and flag the MCS transects
during the field reconnaissance visit.
4.4.3 Field-based site verification
Field-based site verification activities are largely analogous to office-based verification but
are conducted at the site. Table 4-5 presents field-based site verification procedures in detail.
Field-based site verification forms (Figures 4-7 and 4-8) are filled out for all site visits (including
revisits), regardless of whether the site is sampled or not. These forms are returned with all
other data forms from a sampling event.
4.4.4 Identifying the MCS and making adjustments in sample locations in the field
Identifying the MCS is a necessary step in establishing the set of sample locations at each
EMAP site. Once the MCS is identified, the sampleability (based on safety) of the 500-m
primary and secondary main-channel shoreline transects must be evaluated. To reduce bias in
sample location, all adjustments in the MCS transect, either laterally (shift) or longitudinally
(slide), must adhere to the rules below. Figure 4-4 presents a flow chart for identifying the MCS,
associated sample locations, and any adjustments that may be required.
The MCS is defined as the interface between the main channel (the channel with the most
discharge) and landward terrestrial habitat. It is usually the most obvious and closest shoreline
that borders the main channel. By definition, it is a terrestrial shoreline that an electrofishing
boat can safely navigate. In relatively straight, un-braided, single-channel reaches, the MCS will
likely follow a high bank that bounds bankfull channel flow. This situation is also likely in
reaches that are extensively modified (e.g., riprapped) and along outside bends where the main
channel is in direct contact with a high bank that separates floodplain terraces from the active
73
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channel. In complex reaches (deltaic, braided, or with at least two significant channels), or
where a new floodplain has formed below an abandoned pre-regulation floodplain (as has
occurred along the upper Missouri River), the MCS will not necessarily follow the high bank. In
these cases, the MCS is the interface between the main channel and terrestrial habitat that is
navigable for electrofishing and is closest to the position of the X-site moving laterally along the
cross-channel transect. The following rules must be followed when the MCS is shifted laterally:
1. If the target shoreline habitat adjacent to the X-site is an apparent island that is
obviously less than 2 km in length (estimated in the field orfrom the dossier), the
shoreline of the bar or island is probably not the MCS. The MCS will instead likely fall
on the next navigable shoreline landward along the line of the cross-channel
transect.
2. If there is no navigable channel (> 5 m wide; > 2 m maximum depth) along the MCS
landward of the apparent island, the MCS is located on the island regardless of its
size.
3. If the location of the thalweg is unclear (e.g., discharge in two channels appears
equal), and there are no other clues to the location of the thalweg (e.g., location of
the navigation channel) then the MCS transect is established on the shoreline
nearest to the original X-site location if the above rules are not violated. If the
nearest shoreline is not navigable, the MCS transect is established on the next
closest navigable shoreline.
4. Primary and secondary MCS transects are shifted together. For example, if the
primary MCS transect must shift landward to get off an island < 2 km in length, the
secondary MCS would also shift laterally to the same shoreline.
5. All 500 m of the transect is shifted together so that part of the 500 m transect is not
on one shoreline and the rest on another shoreline.
6. The opposite shoreline may not be used to replace the target MCS specified in the
dossier.
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Once the MCS is identified, the sampleability of the primary MCS transect (the 500 m
upriver from the intersection of the cross-channel transect and the MCS; see Figure 4-1) must
be evaluated. If a tributary or unconnected secondary channel (backwater or slack water)
greater than 5 m wide merges with the main channel or if there are less than 500 m of safely
sampleable MCS habitat along this reach, the entire set of MCS sampling stations should be
"slid" up- or downriver along the MCS a distance that places a sufficient buffer between
sampling stations and the safety hazard or tributary/secondary channel. For example, if a
navigation lock is 200 m upriver from the X-site or cross-channel transect, the 500-m primary
MCS transect can be slid approximately 400 m downriver from the X-site or cross channel
transect so that a 100-m buffer exists between the end of the transect and the lock. The degree
of hazard presented and the length of the buffer needed is up to the crew leaders (explain
decisions in comments on the field-based verification form). If necessary, all 500 m of the
primary and secondary MCS transects may be slid. If a connected secondary channel (running
water but not a tributary) greater than 5 m wide at the mouth and with a maximum depth > 2 m
is encountered along the MCS and it is safely navigable, the MCS transect will follow this
shoreline into the secondary channel. Tributaries and secondary channels less than 5 m wide at
their mouth are ignored when locating the MCS transect. Several rules apply when sliding the
MCS transect:
1. The MCS transects must not be slid more than 500 m up- or downriver so that the
both the up and downriver end of the primary MCS transect are > 500 m from
intersection of the cross-channel transect and the shoreline.
2. The opposite shoreline may not be used to replace the target MCS specified in the
dossier.
3. Safety or access issues that only apply to riparian plots are not grounds for sliding the
MCS transect. Unsafe or inaccessible riparian plots are not sampled (see Section 7).
4. If an up- or down-river slide in one of the transects will cause the primary and
secondary transect to overlap, the non-slid transect must be slid in conjunction with
the adjusted MCS transect. For example, if the downriver end of the primary MCS
transect is slid 100 m downriver, the upriver end of the secondary MCS transect is
also slid downriver 100 m so that they will not overlap.
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5. If a safety hazard or tributary or unconnected secondary channel > 5 m wide at the
mouth is encountered along the secondary MCS transect, the reach may be slid up-
or down-river to avoid the tributary or channel (following all the rules above). The
upriver end of the secondary transect must be < 500 m from the downriver end of the
primary 500 m MCS transect after sliding.
6. If the primary or secondary transect must be slid up- or down-river but the other
transect cannot be slid an equal distance because of a safety hazard, then the
transects can be slid in opposite directions (following all the rules above).
7. If the transects cannot be slid in opposite directions to avoid a hazard or tributary, the
secondary transect should be truncated in favor of the primary MCS transect (but is
still sampled to the degree possible). Flag any data not collected as missing and
explain the reasons.
There are many possible combinations of safety hazards and adjustments that may be
encountered in the field, not all of which are treated here. The operational goal of crew leaders
should be to conduct as much of the complete sampling suite as is possible without subjecting
the crew to unacceptable risk. Flag and explain any non-standard methods used or sampling
decisions made in the field. Do the best you can and consult the authors of this section for
additional guidance in special cases that do not seem to fit the circumstances described herein.
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Table 4-5. Field reconnaissance and field-based site verification procedures.
1. Fill out the header information on the appropriate form (field reconnaissance form or field-
based verification form) including site ID, the date the form was filled out, the team (see
Section 2.3.2), the verifier, and the closest river mile.
2. Transfer the original (dossier-based) or adjusted (from office verification) X-site coordinates,
cross-channel transect azimuth, and MCS intersections from the office-based site verification
form or field reconnaissance form. Do not record negative numbers for longitudes.
3. Using all available data, answer the questions about apparent frame errors or safety hazards
(or confirm the answers from the office-based site verification form/field reconnaissance form).
4. Assign a site status code to the site from among the options in Table 4-3. Sites classified as
UNK during office-based verification require field reconnaissance prior to sampling. Use
flag/comments to explain status assignments if necessary. Hazards include (but are not limited
to) falls, rapids, high and low dams, locks, and barge fleeting areas. A site is considered target,
not sampleable (TNS) if it appears that electrofishing cannot be safely conducted or if the main
channel sample locations on the cross-channel transect cannot be safely sampled after all
allowable adjustments are made (steps 5-12 below). If only a subset of littoral or riparian
sampling is prevented by unsafe conditions, the site is still considered sampleable. If the site is
classified as NTS or TNS, explain the reasons in the comments and site verification is
complete; otherwise move on to step 5.
5. Following the rules in Section 4.4.4 determine if a lateral shift or longitudinal slide is required for
either the primary or secondary MCS. The need for shifts or slides may only be apparent after
going through steps 8-12.
6. Longitudinal slides only apply to MCS transects. The X-site and cross-channel transect may
not be slid up- or down-river.
7. Navigate to the X-site using coordinates from the dossier, office-based site verification or field
reconnaissance forms. It is strongly recommended that the relevant site coordinates (i.e.,
points 1-6, 9 and 10 from the dossier) be pre-loaded into the boat's GPS unit.
8. If the location of the X-site is incorrect (e.g., not in the thalweg, on an island or the floodplain), a
new location should be established in the field. Locate the position of the estimated thalweg
closest to the X-site along the cross channel transect. Using the boat's GPS, acquire the
coordinates of the actual thalweg sample location and record them on the field reconnaissance
or field-based site verification form.
Continued
77
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Table 4-5. Field reconnaissance or field-based site verification procedures, continued.
9. Verify the azimuth of the thalweg and/or the general direction of the MCS while stationed on the
original X-site (if not moved in step 6) or generate a new one if the thalweg sample location is
not the same as the X-site. Maintain the boat position as close as possible to the X-site or its
substitute and back-sight along the direction of thalweg flow with a bearing compass. If the
safest or most convenient location is on the MCS, back-sight along the shoreline upriver.
Generate the approximate azimuth of the new cross-channel transect by calculating an azimuth
perpendicular (offset 90 degrees) to the thalweg/MCS orientation. Record this azimuth on the
field reconnaissance or verification form. If the original X-site is actually in the thalweg and the
field-calculated cross channel transect azimuth is within 5 degrees of the original cross-channel
transect azimuth, the original dossier azimuth value should be used.
10. While stationed on X-site or at the thalweg sample location (if they are not the same point),
visually extend the cross-channel transect laterally to the suspected MCS by sighting along the
bearing compass using the azimuth from the dossier/office-based site verification form or the
new value calculated in step 9. The intersection of the cross-channel transect and the MCS
should be identified using a fixed feature on the shoreline that is closest to the intersection.
11. Navigate to the river-right and river-left sample locations on the cross-channel transect. If they
appear to be in the wrong location (not half the distance from the shore to the thalweg) or are
unsampleable, they should be adjusted by moving them to the closest sampleable location
along the cross-channel transect. Record the final GPS location for each site on the form. Be
sure to note with a flag any adjustment made in the sample locations.
12. Navigate towards the MCS using the original or adjusted azimuth (from step 9). Determine if
the MCS as identified in the dossier or through office-based site verification is correct. See
Section 4.4.4 for a description of the MCS. If the MCS needs to be adjusted, locate the correct
MCS along the original or corrected azimuth. Verify or acquire new coordinates for the
intersection of the cross-channel transect and the MCS. Record these on the form.
13. Conditions apparently preventing complete and safe sampling of a subset of indicators do not
cause a site to be classified as unsampleable. All data that can be safely collected should be
collected from a given site.
14. All adjustments in sampling stations should be explained in the comment or explanation
sections of the site verification form.
15. Locate and flag the primary and secondary MCS transects (Section 4.5) and acquire the GPS
coordinates for the up- and down-river end of each MCS transect.
16. Make a sketch of the site (Figure 4-10), including any hazards, or frame error corrections. Take
digital photographs of the site (additional photos should be taken opportunistically during
sampling).
17. Record crew personnel.
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4.5 Locating and flagging the 500 m MCS transects
Because the position of one MCS can effect the other MCS, the two sampling crews
should communicate to make sure all adjustments are mutually understood. The first sampling
crew (fish or river) at the site locates and flags the primary and secondary 500-m MCS
transects after the intersection between the cross-channel transect and the MCS has been
verified and flagged using all the rules detailed above. If both crews arrive at the site at the
same time, crew leaders should decide which crew is responsible for flagging the stations. The
simplest method is to use the trip odometer on a GPS unit to determine distance along the
shoreline from the boat. A heavy steel washer with a flag streamer can be thrown from the boat
onto the bank at the station location to avoid landing the boat. Other methods may be used and
should be available as a backup if GPS reception is poor. For the purposes of EMAP-GRE,
dikes, wing dams, and other man-made structures that project into the channel are treated as
fish habitat at the site but are not treated as extensions of the river's shoreline. The contour of
the MCS cuts across the base of these structures. During site layout, stations on the primary
and secondary transect are flagged at 100-m intervals to 500 m. The intermediate littoral
stations on the primary MCS transect are spaced 50 m apart and are established during littoral
sampling.
4.6 Determining sampleability during the sampling visit
Even after site verification has indicated that a site is target and sampleable, conditions
encountered at the time of sampling may make sampling the site unsafe or impossible. The
most likely situations include high flows, high turbidity, barge activity, or gear failure. The
decision whether to sample on a given day is up to the crew leader(s). If the decision is made
not to sample, the crew should schedule another visit for the site and perhaps attempt to
sample an alternate target site. All reasons for aborting the site sampling attempt should be
noted on the Field-based site verification form Any data forms that are completed should be
returned (with appropriate flags) to the data manager. The field-based site verification form is
also filled out during site revisits.
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4.7 Site photographs
Site photographs should be taken at all sites using a digital camera (Table 4-6). In
addition to the recommended scenes, photos may also be taken of any stressors or other
unusual features at or around the site. Image file names (or the series) should be recorded on
the field-based verification form (Figure 4-10). All photos should be downloaded into a folder
named with the Site ID and backed up. The image files should be delivered to the EMAP data
center on a disk after the sample season.
Table 4-6. Recommended site photographs.
Image scene
Comments
X-siteand MCS
At cross-channel transect - MCS intersection: along MCS
looking upriver
At cross-channel transect - MCS intersection: along MCS
looking downriver
At cross-channel transect - MCS intersection: riverward
On riparian bank at bank station A looking landward
On riparian bank at bank station E looking landward
On riparian bank at bank station K looking landward
Other 1 -4
If possible, include a placard with site
name and sampling date.
Taken at the shoreline
Taken at the shoreline
Taken at the shoreline
Taken on the riparian bank
Taken on the riparian bank
On riparian bank
Stressors, notable features
4.8 Equipment and supplies
A list of the equipment and supplies required to conduct site verification and to lay out
the sampling reach is presented in Table 4-7. Generic supplies required for all EMAP-GRE field
sampling are listed in Table 2-5.
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Table 4-7. Equipment and supplies for site verification.
Qty
1
1
1 set
2 rolls
1
1
Item
Dossier of site and safety/access information
Gazetteers, topographic maps, and/or other navigation aids
Office-based, field reconnaissance, and field-based site verification forms
Biodegradable flagging, 2 colors (one for 100-m MCS intervals; one for 50-m
intervals)
Laser range finder (>1000 m range) (optional)
Digital camera and extra memory cards
4.9 Literature cited
EPA EMAP Design Web Site, http://www.epa.gov/nheerl/arm.
Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J., Lazorchak, J. M.,
McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and Monaco, P. A. Unpublished
draft. Environmental Monitoring and Assessment Program - Surface Waters Western Pilot Study:
Field Operations Manual for Non-Wadeable Rivers and Streams, U.S. Environmental Protection
Agency, Washington, DC.
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J., Lazorchak, J. M.,
McCormick, F. H., Peterson, S. A., Ringold, P. L., Magee, T. and Cappaert, M. R. Unpublished
draft. Environmental Monitoring and Assessment Program - Surface Waters Western Pilot Study:
Field Operations Manual for Wadeable Streams, U.S. Environmental Protection Agency, Office of
Research and Development, Washington, DC.
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Draft
EMAP-GRE OFFICE-BASED VERIFICATION (front) "«««"
toy flrtifsls):
SITE ID: GRW0444S- DATE: / / 2 0 0
TEAM:
ANNUAL VISIT
NUMBER:
D1 D2
VERIFIER:
Rl¥ER
MILE:
Sample Site Coordinates and Atonrth from Dossier Latitude Longitude AJmuth Flag
X-site DLef!
D Right
Coordinates of intersection of cross-channel
transect and main channel shoreline.
Coordinates of river right water
port, half way
between X-sle and river right shoreline.
Coordinates of let
quolity'pionkion sarr.p'e po n* half way
betwaen X-site and river left sbiyeiire.
Site Status and Sampling Stations Fiag
is the s-site located in a IV •SKX.n River mainsterr. "sen/air1' (non-target XIT> if >es i Y N Unk
Is the x-site located in a iwer different from the design file designation? (TWS if iss) Y N Unk
Is the x-site located in a section (e,g, . state, reach) d fferent from the design file designal on? (TWE If yesl Y N Unk
Is the x-site in the channel? Y N Unk
the x-site on the NHD to be in the tiatweg'? Y N Unk
Does the cross-channel transect - MCS intersection appear to be perpendicular to genera! main channel morphology? Y N Unk
Does the river right water qyaity/plenkton sample point appear to be sampleable? Y N Unk
Does the river left ivater quality/plankton sample point appear to be sampfeable? Y N Unk
Is the x-site a and of in comments}? Y N Unk
Does the main channel shoreline (MCSJ appear to be n the correct position? Y N Unk
Site Status Description - Check One
Q TOO - Apparently target using office-based verification Q TNS - Targe
lexplain in comments.). haards lex
D TWS - Apparen% target byt belongs to another Q NTS - Non-t
strata {explain in comments.) other reaio
Q] TWE - Apparently target but belongs to another Q] UNK - Unkn
section (state or reach.) (Explain in comments.)
New Coo
1 1 No Changes From Previous
Coordinates of ipitersection of cross-channel
transect and main channel shoreline.
of river right
quality/plankton sample point, half way
anil river right
Coordinates of river left water
sample point, half
between thalweg and river left shoreline.
rdinates and Aimuth from <
Latitude
t but unsampleabie due to safety
slain in comments.) Flan
i; explain in comments.)
swn. reqires FIELD RECONAISSANCE.
Jffice Verificaton
Longitude Aimuth
Flag
Flag codes; K~no measurement made. U^suspecf tneasurement;F1, F2. etc~mlsc flag® assigned by field crew. Explain in comments.
River ri§ht = Right shoreline as yoy look downstream
River left = Left shoreline as you look downstream
10.
Figure 4-4. Office-based site verification form (front).
-------
I
1 L |y bMAP-GKb OhHCfe-BASbi) VtKIHCAIIt
Draft
SITE ID: GRW04449- DATE: /
Name
Ramp Shoreline D River D Rivef Lei
Location from x-site d D
Distance from x-site (miles)
Nearest town
MI r-nnmm „ , , RiWtWSi _
INFORM (back) D^MH*): |
•
k
1 n n n ANMUAL VISIT
j ,2.°,°, , NUMBER: D1D2
D D Left
D D Downriver
Driving directions to primary boat ramp
information Sources • Names, Affiliations, Phone Numbers
Flag Comments {Description of MCS position, safety haards, etc.)
Flag code*: K=no measurement made, INsuspeet mtt*yrefn*nt;F1 , F2, ete=ml8c flap assigned by field crew. Explain In comments,
• •
Figure 4-5. Office-based site verification form (back).
83
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EMAP-GRE FIELD FORM (front)
Draft
SITE ID: GRW04449-
DATE:
/ 2 0 0
TEAM:
ANNUAL VISIT
NUMBER:
D1 D2
CREW
LEADER:
RIVER
MILE:
Sample Site Coordinates Worn Dossier or
Office Verification
Latitude
Longitude
Aimuth
Flag
X-site
n
Coordinates of of
transect and main channel shoreline.
Coordinates of river right weter
point, half way
between X-site and river right shoreline.
Coordinates of river left water quality/plankton
sample point, half wey X-site and
river left shoreline.
Site Status and Sampling Stations
Flag
Is the x-site located in a Missouri River mainstem reservoir? {non-target MTS if yes)
Y N ynk
Is the x-site in a river from the He (TWS if yes)
y N
Is the x-site in a (e.g., from ths file {TWE if yes)
Y N ynk
Is the x-site in the channel?
Y N ynk
Does the x-site on the NHO to be in the thatweg?
Y N Unk
the cross-channel - MCS intersection to tie to main
Y N ynk
Does the ri¥er water point to be sampleable?
Y N
the river let water to be sampfeable?
Y N Unk
Is the x-site near a safety hazard (describe distance from and nature of hazard in comments)?
Y N ynk
Does the main channel shoreline (MCS) as indicated on the dossier imagery match the field situation (see section *
Y N ynk
Site Status Description - Check One
LJ TSA - Target and sampleable.
Q TWS - Apparently target but belonfs to another
strata (river), (Explain in comments),
L~] TWE -Apparently to another or reach,
be sampled, {Explain in comments).
TNS - Target but unsampleable dye to safety
(explain in comments.)
UTS - Non-target (X-site in Missoyri Reservoir or
other reason, explain in comments.) Not sampled.
New Coordinates and Ainuth from Field Recon
Latitude
Longitude
Ainuth
Flag
D
No Changes
From Previous
Thalweg Sample
Locations
River right = Right shoreline as you look downstream
River left = Left shoreline as yoy look downstream
12,
Figure 4-6. Field-based reconnaissance form (front).
84
-------
•
ISPIS EMAP-GRE FIELD RECONNAISSANCE FORM (back) •
\ ml ^™
Dra« StTEIO:GRVTO4448- DATE: _ J_ f 200 ANN D 1 D 2
of
Flag Comments
• Flag code®: K - Ko messyrement mades U - Syspect messufement. F1,F2. etc. - misc. flugs assigned by etch field cfew. , _ ^H
Explain all flags in somnnent section, ' ^« ^H
Figure 4-7. Field-based reconnaissance form (back).
85
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Draft
FIELD-BASED VERIFICATION FORM (front)
SITEID:GRW04449- DATE: /
Reyiewetf
by (Inilialsj;
/ 2 § 0
TEAM:
ANNUAL VISIT D 1 D 2
CREW
LEADER:
RIVER
MILE:
Sample Site Coordinates from Dossier/Office
Verification/Field Recon
Latitude
Longitude
Aimuth
Flag
X-site
of of cross-channel
transect and main channel shoreline.
of river right
quality/plankton sample point, half way
X-site and river nght shoreline-
Site Status and Sampling Stations
Flag
Y N Unk
Is the x-site in a river different from the file designation"? (TWS if yes)
Y N Unk
Is the x-site located in a section (e.§,t slate, reach) different from the design fife designation? (TWE if yesj
Y N Unk
Is the in the
Y N Onk
Does the on the NHD to to in the thalweg'
Y H Unk
Does the cross-channel X to to to main
Y N Unk
Does the rwer right quality/plankton to be
Y N Unk
Does the river left water quatity'plankton sample to be
Y N Unk
Is the x-site near a safety hazard (describe distance from and nature of hazard in comments)?
Y N Unk
the main (MCS) as on the imagery match the 4.4.4)?
Y H Unk
Site Status Description - Check One
EH TSS - Target and sampled.
LJ TWS - Target but belongs to another strata. May be
(explain In comments.)
I I TWE - Target but belongs to another state or reach.
May be sampled {explain in comments.!
fl TNS - Target but unsampieable due to safety
haards (explain in comments.)
[~1 NTS - Non-tarpst (X-site In Missoyri Reser¥oir or
other reason, explain in comments.) Not sampled.
Flag
Flag
Comments (Description of MCS position, safety haards, etc.)
Flag codes: K = No measyrement made, U = Syspect measurement, F1.F2, etc. = misc. flags assigned by each Held crew.
Explain ail flags In comment section.
14.
Figure 4-8. Field-based site verification form (front).
86
-------
1
• ^>
Draft SITE ID: QRW04449- DAT6:
'ICATION FORM {back)
/ / 0 n ft ANNUAL VISIT
, , ,' , , ,',,.. , NUMBER:
1
•
^m
D1 D2
Final Sampled Coordinates and Almuth Latitucte Longitude Flag
1
Thalweg Sample locations .
Gc-ord naler;. of nver ngh* vvaier qu^!ity.;pionk;C'n sample
point, uilf say between itateeg art t:yerr ghi shoreline
" d' '11
point, talf way between thalweg ard river left shoreline.
Coord of end of MCS (Sits A)
Coordinates of end of MCS (Site K)
Coord of end of MCS fSOO rn)
Coord of end of secondary MCS (Om)
Sketch of X-site (indicate direction of flow!
FItos
Photo description
and MCS Site card)
At cross -channel transect - MCS intersection: along MCS ypriver
At cross-channel transect - MCS intersection: along MCS downriver
At cross-channel transect - MCS intersection: rwerwird
On riparian bank at bank station A looking landward
On riparian bank at bank station E looking landward
On riparian bank at bank station K looking landward
Otfier_
Other_
1
Filename
Flag
15, |
I
Figure 4-9. Field-based site verification form (back).
87
-------
Start
Are WQ
sample
locations hall
way between
thalweg and
MCS7
Is cross-
channel
transect
perpendicular
to main-
ohannef?
Does shore
appearto
be tie
actual
MCS?
Sbde shore transects
unlit fish able. See
slide rules in
Sect! on 4.4.4
Adjust channel
transect perpendicular
to main-channel at
closest point to X-site.
Adjust channel
transect station
locations.
Shift MCS to aetuai
yes. See Shift rules
in Sect!on 444.
Sliding does not yield
a ny safely fish abl e
shore.
No stations are
sample able.
Site is TOO until ft eld
verification (ISA) and
sampling are complete
(TSSJ.
Site status is TNS and
must be replaced.
Figure 4-10. Flow chart for adjusting layout of sites known to be target. MCS = main channel shoreline. Other acronyms defined in
Table 4.3.
88
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Section 5
Water Chemistry and Plankton
Terri M. Jicha1, Ted R. Angradi1, and Brian H. Hill1
In EMAP-GRE, water chemistry data will be used to define reference conditions and to
identify stressor gradients. Great River stressors associated with water chemistry may include
nutrient enrichment, inorganic contamination, hypoxia, temperature stress, turbidity, and
suspended sediment. Water chemistry sampling includes depth-integrated water samples for
laboratory analysis and depth-integrated in-situ measurements, including dissolved oxygen,
conductivity, and pH. At the base location, turbidity is measured, and chlorophyll, total
suspended solids (TSS), and geochemical marker samples are collected from subsamples of a
composite water sample.
Plankton includes algae (phytoplankton) and microinvertebrates (zooplankton)
suspended in the water column. Zooplankton are potentially useful indicators of environmental
condition. They are important to the food web of large rivers because they link primary
producers (algae) to larger invertebrates, and to fish and other vertebrates (Baker et al. 1997).
Plankton assemblage structure, body size distribution, and trophic structure are likely sensitive
to a number of anthropogenic disturbances, including flow regulation, habitat alteration
(including floodplain disconnection), invasive species, and contamination by nutrients, metals,
and herbicides.
Water chemistry and plankton sampling are combined in this section of the manual
because the activities are conducted together at the same sampling locations in the channel
and the data are recorded on the same forms. Water samples are collected from a transect
across the main channel rather than from littoral areas, because the emphasis is on an
unbiased and representative composite sample from the site rather than on relating water
chemistry to the biota at a site. Aspects of this section are adapted from Peck et al.
(Unpublished drafts).
5.1 Water samples
Water sampling is conducted by the river-sampling crew. The coordinates of the sample
U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
89
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locations are specified in the site dossier (Section 4). The three sample locations are in the
thalweg, and half the distance from the thalweg sampling location to each shoreline along the
cross-channel transect (Figure 4-1). The water sample consists of two depth-integrated 4-L
cubitainer samples (composite water samples 1 and 2) and one 500-mL grab sample.
A 2-L subsample of composite water sample 1 and 2, combined will be shipped to the
laboratory for determination of major cations and anions, nutrients, organic carbon, and
dissolved metals. The 500-mL sample is collected as a single grab at the thalweg sample
location and is used to determine total alkalinity. Subsamples from composite water sample 1
and 2, combined (the portion not shipped) will be used to measure chlorophyll a, geochemical
markers, TSS, and turbidity.
The 2-L composite water sample and the 500-mL grab sample are packed on ice in
coolers and shipped as soon as possible (Section 3.2.2) to the laboratory for analysis. The rest
of the composite water sample is processed at the base location. Procedures for collecting
water samples are described in Table 5-1. Subsample volumes for depth integrated composite
samples are given in Table 5-2.
Whenever possible, schedule site visits to occur in a downriver to upriver direction to
avoid re-sampling the same parcel of water. This is especially important when sampling a group
of adjacent sites over a short period of time.
5.2 Water-quality measurements
5.2.1 Dissolved oxygen, conductivity, pH, and temperature
In-situ measurements for DO, conductivity, pH, and temperature are made at each
subsample depth at each of the three sample locations at the site. The DO and pH meters are
calibrated daily at the base location or the sample site. Calibration requirements are described
in Section 3.
5.2.2 Water clarity and turbidity
Water transparency will be estimated using a Secchi disk at each sample location
(Table 5-3). Turbidity will be measured using a turbidimeter at the base location or mobile lab.
Turbidity readings in NTUs are made for a subsample from composite water sample 1 and 2,
combined (Table 5-4).
90
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5.2.3 Chlorophyll a, TSS, and geochemical markers
Chlorophyll a, TSS, and geochemical marker samples for laboratory analysis will be
collected by filtering a subsample from composite water sample 1 and 2 (combined) at the base
location or mobile lab. Geochemical markers include percent organic matter, percent nitrogen,
and stable isotopes (513C, 515N). The chlorophyll a filters must be kept refrigerated and dark
until frozen. Geochemical marker and TSS filters are dried before shipment. Procedures for
filtering water are described in Table 5-4.
5.3 Collection of plankton samples
Depth-integrated composite phytoplankton and zooplankton samples are collected in the
main channel at the water sampling locations. Phytoplankton samples are collected as a ~2-L
composite of pumped river water. Macrozooplankton samples are collected by pouring about
180 L of pumped river water through a plankton net with 63-um mesh. Microzooplankton are
collected by filtering 18 L of pumped river water through 20-um mesh. Microzooplankton
samples are not prefiltered through the macrozooplankton net. Table 5-5 describes plankton
sampling procedures in detail. Composite plankton samples are preserved with formalin
(phytoplankton) or a formalin-sugar solution (zooplankton) and stored until they can be
transported or shipped to the laboratory.
91
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Table 5-1. Procedure for collecting water chemistry samples.
1. Fill in the site ID and date on the front and back of the water chemistry and plankton form
(Figure 5-1 and 5-2).
2. Calibrate the DO meter (if the meter has not already been calibrated at the base location),
record calibration details in the log book. Indicate on the water chem istry and plankton form
(Figure 5-1) whether or not the DO and pH meter have been calibrated on the day of sampling.
3. Using a fine-point waterproof marker (e.g., Sharpie), fill out a pre-printed sample label (Figure
5-3) for one 4-L cubitainer and one 500-m L bottle (these containers will be shipped to the lab).
On each label, circle the sample type (WCC = water chemistry composite in cubitainer; ALK =
500-m L bottle), affix the labels to the cubitainer and bottle, and cover them with clear tape.
Record the sample ID numbers (from the labels) on the forms (water sample ID for the 4-L
cubitainer; alkalinity sample ID for the 500-mL grab). On a second 4-L cubitainer, write the site
ID and date directly on the cubitainer with a waterproof marker (this container is not shipped).
Containers may also be labeled at the base location prior to departing for the sample site. If
the site has been identified as a QA/QC (Section 5.4) site for water chemistry, additional
containers will be needed for field blanks and duplicate samples.
4. Attach the sounding weight to the winch cable. Attach the end of the hose from the peristaltic
pump (Tygon size 24) and the Guzzler pump (garden hose with check valve on the end) and
the DO/conductivity and pH probe(s) (or datasonde) to the cable above the sounding weight
(Figure 5-4).
5. Using the coordinates provided in the site dossier, navigate to the first sampling location
located halfway between the thalweg sample location and one of the shorelines (river-right or
river-left).
6. At the sample location, anchor if possible. Otherwise, the driver should hold the boat on the
sample location facing upriver. Remove the lids from both of the 4-L amber cubitainers and
pull them open. One cubitainer (label covered with tape) will hold composite water sample 1;
the other cubitainer (site ID written on side) will hold composite water sample 2. Do not blow
into the cubitainers to inflate them; this will contaminate the samples. Sun block, insect
repellent, etc., will contaminate the sample so avoid touching the inside of the cubitainer or cap.
7. From the sonar unit, determine and record the depth under the boat on the form. If the depth is
> 2 m, a subsample will be collected at 0.5 m off the bottom, at mid-depth and at 0.5 m from
the surface (each subsample is about 445 mL (Table 5-2): 4L composite volume/three sample
sites/three subsample depths). If the depth is < 2 m and > 1 m, subsamples will be collected
only at 0.5 m off the bottom and 0.5 m below the surface (each subsample is about 665 mL). If
the depth is < 1 m, the entire 1300-mL subsample for the location is collected at mid-depth.
The pumps may lack sufficient lifting power beyond about 6 m. If a sample location on the
cross-channel transect cannot be sampled, the subsample volume for each remaining location
should be adjusted so that the total sample volume for the site is maintained.
Continued
92
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Table 5-1. Procedure for collecting water chemistry samples, continued.
8. Lower the hose/sensor cluster to the deepest subsample depth using the depth dial on the
winch, by counting marks on the cable, or by allowing the sounding weight to contact the
bottom and then raising it to the proper depth. Adjust the depth to account for the distance
between the bottom of the weight and the sensors. Turn on the peristaltic pump and the
DO/conductivity/pH meter(s). Make sure the sounding weight is not bouncing along the bottom
during sampling. Use plastic cable ties to secure hoses and the sensor cable(s) to the winch
line as the weight is lowered. Pump overboard more water than the entire length of the
peristaltic pump hose can hold (determined beforehand). Once the hose is refreshed, pump
about 100 ml_ into each cubitainer to rinse it out. Repeat this two more times making sure rinse
water comes in contact with all interior surfaces.
9. Pump the first subsample (of 445, 665, or 1300 ml_ depending on the number of subsample
depths) into cubitainer 1 and cap it. Use graduations on the cubitainer to estimate when the
subsample volume is attained. Pump a second subsample of equal volume into cubitainer 2
and cap it (total of 888 ml_ per depth at each sample location). Record sample depth, DO
(mg/L) and conductivity (uS/cm), temperature and pH following procedures in the instruments
operating manuals. Record the actual sample depth (from the surface). Flag depths (K flag) for
which no instrument readings are made.
10. Collect phytoplankton and zooplankton subsamples using the procedures described in Table 5-
5.
11. Repeat steps 6-9, as appropriate, for the other subsample depths at the sample location.
Record the total depth or sample depth each sample location on the form. Estimate Secchi
depth (Table 5-3).
12. Repeat steps 6-10 for the two other water quality sample locations at the site. By sliding the
cable ties down the winch line as the line is winched up, it should be possible to move between
stations without having to disassemble the sampling apparatus. At the thalweg sample location
only, collect a 500-mL grab sample for alkalinity. Rinse the bottle and cap three times with river
water (discard downriver). Take the sample by filling and capping the bottle under water at
arms-length; cap the bottle underwater with no head space. Alternatively, fill a clean bucket
with river water from the surface at the thalweg location and take the sample by filling and
capping the bottle under water.
13. After all the water and plankton has been collected, the boat can be beached at the first
shoreline sample station for sample processing. Place the water samples in a cooler with ice.
5.4 QA considerations for water chemistry and plankton sampling
Instrument calibration and maintenance, avoiding contamination, and proper container
labeling and tracking are all essential for maintaining high QA standards for water chemistry
sampling. Table 5.6 describes some specific QA considerations for water chemistry sampling.
At the second site sampled and at the 11th and 21st site sampled by a crew in a season,
field duplicates and field Dl blanks are required. Field duplicates and blanks are processed and
tracked using the same procedures as for regular water samples.
93
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5.5 Equipment and supplies
A list of the equipment and supplies required for collecting water and plankton is
presented in Table 5-7. Generic supplies required for all EMAP-GRE field sampling are listed in
Table 2-5.
Table 5-2. Subsample volumes for depth-integrated composite water samples.
Volumes are totals for each depth for each of three sample locations on the
cross-channel transect. If a sample location on the cross-channel transect
cannot be sampled, the subsample volume for each remaining location should
be adjusted so that the total sample volume for the site is maintained. This table
should be reproduced and affixed to the top of the peristaltic pump or other
convenient location for reference in the field.
Sample type
(Total volume per site)
Water chemistry
(8L)
Phytoplankton
(1935 ml_)
Macrozooplankton (63-um mesh)
(180L)
Microzooplankton (20-um mesh)
(18 L)
Total depth at sample location
> 2 m
(3 depths)1
888 ml_
215 ml_
20 L
2 L
< 2 m and > 1 m
(2 depths)2
1333 ml_
322 ml_
30 L
3 L
< 1 m
(1 depth)3
2667 ml_
645 ml_
60 L
6 L
1 Collect a subsample 0.5 m above the bottom, at mid-depth and 0.5 m below the surface.
2 Collect a subsample 0.5 m above the bottom and 0.5 m below the surface.
3 Collect a subsample at mid-depth.
Table 5-3. Procedure for determining Secchi depth (after Strobel and Heitmuller 2001).
1. Remove sunglasses. If the water is relatively clear, lower the Secchi disk over the shaded side
of the boat until it disappears. Slowly raise the disk until it becomes visible and record the
depth indicated on the line, (interpolate to the nearest 10 cm). If the disappearance depth is <
0.5 m, retrieve the disk and go to step 2.
2. Use the "Secchi on a stick" and read the disappearance depth from the scale on the stick to the
nearest cm. In current, it may work best to estimate Secchi depth while drifting.
3. If the disk is visible resting on the river bottom, mark the appropriate box on the form.
94
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Table 5-4. Base-location procedures for turbidity measurements and for subsampling
and filtering a water sample for chlorophyll a, TSS, and geochemical
markers (based, in part, on personal communication with Anthony
Aufdencampe, Stroud Water Research Center, Avondale, PA).
1. Shake composite water samples 1 and 2 vigorously and completely pour both composites into
an 8-L plastic churn-splitter. Use a subsample dispensed from the churn to rinse the
cubitainers into the churn. Churn for five strokes with the dasher touching the bottom of the
churn on each stroke. Dispense subsamples while continuing to churn.
2. Dispense 2000 ml_ from the churn back into the labeled 4-L cubitainer that originally held
composite water sample 1. Cap the cubitainer and return it to the cooler with ice. This is the
sample that will be shipped to the lab for analysis.
3. Turbidity. Dispense about 75 m L of composite from the churn into a beaker for turbidity
analysis. Follow the operating instructions for the turbidimeter and make three replicate
readings (three subsamples from the 75 ml_ of composite). Record the values in NTUs on the
water chemistry and plankton form. Record the temperature of the sample. Clean sample tubes
as required in the turbidimeter operating instructions.
4. Set up the filtering apparatus by connecting a vacuum pump to the filter reservoir.
5. Ch lorophyll a. Filter chlorophyll in shade or subdued light. Place the filter holder on the
reservoir and position a 47-mm Whatman GF/F glass-fiber filter on the manifold (not pre-ashed
or pre-weighed). Handle filters with forceps. The top of the filter is the side opposite the
"checked" or "gridded" side (i.e., filters are placed "grid to grid" on the manifold). Wetting the
manifold screen with Dl water will allow the filter to adhere better. Secure the top of the
apparatus.
6. Dispense (while churning) between 50 to 250 ml_ at a time using a graduated cylinder. If the
water appears clear, dispense 250 ml_ at a time; if the water appears turbid or green, dispense
50 to 100 mL at a time. Turn on the vacuum pump (or start pumping with a hand pump). Try
not to exceed a vacuum of 7 psi (15 inches of Hg) during filtration. Pour the entire contents of
the graduated cylinder into the filter funnel each time. Continue dispensing until the filters
begins to clog and filtration slows. Keep track of the volume dispensed (to the nearest mL). It
is important to filter as much water as possible, in order to collect enough sample for analysis.
The total volume filtered can range from 50 mL to 1500 mL, depending on turbidity. If the filters
clog completely before all the sample in the filter funnel has been filtered, discard the sample
and filter and prepare a new sample with a smaller volume of water.
7. Record the final volume filtered to the nearest mL on the form and sample label (there is no
filter ID for chlorophyll filters). Remove the filter with forceps and place it in a foil-wrapped
scintillation vial. The filter may be folded in half.
8. Geochemical markers. Place the filter holder on the reservoir and position a pre-weighed and
pre-combusted (450° C, 4-6 h) 47-mm Whatman GF/F glass-fiber filter on the manifold. Handle
filters with forceps The top of the filter is the side opposite the "checked" or "gridded" side (i.e.,
filters are placed "grid to grid" on the manifold). Record the filter ID number from the filter
container on the water chemistry and plankton form. Wetting the manifold screen with Dl water
will allow the filter to adhere better. Secure the top of the apparatus.
Continued
95
-------
Table 5-4. Procedure for filtering a water sample for chlorophyll a, TSS, and
geochemical markers, continued.
9. Dispense (while churning) between 50 to 250 ml_ at a time using a 100 or 250 ml_ graduated
cylinder. If the water appears clear, dispense 250 ml_ at a time; if the water appears turbid or
green, dispense 50 to 100 ml_ at a time. Turn on the vacuum pump (or start pumping with a
hand pump). Try not to exceed a vacuum of 7 psi (15 inches of Hg) during filtration. Pour the
entire contents of the graduated cylinder into the filter funnel each time. Continue dispensing
until the filters begins to clog and filtration slows. Keep track of the volume dispensed (to the
nearest ml_). It is important to filter as much water as possible, in order to collect enough
sample for analysis. The total volume filtered can range from 50 mL to 1500 ml_, depending on
turbidity. If the filters clog completely before all the sample in the filter funnel has been filtered,
discard the sample and filters and prepare a new sample with a smaller volume of water.
10. Record the filter ID and final volume filtered to the nearest mL on the form and the EPA sample
label (transfer the filter ID from the small label on top). Remove the filter funnel with the
vacuum still on. Turn the vacuum off and carefully peel the filter off the manifold screen and
return the filter to its container (dirty side up) using blunt forceps. If the filter tears, start over.
11. Total suspended solids (TSS). Place the filter holder on the reservoir and position a pair of
pre-weighed 47-mm membrane filters on the manifold (they will be labeled and prepackaged as
a pair). Handle filters with forceps and make sure the top filter from the filter holder is on top in
the manifold. Wetting the manifold screen with Dl water will allow the filters to adhere better.
12. Record the filter pair ID number on the water chemistry and plankton form. Secure the top of
the apparatus.
13. Dispense (while churning) between 50 to 250 mL at a time using a 100 or 250 mL graduated
cylinder. If the water appears clear, dispense 250 mL at a time; if the water appears turbid or
green, dispense 50 to 100 mL at a time. Turn on the vacuum pump (or start pumping with a
hand pump). Try not to exceed a vacuum of 7 psi (15 inches of Hg) during filtration. Pour the
entire contents of the graduated cylinder into the filter funnel each time. Continue dispensing
until the filters begins to clog and filtration slows. Keep track of the volume dispensed (to the
nearest mL). It is important to filter as much water as possible, in order to collect enough
sample for analysis. The total volume filtered can range from 50 mL to 1500 mL, depending on
turbidity (membrane filters have a 10-20% lower capacity than OFF filters). If the filters clog
completely before all the sample in the filter funnel has been filtered, discard the sample and
filters and prepare a new sample with a smaller volume of water.
14. Record the filter ID and the final volume filtered to the nearest mL on the form and on the EPA
sample label (transfer the filter ID from the small label on top). Remove the filter funnel with the
vacuum still on. Turn the vacuum off and carefully peel the filter off the manifold screen and
return the filter to its container (dirty side up) using blunt forceps. If the filter tears, start over.
15. Repeat step 11-14 for a second TSS filter pair. The total volume filtered should be about the
same as for the first TSS sample.
16. Rinse the filter apparatus, churn, and graduated cylinder with Dl water.
96
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Table 5-4. Procedure for filtering a water sample for chlorophyll a, TSS, and
geochemical markers, continued.
17. Complete filling out sample labels (Figure 5-3) for each filter or filter pair (TSS). Place a label
on the chlorophyll vial and cover with tape. Place the labels on the bottom of the"Petrislide"
containers (TSS and geochemical marker filters). Be sure they do not cover the lid or obscure
other information. Keep chlorophyll filters cold (near 4° C) until they can be frozen.
18. Chlorophyll filters are preserved by freezing. Geochemistry and TSS filters are preserved by
drying in an oven. Dry filters overnight in a drying oven at 30 - 50° C with the Petrislide
container upright and the lids ajar (the Petrislide containers will melt at >70° C). Drape a sheet
of aluminum foil over the filters to keep out dust. If a drying oven is not available, filters will
usually dry in 24-48 h in an air-conditioned room by setting the filters out on a table with lids
ajar and a sheet of aluminum foil draped on top to keep out dust. Store dried filters in a dry
location prior to shipment. Do not ship filters that have not been dried.
97
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Table 5-5. Procedures for collecting plankton samples. Plankton sample collection is
integrated with water sample collection (Table 5-1).
1. Navigate to the first sampling location half the distance between the thalweg and one of the
shorelines.
2. Phytoplankton. Follow steps 1-8 in Table 5-1 (Procedure for collecting water chemistry
samples). After the water subsample has been collected, continue pumping water at that depth
for a phytoplankton sample. Pump a subsample into a graduated cylinder. Total sample
volume at each location will be 650 ml_ (1,950 L total sample volume/three sample locations -
total volume is less than 2 L to allow room for preservative ). Subsample volume at each depth
will be =21 5, 322, or 645 ml_ (Table 5-2), depending on the number of depths at the sample
location (e.g., 322 ml_ = 1,935 ml_ total composite/three sample locations/two depths). Pour the
subsample into a 2-L bottle and cap the bottle (this same bottle will hold the entire composite
phytoplankton sample for all depths and all three sample locations).
3. Microzooplankton. After the phytoplankton sample has been collected, continue pumping at
that depth into the graduated cylinderfor a microzooplankton sample. Total microzooplankton
sample volume for each location is 6 L, so if there are three depths to be sampled, the
subsample volume will be 2 L; for two depths the subsample volume will be 3 L; and for 1 depth
the subsample volume will be 6 L. Pour the subsample from the graduated cylinder into a PVC
microzooplankton filter with 20-um mesh. Turn off the peristaltic pump. Alternatively, the
Guzzler pump (see step 4) can be used to collect the microzooplankton composite.
4. Macrozooplankton. Pump the Guzzler pump to prime it. Pump over-board for about 15
seconds to refresh the hose contents. Total sample volume for each sample location is 60 L,
so if there are three depths, the subsample volume will be 20 L for a macrozooplankton
sample; for two depths the subsample volume will be 30 L; and for 1 depth the subsample
volume will be 60 L Table 5-2. Pump into a graduated plastic bucket and pour the bucket
contents through the plankton net (63-um mesh) 10 L at a time.
5. Repeat steps 2-4 for each depth at the sample location. Be sure to pump water overboard
before collecting plankton samples to refresh the hose contents at each new depth. Keep track
of the volume of water pumped for zooplankton.
6. Using a minimum amount of filtered (20-um mesh) river water from a wash bottle, rinse the
contents of the microzooplankton filter into a 100-mL bottle (use a funnel) and cap the bottle
(this same bottle will hold the entire composite microzooplankton sample).
7. Wash the (macro)plankton net contents down to the cod end using the on-board washdown
pump. Release the pinchcock and discharge the contents of the cod end into a 250-mL sample
bottle. Using a minimum amount of filtered river water from a wash bottle, rinse the cod end
into the sample bottle and cap the bottle (this same bottle will hold the entire composite
macrozooplankton sample).
8. Repeat steps 2-7 at each sample location. By sliding the cable ties down the winch line as the
line is winched up, it should be possible to move between stations without having to
disassemble the sampling apparatus. Be sure to pump water overboard before collecting the
sample to refresh the hose contents at the new location/depth.
Continued
98
-------
Table 5-5. Procedures for collecting plankton samples, continued.
9. Using a small beaker, add 80 mL of borax-buffered 100% formalin to the 1,950 ml_
phytoplankton composite (4% formalin final concentration). Use gloves and safety glasses
when handling formalin.
10. Using filtered (20-|jm mesh) river water, raise the level in the macrozooplankton (250 mL) and
microzooplankton (100 mL) sample bottles to 2/3 full. Put half of an Alka-Seltzer® tablet in
each bottle and let it dissolve. Top off each bottle with the chilled 12% borax-buffered formalin-
sugar solution to achieve a final preservative strength of 4% formalin. Record the total volume
filtered for macrozooplankton and for microzooplankton on the water chemistry and plankton
form.
11. Prepare labels (Figure 5-3) for outside the jars (if not pre-labeled). Fill in the site number, enter
the sample date, and composite sample volume (phytoplankton) or the volume filtered
(zooplankton) and site visit number. Place the labels on the jars and cover them with clear
tape. Record the sample ID and other data on the water chemistry and plankton form.
12. Seal each capped jar with plastic electrician's tape. Store the preserved samples upright in a
container to await transport or shipment to the laboratory (see Table 3-7).
99
-------
Table 5-6. QA considerations for water chemistry and plankton sampling.
The laboratory water quality analyses are very sensitive, and attention to detail is needed to
avoid sample contamination. Possible sources of contamination include substances on
sampler's hands, boat exhaust, and sediment from the river bed.
Be vigilant about refreshing the hose contents at each new depth or sample location.
Cap the composite water samples (4-L cubitainers) between subsamples to avoid atmospheric
contamination.
Keep the 12% formalin-sugar solution (for preserving zooplankton) on ice in the boat.
Whenever possible, schedule site visits to occur in a downriver to upriver direction to avoid re-
sampling the same parcel of water. This is especially important when sampling a group of
adjacent sites over a short period of time.
Calibrate the dissolved oxygen meter before or during every site visit.
Avoid contacting the river bed with the sounding weight while collecting water and plankton.
Avoid sending the sounding weight down through or adjacent to snags. If snags are observed
or suspected under the boat, move several meters away to sample and flag the samples.
If possible, the same crew member should estimate Secchi depth at each sample location and
site.
Using a survey pole or other gage, occasionally confirm the accuracy of the boat's sonar depth
readings.
Always pour the entire 4-L grab samples (water chemistry composites 1 and 2) into the churn
for subsampling. Rinse the cubitainers into the churn with an aliquot from the churn.
Make certain that the TSS and geochemical marker filters are thoroughly dried before shipping
them to the lab.
100
-------
Table 5-7. Equipment and supplies for water quality and plankton sampling. Generic
supplies required for all EMAP-GRE field sampling are listed in Table 2-5.
Qty
1
1
1
1
1
25
1pr
1
1 set
4
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
Item
Winch with calibrated cable or depth dial and 30, 50, or 100 Ib sounding weight depending on
water velocity
Masterflex 1 1 5V peristaltic pump or equivalent with 1 0 m of size 24 Tygon hose (Fisher Scientific
1 3-31 0-490 or equivalent)
Guzzler hand pump (Model 400h with aluminum epoxy coated clamp ring, 3/4" mail garden hose
fittinqs and 74MGH check valve on submerged hose end) wvvw.thebosworthco.com. Mount the
pump on a board large enough to step on with 2 feet.
3/4" flexible garden hoses (10m section with female fittings both ends, 2 m section with 1 female
fitting)
Modified sounding weight hanger or other apparatus for attaching sensors and hoses to winch line
(See Figure 5-4)
Plastic cable-ties long enough to secure hoses and sensor cables to the winch line or cable
Side cutters for removing cable ties
Nephelometric turbidimeter (used at the base location)
Turbidity standards (e.g., 1, 10, 100, 250 NTU)
4-L amber pre-DI-washed cubitainer for water samples. Only two are needed per site; extras
should be carried in case of contamination (Fisher Scientific 1 1-375-1 15B or equivalent).
500-mL Nalgene bottle for pH and alkalinity subsample
DO/Conductivity meter with 7.5-m cable and manufacturers manuals
pH meter with 7.5-m cable and manuals
200-mm Secchi disk with calibrated chain or line (0.5-m increments)
200-mm Secchi disk affixed to a calibrated stick (0.01 -m increments)
Plankton tow net with 63-um mesh net, drain hose and pinchcock (Wildco 426-A28 or equivalent)
250-mL plastic graduated cylinder for filtering samples
100-mL plastic graduated cylinder for filtering samples
Graduated plastic bucket
20-um mesh plankton filter (sieve or home made from PVC pipe, coupler and Nitex). Put a bead
of silicone caulk around inside edge of pipe where it meets the Nitex.
Square HOPE sample jars, 250-mL capacity (for macrozooplankton samples) (Fisher Scientific
03-31 1-3D or equivalent)
Square HOPE sample jars, 1 00-mL capacity (for microzooplankton samples)
(Fisher Scientific 03-31 1-3C or equivalent)
Square HOPE bottle, 2-L capacity (for phytoplankton samples) (Fisher Scientific 03-31 1-3G or
equivalent)
Vacuum pump
8-L churn sample splitter (Wildco 1831-C80 or equivalent)
Continued
101
-------
Table 5-7. Equipment and supplies for water quality and plankton sampling,
continued.
Qty
1
1
1
1
1
1
1 box
1
1
2pr
1 pr
2
1
1 set
1 L
1
1 L
2
1 roll
Item
Lab thermometer for turbidity samples
Chlorophyll filter apparatus (including tubing) (Fisher Scientific 09-740-23E or equivalent)
Whatman GF/F filters (47 mm) for chlorophyll analysis (Fisher Scientific 09-874-71 or
equivalent)
Pre-weighed and pre-combusted (450° C, 4-6 h) Whatman GF/F filter (47 mm) for
geochemical markers in a Millipore Petrislide container (Provided by lab).
Pre-weighed membrane filter pair (47 mm, 0.45 urn) for suspended sediment
in a Millipore petrislide container (Provided by lab).
Scintillation vials for chlorophyll filters (Fisher Scientific 03-337-14 or equivalent)
Aluminum foil to wrap chlorophyll vials
Small funnel for transferring plankton samples from nets into jars
Wash bottle
Powder free lab gloves
Filter forceps
100-mL plastic beakers for turbidity procedures and for adding formalin to phytoplankton
samples (Fisher Scientific 02-591-27 or equivalent)
Water chemistry and plankton form
Sample labels
Borax-buffered formalin (100%) for preserving phytoplankton
Gloves, safety glasses and apron for handling formalin
Buffered formalin-sugar solution (12%) for preserving zooplankton
Alka-Seltzer® tablets to anaesthetize zooplankton prior to preservation
Plastic electrician's tape for sealing plankton sample jars
5.6 Literature cited
Baker, J.R., D.V. Peck, and D.W. Sutton (editors). 1997. Environmental Monitoring and
Assessment Program Surface Waters Field Operations Manual for Lakes. EPA/620/R-
97/001. U.S. Environmental Protection Agency, Washington, DC.
Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and
Monaco, P. A. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Non-Wadeable
Rivers and Streams, U.S. Environmental Protection Agency, Washington, DC.
102
-------
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Ringold, P. L, Magee, T. and
Cappaert, M. R. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams,,
U.S. Environmental Protection Agency, Office of Research and Development,
Washington, DC.
Strobel, C.J. and T. Heitmuller. 2001. National Coastal Assessment Field Operations Manual.
U.S. Environmental Protection Agency, EPA/620/R-01/003. 71 p.
103
-------
SITI ID: GRW04449-
EMAP-GRE WATER CHEMISTRY AND PLANKTON (front)
DATE: / / 2 0 0
ANNUAL VISIT
NUMBER:
n1
LJ '
Water Chemistry
Water Sample
DO/PH Calibration
Sample ID
2 L composite
Altitude at Calibration (m)
Sample
2 L composite 1
D Yes
SOD mL grab
D Yes
Composite of 3
Stations?
Was DO meter calibrated on day of sampling? Q] Yes Q No
D Yes D No
D Yes D No
Was pH meter calibrated on slay of sampling? n Yes Q No
2 L Composite Field
Duplicate Sample ID
Alkalinity
Sample ID
01 Blank Sample ID
Alkalinity
Duplicate
Sample ID
Water Quality Measurements
"' Set below for rules
on which depths to
take readings;f!ag
depths not used.
0.5 m from
surface
River Left
Mid depth
sampfe Oepth
0.5 m from
bottom
0.5 m from
surface
Sample rfepfft
Thaiwefl
Mid depth
0.5 m from
bottom
River Right
Mid depth
e depfft
Samp/e
bottom
Sample depth
Depth xx.x m
DO (mfl/L)
Conductivity (yS/cm)
Temperature (
PH
Flag
Phytopiankton Composite Desired Sample (1935-m[ composite excluding preservative!
Sample ID
Composite vol. fmL)
Mymber of Locations SarrsplsJ fQ-3|:
63-um fl^acroEoplankton Composite Sample (1SO-L composite filtration desired)
Sample ID
Volume filtered (L)
Number of Locations Sampled (0-3|:
20-um MicroEoptankton Composite Sample {18-L composite filtration desired)
Sample ID
Volume filtered (L)
Number of Locations Sampled |Q-3):
o::-:e volumes
Flag codes: K=no measurement made. U=suspect measurement;F1, F2. etc=misc flags assigned by field crew.
Explain in comments.
16.
Figure 5-1. Water chemistry and plankton form (front). This version of the form has a
typographic error in the information at the bottom of the form: no sample is
collected at mid-depth if depth at the station is <2 m and >1 m.
104
-------
EMAP-GRE WATER CHEMISTRY AND PLANKTON (back)
9547
SITE ID: GRW04449-
/ / 2 0 0
ANNUAL VISIT
NUMBER:
. D 1 D2
Wafer Chemistry (cont)
Secchi Depth (cm)
Is disi< visible resting on bottom?
FLAG
River Left
Y N
Thalweg River Right
Y N Y N
TyrbWity
Measurement Replicate 1 Replicate 2
Turbidity (KTU's)
Sample Temperatures (C)
FLAG
Replicate 3 Oyplleats 1 Duplicate 2 Duplicate 3
Chlorophyll a Filtration {GFF filterl
Sample TO
Volume filtered (mL)
FLAG
Duplicate Sample ID Duplicate Volym® filtered |mL) FLAG
Geoehemteat Markers (GFF filter)
Sample ID
Duplicate Sample ID
Volume filtered (mL)
Duplicate ¥ol. filtered (mLj
Filter ID FLAG
G
Dupl cate Filter ID FLAG
G
Total Fitter Pair 1
Sample ID
Volume filtered {mL)
Filter Fair ID FLAG
M
Tota Sysp@nded Solids
Sample ID
Duplicate Sample ID
Volume filtered (mL)
Duplicate Vol. filtered (mL)
% tterwbrawe Filter Pair 2
Filter Pair ID FLAG
M
Duplicate Filter Pair ID FLAG
M
FLAG
Flag codes; K = No measurement made, U = Suspect measurement,, F1,F2, etc. = rnisc, ftags assigned by each field crew,
Explain all flags m comment section.
17.
Figure 5-2. Water chemistry and plankton form (back).
105
-------
WC AL
I
visit 1
300213
ZOO PLANKTON
BZ LZ
/
Volume filtered L
visit nymber 1 2
300215
Sample type
GRW04449-
Comp/filtered vol._
Site visit 1 2
I®
PHYTOPLANKTON (PP)
{4% formal In)
/
Composite volume.
visit 1 2
300214
FILTERS
CF GF SS1 SS2
GRW0444i-
I / 200_
Volume filtered
m
ml
visit number 1
300218
Figure 5-3. Labels for water quality and plankton samples. WC = water chemistry composite,
BZ = macrozooplankton, LZ = microzooplankton, AL = alkalinity, CF =
chlorophyll filter, GF = geochemical markers filter, SS1 = total suspended solids
filter pair 1. The bottom number on the labels is a unique sample ID. The bottom
continuation label is used if additional containers are needed for a sample. Not
actual size.
106
-------
SP^* w^iiir*^ ^G^i^S^M^
IS;-?
_
^"" --^ ,,
•2*^
** ""a--^=.i - jp4?te* --i-^^S^'jfflfe?----^"-" '>J**!r 2fe5*!^ri=ti^Sl
ie^S^*" ~J?*ill»'-';^y' ^»*s;^::" -*;=-"'^V-Z.~.nf= --">"^~ :**H--X--
*" V l2«?"3'fe%siiiaEJ^?^^,- 3ss
Figure 5-4. Apparatus for attaching instrument sensors and sample-collecting hoses to the
sounding weight (30 Ibs in this picture). A sounding-weight hanger-bar has been
fabricated to accommodate a Hydrolab DataSonde. The DataSonde is attached
to the hanger bar, facing forward, with stainless hose clamps (covered by tape in
this picture). The hose to the peristaltic pump (small diameter) is taped to the
DataSonde. The hose to the high volume Guzzler pump (large diameter) is
attached to the hanger bar with plastic cable ties. A one-way valve is attached to
the end of the Guzzler hose. When the apparatus is lowered, the instrument
cables and hoses are attached to the winch line with cable ties. Other
configurations are acceptable.
107
-------
Blank Page
108
-------
Section 6
Aquatic Vegetation
Ted R. Angradi2, and E. William Schweiger1
Aquatic vegetation has multiple ecological functions in Great River Ecosystems. Aquatic
plant beds generate dissolved oxygen, stabilize bed sediments, filter suspended sediment, and
immobilize nutrients and toxic substances (Rogers and Theiling 1999). Aquatic plants are an
important food source for waterfowl and other wildlife, and they provide substrate for
invertebrates and habitat for fish (Rogers and Theiling 1999). Submerged aquatic vegetation
(SAV) is sensitive to anthropogenic stressors, including excessive turbidity, sedimentation, flow
modification, and exotic herbivores. Relating SAV community structure, abundance, and
distribution to stressors can provide a biological basis for water quality criteria. For example,
understanding the influence of turbidity on SAV beds could lead to development of light-related
water-quality criteria for Great Rivers (UMRCC 2003). The EMAP-GRE method is adapted from
Yin et al. (2000).
6.1 Aquatic vegetation sampling
Littoral aquatic vegetation is sampled from the main-channel shoreline (MCS) by the
river crew. Plant cover and species occurrence is noted visually, and total and species-specific
density is estimated from samples collected with a vegetation rake. Samples are processed in
the field, although voucher specimens should be retained for identification when necessary.
6.1.1 Sample locations
At each site, two 500-m main channel shoreline (MCS) transects starting at the
intersection of the cross-channel transect and the MCS (Figure 4-1), are located and flagged
out by either the fish- or river-sampling crew, depending on which crew arrives at the site first.
The primary transect is initially flagged at 100-m intervals; intermediate littoral stations at 50-m
National Park Service, 1201 Oakridge Drive, Fort Collins, CO 80525
U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
109
-------
intervals are located and flagged using a handheld GPS or by visual estimation during littoral
sampling (use a different flag color than at the 100 m stations). At every other shoreline station
(A, C, E, G, I, K; Figure 4-2) visual and quantitative methods are used to quantify aquatic
vegetation in a 2 x 5 m littoral plot (Figure 6-1). Plots are slightly offset from the station location
to avoid the effects of disturbance from other littoral sampling (macroinvertebrates, sediment,
and periphyton) and are slightly offset from the wetted margin to avoid the effects of wave
action. For guidance on where to collect aquatic vegetation samples in a dike field, see Figure
10-1. In swift, turbid reaches such as the lower Missouri River, aquatic plant beds will rarely be
encountered in the main channel.
Ralce sample
(2 x 0.36 m)
Shoreline
2 K 5 m
Flow
Subplot
1 m offset
from
wetted
margin
A
3 - 5 m
offset from
V
I
Figure 6-1. Littoral plot for aquatic vegetation sampling. Not to scale.
110
-------
6.1.2 Sampling procedures
In each littoral plot, percent cover by life form and taxa richness are estimated visually.
Habitat characteristics of each plot, including depth, velocity, and substrate are also quantified.
The relative abundance of each submersed taxa is estimated from a 0.72-m2 rake sample
(Figure 6-2) in each of three subplots. The density of vegetation is estimated from the amount
of vegetation caught on the rake. Procedures for aquatic vegetation sampling are described in
Table 6-1.
Figure 6-2. Double-headed sampling rake. Each tine is marked into 5 sections of equal
length. In this sample, total density would be scored as "2" based on rake
fullness.
111
-------
6.1.3 Taxa codes and voucher specimens
Nomenclature and synonymy follow protocols and references established in Yin et al.
(2000). Table 6-2 lists expected species and their codes. Codes are derived from the USDA
Plants database (http://plants.usda.gov/) and Yin et al. (2000). Codes and nomenclature for
species not in Table 6-2 (identified in the field or for voucher specimens in the lab) should be
derived from the Plants database. If genus is known but not species, use the first four letters of
the genus with "?" inserted between the second and third letters or use the genera level USDA
code.
112
-------
Table 6-1. Aquatic vegetation sampling procedures.
1. Fill in the site ID and date on the aquatic vegetation form (Figure 6-3).
2. Go to littoral sample station A (aquatic vegetation is sampled at stations A, C, E, G, I, and K).
Visually establish a 2 m wide x 5 m long littoral plot starting 3 - 5 m upriver from the flagged
station and offset 1 m from the wetted margin (Figure 6-1). If there is no aquatic vegetation in
the plot, go to step 12.
3. If vegetation is present, visually estimate the percent cover, by life-form categories, of
vegetation within the entire 2x5 meter plot. Life-form categories include non-rooted floating-
leaved, rooting floating-leaved, and emergent. Use the following cover codes:
Cover (%)
81-100
61- 80
41- 60
21-40
1-20
None
Cover code
5
4
3
2
1
0
4. Estimate the non-submersed taxa richness at or above the water surface in the 2x5 meter
plot. If taxa are not readily distinguishable, use distinct morphological variants as proxies for
taxa.
5. Prior to rake sampling, evaluate the sampleability of the 2 x 5 meter plot at the station. If the
plot is not sampleable due to safety concerns or excess depth, place a flag in the "Plot
characteristics" section. Explain the flag in comments.
6. If rake sampling is possible, visually divide the plot into three evenly-spaced subplots
(numbered from down river: 1,2,3; Figure 6-1) and collect a rake sample by extending the
rake 2 meters outward from the shoreline at the midpoint of each subplot and lowering the
head to the bottom. Drag the rake along the bottom back to the shoreline for 2 m. Twist the
rake 180 degrees as it is raised out of the water.
7. The density of vegetation is estimated from the amount of vegetation caught on the upper set
of tines on the rake (after it has been twisted 180 degrees) or the "rake fullness". Each tine
should be marked into 5 even vertical segments (requiring 4 horizontal lines on each tine,
Figure 6-2). Vegetation density ranges from 0 to 5. If there is no vegetation on the rake, the
density is 0; if there is vegetation on the rake but the amount does not reach the first horizontal
mark, the density is scored as "1"; if the vegetation on the rake reaches between the first and
second horizontal mark (from the bottom) the vegetation density is scored as "2", etc. If
vegetation is not evenly distributed on the rake, visually distribute the catch across the tines
before assigning a density score.
8. Estimate the total submerged vegetation density of each rake sample (Figure 6-3). Next,
record the taxa code (Table 6-2) and the density code (0 - 5) for each taxa collected on the
rake on the back of the form (Figure 6-4). Record "UNKN" for unknowns (QE codes 3 and 4,
see below). Each taxa may have a density up to 5. In contrast with the surface richness
estimate, submersed taxa are also sampled. Use a continuation form(s) if necessary (Figure 6-
5).
113
-------
Continued
Table 6-1. Aquatic vegetation sampling procedures, continued.
9. For all taxa code designations used, characterize the quality of the taxonomic evaluation in the
"QE" field using the following scale:
Taxonomic evaluation QE code
Species code matches definition in Table 6-3 1
Genus certain 2
Genus and species suspected 3
Unknown 4
10. Collect specimens for taxa with a QE code other than 1. Place each specimen in a plastic bag
with an Aquatic Plant Specimen Tag (one taxa per bag; Figure 6-6). Record the tag number on
the form. Attempt to collect at least two specimens with roots and flowers (if possible).
Duplicate specimens may be placed in the same bag and have the same specimen tag
number. Place bagged specimens in a larger bag. Place an Aquatic Plant Specimen Label
(Figure 6-6) on the outer bag and cover with clear tape. Place the specimens in a cooler on
ice.
11. Based on the sediment brought up with the rake samples or visually from the shoreline, classify
the dominant substrate in the 2 x 5 m plot and note the presence/absence of detritus (Figure 6-
3).
12. Assign a depth category to the 2 x 5 m plot based on the midpoint of the rake samples.
Estimate water velocity in the plot using the qualitative scale on the form (Figure 6-3).
13. Search for aquatic plants along the shoreline between littoral plots. If present, check the
appropriate box (Figure 6-3) and list taxa codes in comments.
14. Repeat steps 2-13 at the remaining five shoreline stations.
15. Place each bagged plant specimen in a larger bag. Place an Aquatic Plant Specimen Label
(Figure 6-6) on the outer bag and coverwith cleartape. Record the sample ID from the label
on the form (Figure 6-4). Place the specimens in a cooler on ice.
114
-------
Table 6-2. Taxa codes for aquatic plants. Life form codes: A = filamentous algae; N = non-rooted floating-leaved; E =
emergent; F=Floating; S = submersed; U = not applicable.
Code
ACSA2
ALPL
AM7AR
AMTU
AMTR
AMCO
AMCOP
AP70C
ASIN
AZCA
AZME
AZ70L
BARO
BIAR
BOCY
CASES
CA7RE
CEOC2
CEDE4
CHSE4
CH7AR
CODI5
CYER2
CY7PE
CYST
DEVE
DEIL
DI7GI
DIVI5
DUAR3
ECCR
ECES
ECMU2
ECWA
ECPR
EL7EO
ELCA7
ELVIS
EQ7UI
ERFR
ERPE
ER7IG
ALGA
Life form
E
E
E
E
E
E
E
E
E
N
N
N
E
E
E
E
E
E
S
E
S
E
E
E
E
E
E
E
E
E
E
E
E
E
E
E
S
E
E
E
E
E
A
Scientific name
Acer saccharinum
Altissima plantago-aquatica
Amaranthus spp.
Amaranthus tuberculatus
Ambrosia trifida
Ammannia coccinea
Ammannia coccinea purpurea
Apocynum spp.
Asclepias incarnata
Azolla caroSniana
Azolla mexicana
Azolla spp.
Bacopa rotundifolia
Bidens aristosa
Boehmeria cylindrica
Calystegia sepium sepium
Carex spp.
Cephalanthus occidentalis
Ceratophyllum demersum
Chamaesyce serpens
Chara spp.
Commelina diffusa
Cy perns erythrorhizos
Cyperus spp.
Cyperusstr'gosus
Decodon vertidllatus
Desmanthus illinoiensis
Digitaria spp.
Diospyrus virginiana
Dulichium arundinaceum
Echinochloa crus-galli
Echinochloa esculenta
Echinochloa muricata
Echinochloa walteri
Eclipta prostrata
Eleocharis spp.
Elodea canadensis
Elymus virginicus
Equisetum spp.
Eragrostis frankii
Eragrostis pectinacea
Erigeron spp.
filamentous algae
Common name
silver maple
American waterplantain
pigweed
roughfru it amaranth
great ragweed
valley redstem
valley redstem
dogbane
swamp milkweed
Carolina mosquitofern
Mexican mosquitofern
mosquitofern
disk waterhyssop
bearded beggarticks
false nettle
false hedge bindweed
sedge
common buttonbush
coon's tail
matted sandmat
chara
climbing dayfiower
redrootflatsedge
flatsedge
straw-colored flatsedge
swamp loosestrife
prairie bundlefiower
crabgrass
common persimmon
threeway sedge
barnyardgrass
Japanese millet
rough barnyardgrass
coast cockspur grass
false daisy
spikerush
Canadian waterweed
Virginia wildrye
horsetail
sandbar lovegrass
tufted lovegrass
fleabane
filamentous algae
Code
NELU
NULU
NYTU
PAFL5
PESE6
PHAR3
PHAU7
PH7RA
PHLA3
PH7YS
POPR
P07A
POACEA
POAM8
POAME
POHY2
POL013
POPE2
POPU5
P07LY
POC014
PODE3
POAL8
POCR3
POEP2
POF03
NLPW
PON02
POPU7
PORI2
POZO
RAL02
RA7NU
RATR
RIFL4
RINA2
RONA2
RUCR
RU7ME
SACU
SALA2
SARI
SA7GI
Life form
F
F
F
E
E
E
E
E
E
E
E
E
E
E
E
E
E
E
E
E
E
E
S
S
S
S
S
S
S
S
S
S
S
S
N
N
E
E
E
E
E
E
E
Scientific name
Nelumbo lutea
Nuphar variegata
Nymphaea odorata tuberosa
Paspalum fluitans
Penthorum sedoides
Phalaris aruninaceae
Phragmites australis
Phragmites spp.
Phyla lanceolata
Physostegia spp.
Poa pratensis
Poa spp.
Poaceae
Polygonum amphibium
Polygonum amphibium v. emersum
Polygonum hydropeperiodes
Polygonum pensyh/anicum
Polygonum pensylvanicum
Polygonum punctatum
Polygonum spp.
Pontederia cordata
Populus deltoides
Potamogeton alpinus
Potamogeton crispus
Potamogeton epihydrus
Potamogeton folbsus
Potamogeton foliosus/pusillus
Potamogeton nodosus
Potamogeton pusilus
Potamogeton richardsonii
Potamogeton zosteriformis
Ranunculus longirostris
Ranunculus spp.
Ranunculus trichophylus
Riccia fluitans
Ricc'ocarposnatans
Rorippa nasturtium-aquaticum
Rumex crispus
Rumex spp.
Sagittaria cuneata
Sagittaria latifolia
Sagittaria rigida
Sagittaria spp.
Common name
American lotus
yellow pond lily
white waterlily
horsetail paspalum
ditch stonecrop
reed canary grass
common reed
reed
lanceleaffogfruit
dragonhead
Kentucky bluegrass
bluegrass
grass family
water knotweed
longrootsmartweed
swamp smartweed
Pennsyh/ania smartweed
Pennsyh/ania smartweed
dotted smartweed
smartweed
pickerelweed
eastern cottonwood
alpine pondweed
curly pondweed
ribbonleaf pondweed
leafy pondweed
narrow-leaved pondweeds
longleaf pondweed
small pondweed
Richardson's pondweed
flatstem pondweed
longbeak buttercup
buttercup
thread leaf crowfoot
slender liverwort
liverwort
watercress
curly dock
dock
arumleafarrowhead
broadleaf arrowhead
stiff arrowhead
arrowhead
Continued
115
-------
Table 6-2. Taxa codes for aquatic plants, continued.
uoae
FOAC
FRPE
GAOBO
GA7LI
ZODU
HILA2
IPLA
JUEF
LEOR
LE7ER
LEMI3
LETR
LEMNA
LEFI
LEFA
LEPA3
LUDE4
LUPE5
LU7DW
LYSA2
MYSI
MYSP2
MY7RI
NAFL
NAGR
NAGU
NAMI
NOSMPL
UNKN
Lire Torm
E
E
E
E
S
E
E
E
E
E
N
N
N
E
E
E
F
E
F
E
S
S
S
S
S
S
S
u
u
scientific Name
Forestiera acuminata
Fraxinus pennsylvanica
Galium obtusum obtusum
Galium spp.
Heterantheradubia
Hibiscus laevis
Ipomoea lacunosa
Juncuseffusus
Leersia oryzoides
Leersia spp.
Lemna minor
Lemna trisulca
Lemnaceae
Leptochloa filiformis
Leptochloa fusca fascicularis
Leptochloa panicoides
Ludwigia decurrens
Ludwigia peploides
Ludwigia spp.
Lythrum salicaria
Myriophyllum sibiricum
Myriophyllum spicatum
Myriophyllum spp.
Najas flexilis
Najas gracilSma
Najas guadalupensis
Najas minor
no sample
Unknown
uommon Name
eastern swampprivet
green ash
bluntleaf bed straw
bedstraw
water stargrass
halberdleaf rosemallow
whitestar
common rush
rice cutgrass
cu (grass
small duckweed
star duckweed
duckweed family
muronatesprangletop
bearded sprangletop
Amazon sprangletop
wingleafprimrosewillow
floating primrosewillow
primrosewillow
purple loosestrife
northern watermilfoil
Eurasian watermilfoil
watermilfoil
nodding waternymph
slender waternymph
southern waternymph
brittle waternymph
uoae
SAEX
SANI
SA7LI
SCFL
SCVA
SC7IR
SE7NE
SEVI4
SIAN
SM7IL
SPEU
SPPO
POPE6
SYLAL7
TYAN
TYLA
TY7PH
ULAM
URDI
UR7TI
UTMA
VAAM3
VEHA2
VICI2
VI7TI
WOCO
W07LF
XAST
ZAPA
ZIAQ
Lire Torm
E
E
E
E
E
E
E
E
E
E
E
N
S
E
E
E
E
E
E
E
S
S
E
E
E
N
E
E
S
E
scienuTic Name
Salixexigua
Salix nigra
Salixspp.
Schoenoplectus fluviatilis
Schoenoplectus tabernaemontani
Scirpus spp.
Senecio spp.
Setaria virdis
Sicyos angulatus
Smilaxspp.
Sparganium eurycarpum
Spirodela polyrrhiza
Stuckenia pecfnatus
Symphyotrichum lateriflorum v. alateriflorum
Typha angustifolia
Typha latifolia
Typha spp.
Ulmus americana
Urtica dioica
Urtica spp.
Utricularia macrorhiza
Vallisneria americana
Verbena hastata
Vitis cinema
Vitis spp.
Wolffa Columbians
Wolffa spp.
Xanthium strumarium
Zannichellia palustris
Zizania aaua ties
uommon Name
sandbarwillow
blackwillow
willow
river bulrush
softstem bulrush
bulrush
ragwort
green bristlegrass
oneseed burrcucumber
greenbrier
giant burreed
big duckweed
sago pondweed
calico aster
narrowleaf cattail
common cattail
cattail
American elm
stinging nettle
nettle
common bladderwort
wild celery
swamp verbena
graybark grape
grapevine
Columbian watermeal
watermeal
rough cocklebur
horned pondweed
wild rice
116
-------
6.2 QA considerations for aquatic vegetation sampling
Table 6-3. QA considerations for aquatic vegetation sampling.
Attempt to standardize the rake sampling among all substrate and vegetation conditions
encountered among sites.
Do not attempt to sample where conditions prevent a good rake sample (e.g., depth >2 m or in
areas of fast current).
Be sure to fill out the header information on the continuation forms properly so they can be
associated with the main forms
6.3 Equipment and supplies for sampling aquatic vegetation
Generic supplies required for all EMAP-GRE field sampling are listed in Table 2-5.
Table 6-4. Checklist of equipment and supplies for aquatic vegetation sampling.
Qty
2
1
1 set
20
20
1 set
1 set
Item
Double headed vegetation sampling rake with 3-m long handle. Buy 2 steel garden
rakes, cut the head off one and weld it to the head of the other at the same angle
from the handle. The rake head should be 36 cm wide and have about 14 5-cm-
long tines on each side. Add a rope extension to the handle and mark the handle in
1 0-cm increments. Mark the tines into 5 equal vertical increments by scoring the
tines or with paint. See Figure 6-2 .
Aquatic vegetation species list (Table 6-3)
Aquatic plant identification guides (not provided by EPA)
1 -gallon self-sealing plastic bags
2-gallon self-sealing plastic bags
Aquatic vegetation forms
Labels (voucher tags, labels)
117
-------
6.4 Literature cited
Rogers, S. and C.Theiling 1999. Submersed aquatic vegetation. In Ecological status and trends
of the Upper Mississippi River System 1998: A Report of the Long Term Resource
Monitoring Program. U.S. Geological Survey, Upper Midwest Environmental Sciences
Center, La Crosse, Wl. April 1999. LTRMP99-T001.
Upper Mississippi River Conservation Committee, Water Quality Technical Section (UMRCC).
2003. Proposed water quality criteria necessary to sustain submersed aquatic
vegetation in the Upper Mississippi River. Rock Island, IL. 6p .
Yin, Y., J.S. Winkelman, and H.A. Langrehr. 2000. Long term resource monitoring program
procedures: aquatic vegetation monitoring. U.S. Geological Survey, Upper Midwest
Environmental Sciences Center, 2630 Fanta Reed Road, Lacrosse, Wl 54603
http://www.umesc.usgs.gov/data_library/vegetation/vegetation_page.html
ftp://ftp.umesc.er.usgs.gov/pub/media_archives/documents/reports/1995/95p00207.pdf
118
-------
EMAP-GRE AQUATIC VEGETATION FORM (front)
Reviewed
by (Initial*):
Draft
SITE ID: GRW04449-
OATE;
1200
ANNUAL VISIT
NUMBER:
D1 D2
Cover and Taxa Richness in entire 2
Station
A
C
E
G
i
K
Station
A
C
E
G
I
K
Non-rooted Rooted
floating floating leaved
543210 54321
543210 54321
543210 54321
543210 54321
543210 54321
543210 54321
0
0
0
0
0
0
x 5m littoral plot
Emergent
5432
5432
5432
5432
5432
5432
1 0
1 0
1 0
1 0
1 0
1 0
Richness
Flag
C
g
4
2
I
0
over Codes:
= 81 - 100%
= 61 - 80%
= 41 - 60%
= 21 - 40%
= 1 - 20%
= 0%
Plot Characteristics in entire 2 x 5m littoral plot
Veiodtv Dominant
Code Depth Coele Substrate
Code
210 210
210 210
210 210
210 210
210 210
210 210
Detritus
presence'
absence
Y
Y
Y
Y
Y
Y
N
N
N
N
N
N
Flag
Code Keys
and Descriptions
X8 = BOULDER (1000 TO 4008 mm) METERSTICK TO CAR
SS z SM.BOULOSR (250 TO 1000mm! BASKETBALL TO METiRSTICK
CB = COBBLE (64 TO 250 iron) TENNIS BALL TO BASKETBALL
OC = COARSE GRAVEL (16 TO 64 mm) MARBLE TO TENNISBALL
OF = ORAViL (2 TO 84 mm) LADYiUO TO MARBLI
SA = SAND {0.0«TO2rnm) GRITTY - UP TO LAOYBUO SIZE
FN = SILT,' CLAY /MUCK NOT GRITTY
HP = BARBPAN FIRM, CONSOLIDATED FINE
SUBSTRATE
WO = WOOD ANY SIZE WOOD
OT=OTHER FLAG AND COMMENT
VELOCITY: 2 = > Im/s, 1 = < Im/s and >0, 0 = amis.
DEPTH: 2 = >1m, 1 => 0,5m and < 1m. 0 = < 0.5m.
Total Submersed Vegetation Density (rake fullness) in Subplots
A C
1 543210 543210
2 543210 543210
3 543210 543210
* Outside i — i i — i
of plots LJ LJ
Station
E
543210
543210
543210
Flag
n
G
5432
5432
5432
n
1 0
1 0
1 0
1
5 4
5 4
5 4
3 2 1
3 2 1
3 2 1
n
0
0
0
K
54321
54321
54321
D
0
0
0
' Presence/absence of vegetation outside of plot, if present, check box and list taxa codes in comments.
Flag
COMMENTS
Fiag eodes: K = Mo measurement made, U = Syspect measurement, F1,F2( etc, = misc. flags assigned by each field crew,
Explain all flags in comment section, -J g
Figure 6-3. Aquatic vegetation form (front).
119
-------
•
SITE
rm
Draft
D; GRW0444S-
EMAP-GRE AQUATIC VEGETATION FORM (back)
I » « n n ANNUAL VISIT r
DATE;, . ,' . , ,» ,Z ,° ,0 , , NUMBER: L
Reviewed
by (Initials)
11 D2 PAG
•
E: 1 OF
Rake of Submersed Taxa
Sample ID
Density of IndivMya! submersed taia from rake samples (rake fyllness)
Sta,
?$$;» Cewte
J 1 I ! t !
Rake Sample '
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
Rake Sample I
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
Rake Sample 3
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
QE
Cede
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
Sp^dmen
Ti|*
Flag
®E Codes: 4 = Unknown, 3 = Genus and species suspected, 2 = Genus certain, 1 = Code match. Collect one specimen if CiE Code rtot = 1 ,
COMMENTS
^^^ Flag codes: K = No measurement made, U = Suspect measurement,, F1,F2, etc. = misc. flags assigned by ^^^
HI each field crew. Explain all flags in comment section. ,« g HI
Figure 6-4. Aquatic vegetation form (back).
120
-------
•
SITE
*J
Draft
ID; 0RW0444S-
EMAP-GRE AQUATIC VEGETATION FORM (cent)
, / 9 ft n ANNUAL VISIT
DATE: / / i U 0 NUMBiR:
Reviewed
by (Initials)
m 02 PAG
•
E: OF
Rake Sample of Submersed Taxa
Sample ID
Density of individual submersed taxa from rake samples (rate fullness)
Stt.
T*)t* Code
R»te Sample 1
543210
S 4 3 2 1 0
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
Rtto Sample 2
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
Rake Sample 3
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
543210
QE
Code
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
4321
Specimen
Tafli
Fla§
QE Codes: 4 = Unknown, 3 = Genus and species suspected, 2 = Genus certain. 1 = Code match. Collect one specimen if QE Code not = 1 .
COMMENTS
^^ Flag codes: K = No measurement made, U = Suspect measurement., F1,F2, etc. = misc. flags assigned by ^^
^^1 each field crew. Explain all flags in comment section. «« ^^1
Figure 6-5. Aquatic vegetation continuation form (one sided form).
121
-------
PLANT
(AP)
GRW0444S-
_!_)
number 1 2
ID
(AP)
number 1 2
0000177
Figure 6-6. Labels and tags for aquatic plant specimens. Plant specimen tags are placed in
the bags containing individual specimens. Specimen labels are placed on the
larger outer bag holding the smaller bags. Use continuation label at upper right if
extra outer bags are needed. Not actual size.
122
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Section 7
Riparian Habitat
E. William Schweiger1 and Ted R. Angradi2
Interactions among aquatic and riparian ecosystem components are important in Great
River ecosystem functioning and condition. Riparian ecosystems contribute to and moderate
the flux of materials and energy between terrestrial and aquatic habitats within GREs. Because
shoreline and riparian characteristics (vegetation, stability, human modifications) can affect
channel form, water velocity, and substrate, they influence ecosystem condition at multiple
spatial scales. This section includes procedures for evaluating riparian vegetation, land cover,
bank morphology, and human influences at each site. Methods in this section are adapted, in
part, from Peck et al. (Unpublished drafts) and Ringold et al. (2001).
7.1 Sample locations
Data collection for the main channel shoreline (MCS) and riparian habitat is conducted
by the river-sampling crew at shoreline stations and riparian plots along the primary 500 m MCS
transect (Figure 7-1; see also Figure 4-2). The MCS is defined in Section 4.2.1. MCS bank
sample stations are located at transects A, E, and K, located 0, 200, and 500 m upriver from
the start of the primary MCS transect (Figure 7-1). Shoreline and macrohabitat type is
evaluated for each of the five intervening 100-m shoreline segments.
7.2 Riparian plots
7.2.1 Location on the main channel shoreline
Riparian plots are established at one of two positions in relation to the MCS (see Section
4.2.1): 1) at or just above the bankfull elevation, or 2) at the main-channel wetted margin. If
there are no limitations to riparian plot access, the preferred location is at or just above the
National Park Service, 1201 Oakridge Drive, Fort Collins, CO 80525
U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
123
-------
bankfull elevation on the MCS. Access is considered limited when reaching the MCS transect
requires more than 10 minutes of walking, relocation of the boat (e.g., into or across a
secondary channel), or crossing posted or other obviously private property with controlled
access. The amount of effort expended to locate riparian plots at the bankfull level is at the
discretion of the crew leader. No specific procedures are provided here for obtaining landowner
permission to access riparian areas on posted or non-posted private property. It is also up to
the crew leader to decide if the limited activities in riparian plots (15-30 minutes per station)
warrant solicitation of landowner permission. Physical access may often be limited on inside
bends where a point bar separates the active channel from a distant high bank. If access to a
plot is incomplete, it is acceptable to only sample a subset of a plot (e.g., just subplot 1,
described below) or to collect measurements remotely (i.e., by viewing vegetation from the
boat). All data collected using non-standard methods should be flagged. An alternate riparian
location should not be substituted for a plot that is not sampled.
The first step in locating the riparian plots is to establish the approximate bankfull
elevation on the MCS. Estimating this elevation during base flow is somewhat subjective. Best
judgement should be used in cases where the bankfull elevation on the MCS is difficult to
locate. If possible, document the bankfull level with a description and site photos (see Section
4). Bankfull elevation corresponds to a flow stage with a recurrence interval of 1 to 2 years
which generally does not inundate the floodplain. Bankfull elevations often correspond to a
"greenline," or the line of first perennial vegetation above the wetted margin of the main
channel. The bankfull elevation may also be detected from the presence of debris caught on
overhanging vegetation. However, subsequent to large floods, this material may be well above
the bankfull level. For braided streams where perennial vegetation may be established on bars
between the channels or on islands, the bankfull elevation or greenline of interest is on the
MCS. On steep banks where the bankfull elevation or greenline is part way up the slope, the
edge of the riparian plots will be located at the top of the bank. If the MCS is gradually sloped,
the bankfull elevation may be more difficult to locate. Often there is a small berm on the MCS at
the bankfull elevation, with subtle changes in vegetation. If MCS bank station A, E, or K is on a
jetty or other artificial structure projecting into the channel, the riparian plot should be positioned
landward of the jetty on the closest point on the non-artificial MCS.
If access to a bankfull elevation location is physically limited (e.g., at an inside bend
124
-------
where the high bank is distant from the river) but is not posted or otherwise restricted, riparian
plots should be established just above the wetted margin of the main channel along the MCS.
Main-
channel
shoreline
transect
segments
Shoreline
station
Riparian
Plot
(30 x 30 m)
Cross-channel
Transect
100
(10x30m)
A .....
E
Figure 7-1. Sample layout for riparian habitat sampling.
125
-------
7.2.2 Riparian plots
Riparian plots are centered on the corresponding MCS bank station (station A, E, or K;
Figure 7-1). The plots extend landward 30 m perpendicular to the main channel and 15 meters
upriver and downriver (Figure 7-2). Each plot is divided into three, 30 x 10 m subplots. Plot
borders are established by stretching a tape 30 m into the riparian corridor orthogonal to the
MCS (flags may be placed at 10 and 20 m). The upriver and downriver extent of each subplot
are estimated by eye. The three subplots are numbered 1, 2, and 3 from the MCS edge
landward.
River bank
10 x 30 m
subplots
Shoreline
staion
30m
Wetted
margin
GPS
Locations
Main-channel
Figure 7-2. Closeup of 30 x 30 m riparian plot.
126
-------
7.3 Bank and channel width measurements
MCS bank angle, height, and width and wetted channel width are determined at three
shoreline stations (A, E, and K; Figure 7-1). Alternate MCS bank stations should not be
substituted for a non-sampled station. MCS bank angles are estimated by viewing the bank
from the wetted channel margin. Bankfull height on the MCS is measured with, in order of
preference, a laser hypsometer, a surveyor's rod, or by visual estimation. If the MCS is
established on a border fill or other similar habitat type, the bank angles and heights may be
low. Channel widths for the main channel are measured from the MCS with a laser rangefinder
sited across the channel to the wetted margin on the opposite bank. Table 7-3 describes
procedures for bank measurements.
Macrohabitat and shoreline type are determined for the 100 m shoreline segments
between MCS bank stations A, C, E, G, I, and K. Shoreline macrohabitat includes continuous
and discrete habitat types (Table 7-1). Continuous macrohabitats (e.g., inside and outside
bends) characterize every river shoreline and may lack a definitive beginning or end. Discrete
macrohabitats (e.g., secondary channels, tributary mouths) are not necessarily found along
every river shoreline. Macrohabitat types may overlap along a MCS. Shoreline type includes
several natural and artificial categories (Table 7-2). Shoreline types are defined as mutually
exclusive. Procedures for estimating the percentage of each shoreline type and macrohabitat
type between each bank station are described in Table 7-3.
127
-------
Table 7-1. Macrohabitat types (adapted from Sappington et al. 1998).
Macrohabitat
Description
Main channel crossover
Outside bend
Inside bend
Pool
Tributary mouth
Connected secondary channel
Unconnected secondary channel
Other
The inflection point of the thalweg. Longitudinal length of the
crossover area generally does not exceed 1.5-2 times the stream
width. Shorelines are often relatively straight in crossover reaches.
The concave side of a river bend often characterized by rip rap or
eroding cut banks
The convex side of a river bend; often characterized by sand bars
Low velocity area upriver of navigation locks or immediately
downriver from dikes or jetties
Mouth of perennial tributary < 10 m across
Lotic channel other than the main channel < 5 m across
Channel < 5 m across, blocked at upriver end by a sand bar
artificial closure, or otherwise; lentic
Any other type of macrohabitat. Describe on form.
Table 7- 2. Shoreline types.
Shoreline type
Description
Stable bank
Aggrading bank
Eroding bank
Blanket-rock revetment (riprap)
Other revetment type
Other
Naturally stabilized due to vegetation or substratum
characteristics
Collecting sediment or extending into the channel; includes point
bars and border fills
Unstable, actively degrading banks
Stabilized by gravel, cobble or boulder-sized material placed at
the river margin
Includes car bodies, cement slabs, trash. Describe on form.
Any other type of shoreline. Describe on form.
128
-------
Table 7-3. Procedure for bank and shoreline measurements.
1. Proceed to shoreline station A (Figure 7-1). Fill in the site ID and date on the bank and
shoreline macrohabitat form (Figure 7-3).
2. Record the coordinates of the shoreline station (Figure 7-2) using a hand-held GPS (applies to
A, C, E, G, I, K).
3. At stations A, E, and K, stand at the edge of the wetted width on the MCS and site across the
channel with a laser range finder and take a reading from an object judged to be at or near the
wetted edge on the opposite bank. If the opposite bank of the main channel is obscured by an
island, take the best reading possible and the flag the data.
4. At stations A, E, and K, estimate bankfull height on the MCS using one of the following
methods: 1) use a laser hypsometer; 2) hold the surveyor's rod vertical, with its base at the
water's edge and estimate (by eye) the height of the bankfull shoreline above the present water
level; or 3) make a visual estimate. If the MCS is very gradual, a meter stick may be used.
Depending on the hypsometer model, the minimum horizontal distance for a bank height
reading may vary (minimum distance is 5 m for Opti-Logic laser hypsometers). Note the
measurement method in comments.
5. At stations A, E, and K measure or estimate the bank width which is the horizontal distance
(not slope distance) from the wetted edge to the bankfull level.
6. At stations A, E, and K, visually estimate the maximum bank angle from the wetted edge to the
bankfull level. Ignore steep breaks in the slope that are <1 m long. Use the classes indicated
on the bank and shoreline macrohabitat form (Figure 7-3).
7. At stations A, E, and K, additional riparian measurements are done before leaving the station
(described later in this section).
8. Proceed to the next shoreline station. On the way, note the shoreline type (Table 7-2 ) and
macrohabitat type (Table 7-1) in the 100-m MCS segment between stations. Estimate the
percent of the segment in each shoreline and macrohabitat type. Ignore types < 5 m in length.
The macrohabitat type may include discrete features (e.g., a tributary mouth) nested within
continuous features (e.g., inside bend shorelines) and macrohabitat types may sum to more
than 100%. Shoreline types are mutually exclusive and must sum to 100%.
9. Repeat steps 2 - 8 as appropriate at shoreline stations C, E, G, I, K.
129
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7.4 Riparian measurements
EMAP-GRE riparian measurements include land cover/use classification, visual
estimation of vegetative cover, canopy density measurements, presence of invasive plant
species, legacy tree characteristics, and human activity and disturbance at the site.
7.4.1 Land cover classification
Land cover in each riparian subplot is classified by codes in Table 7-4 following
procedures in Table 7-5. If a subplot appears atypical from the surrounding area, classification
data for that subplot should be flagged (for example, if a road passes through a subplot). For
riparian plots or subplots on posted land, an attempt should be made at classification without
entering the property.
Table 7- 4. Land cover/land use classification codes.
Code
Description
B Bare, unvegetated, non-forest, non-agricultural, undeveloped (less than 10% vegetative
cover or no obvious vegetation; rock, sand, bare soil, etc.)
D Developed/urban (pavement, buildings, residential yards, industrial/commercial areas,
quarries, highways, etc.)
A Intensive agriculture (row crop, cereal grain, grass seed, nursery, vineyard, orchard,
Christmas trees, etc). Includes bare plowed fields and irrigated pasture.
X Xeric (upland) shrubs or herbaceous. Includes non-irrigated pasture and hayfields.
HM Mesic (riparian) herbaceous (grasses, forbs, etc.). Includes wet prairies or meadows,
sedges, rushes, etc.
SM Mesic (riparian) woody shrubs
F Forested cover (> 10% tree [> 5 m tall] coverage)
130
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Table 7-5. Procedures for classifying riparian subplots.
1. Fill in site ID and date on both sides of the riparian classification and human influence form
(Figure 7-4). Climb the MCS bank (if necessary) and establish the midpoint of the riverside
edge of subplot 1 (Figure 7-2). The 30 x 30 meter plot will extend 15m upriver and downriver
and 30 meters landward from this point.
2. If the land does not appear to be posted or if access is not otherwise restricted, stretch a tape
10 m perpendicular to the bank line landward, establishing the width of the first 10 x 30 m
rectangular subplot (Figure 7-2). The upriver and downriver dimensions (15 m) are estimated
by eye. If there is no access to any or all of the subplots, if it is unsafe, or the vegetation is
impenetrable, all riparian estimates are flagged and done from the bank or closest location
possible.
3. From within riparian subplot 1, assign only one land cover/land use code (Table 7-4) for the
subplot. If more than one code applies, use the code for the dominant land cover/land use and
flag the data.
4. Repeat the process in each subplot. Acquire and record the coordinates of the land edge of
riparian subplot 2 using a handheld GPS (Figure 7-2).
7.4.2 Canopy density
Canopy density is estimated at the river's edge and the land-side edge of subplot 2
(Figure 7-2) using a Convex Spherical Densiometer (Lemmon, 1957). The densiometer must be
taped exactly as shown in Figure 7-6 to limit the number of square grid intersections to 17.
Densiometer readings can range from 0 (no canopy cover) to 17 (maximum canopy cover).
Four measurements (upriver, downriver, landward, and toward the river) are obtained at each
designated point. During measurements, hold the densiometer level approximately 1 m above
the ground. The procedure is described in Table 7-6.
7.4.3 Riparian vegetation structure
Riparian vegetation areal cover is estimated by eye in three vertical layers in each
subplot using methods adapted from Kaufmann and Robison (1998). Areal cover is analogous
to the amount of shadow cast by a particular layer and just that layer when the sun is directly
overhead. The type of woody vegetation (deciduous, coniferous, mixed, or none) is determined
in each of the layers. Consider a layer "mixed" if deciduous and coniferous trees each comprise
more than 25% of the areal coverage. The maximum cover in each layer is 100%, so areal
131
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cover for the combined three layers could sum to 300%. Procedures for estimation of
vegetation coverage are described in Table 7-6.
Table 7-6. Procedure for characterizing riparian vegetation structure.
1. Canopy density. At river's edge (adjacent to MCS station A) hold the densiometer 1 m above
the ground facing landward, away from the main channel. Position the densiometer so your
face is just below the apex of the taped "V" (Figure 7-6). Count the number of intersections (0-
17) that are covered by a leaf, stem, branch, or the bank. In heavy cover, it is usually easier to
count the open intersections and subtract that number from 17. Record the count on the
riparian classification and human influence form (Figure 7-4).
2. Repeat step 1 three more times: facing upriver, downriver, and toward the river.
3. Riparian vegetation structure. From within subplot 1, conceptually divide the riparian
vegetation in into three layers: a canopy layer (>5 m [16 ft] tall), an understory layer (0.5 - 5 m
tall), and a ground cover layer (<0.5 tall). Several vegetation types (e.g., grasses or woody
shrubs) can potentially occur in more than one layer.
4. Record the type of woody vegetation (deciduous, coniferous, mixed, or none) for each layer.
Consider the layer "mixed" if both deciduous and coniferous trees comprise more than 25% of
the areal coverage.
5. Estimate the cover for each category (e.g., big trees, small trees) in each canopy layer (0 =
absent, 1 = sparse [<10%], 2 = moderate [10 - 40%], 3 = heavy [40 - 75%], 4 = very heavy
[>75%]). Estimate areal cover as the amount of shadow that would be cast by a particular layer
alone if the sun were directly overhead.
6. Estimate the percent of bare ground for the ground cover layer.
7. Repeat Steps 3 - 6 for subplots 2 and 3.
8. At the land-side edge of subplot 2 repeat steps 1 and 2 and record the GPS coordinates.
9. Repeat steps 1 - 8 at each riparian plot.
132
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7.4.4 Human influence
For each riparian subplot, the presence/absence and the proximity of human influences
(i.e., activities, disturbance) are recorded. Each human influence is recorded as not present at
the site; as present within subplot 1, 2, or 3; present between the river and subplot 1; present in
the river; present on the opposite bank; or present on the target side of the river outside of
riparian plots. Procedures for characterization of human activities and disturbances in the river
and in the riparian area are described in Table 7-7.
Table 7- 7. Procedure for tallying human influence.
1. From the edge of subplot 1 in riparian plot A, record any human influence categories between
the subplot and the river.
2. Examine the other subplots for human influence. Record any human influence present in any of
the subplots (Figure 7-4). Other influences not included in the list on the form are likely to be
present.
3. From the corners of the riparian plot, search for human influences (on the river ecosystem)
visible outside the plot and not already recorded from the subplots or from between subplot 1
and the river. Human influence outside the subplots may be in the river, on the opposite side of
the river, or on the target side of the river. It is not necessary to circle "N" for every human
influence not present.
4. Repeat Steps 1 through 3 for riparian plots E and K.
7.4.5 Riparian "legacy" trees
Legacy tree data contribute to the assessment of "old growth" characteristics of riparian
vegetation and can indicate historic riparian conditions and the potential for riparian tree growth.
Legacy trees are defined as the largest tree of any species alive or dead in the riparian plot or
within 100 m of the riverside edge of the riparian plot. The legacy tree may be on the floodplain
or on the upland beyond the margin of the floodplain if the floodplain is <100 m wide.
Procedures for identifying and recording the attributes of legacy trees are described in
Table 7-8.
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7.4.6 Invasive plant species
Invasive plant species may alter the structure and function of the riparian ecosystems
which they invade. Stressed aquatic ecosystems are vulnerable to invasion by alien riparian
plant species. Target invasive plant species are given in Table 7-9. These taxa were selected
based on their degree of invasiveness in riparian systems, their capacity to alter ecosystem
function, and ease of identification. Expected distributions, ID codes, and nomenclature are
from the USDA Plants database (http://plants.usda.gov). Do not include species seen in
another subplot as "outside of subplot" observations. Invasive species other than those in Table
7-9 may be noted in a comment. The procedures for tallying invasive plant species are
described in Table 7-8.
Suspected invasive plan specimens may be informally collected for later identification or
confirmation by a botanist. If necessary, these occurrences should be recorded on the field
form before they are sent to the EMAP data manager. There are no formal invasive plant
vouchering requirements for EMAP-GRE.
134
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Table 7-8. Procedures for identifying riparian legacy trees and invasive plant species.
1. Legacy tree. Search the three subplots and an area extending 70 m landward from subplot 3
(Figure 7-1). Identify the largest tree of any species, alive or dead (including snags), in the
search area (one legacy tree for each riparian plot). If only small trees are present, select the
largest example.
2. Estimate the DBH and height of the legacy tree by category. Classify the tree as (D)eciduous,
(C)oniferous, or (N)o trees. Estimate the distance from the tree to the wetted edge of the river.
Record the information on the riparian legacy tree and invasive plants form (Figure 7-5).
3. Assign a taxonomic category to the tree from the list on the form (Figure 7-5).
4. Invasive plant species. Search each subplot for the presence of targeted invasive plant
species (Table 7-9). Check the appropriate box on the form (Figure 7-5).
5. Note the presence of any of the target species in the riparian areas outside riparian subplots.
6. Repeat steps 1-6 at each riparian plot.
7. If any other invasive plant species (not listed in Table 7-9) are observed at the site, note these
in a comment.
135
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Table 7-9. Target invasive plant species.
Common name
Canada thistle
musk thistle
leafy spurge
Russian olive
salt cedar
buckthorn
reed canary grass
giant reed
cheatgrass
teasel, Fuller's teasel
common, lesser
burdock
Japanese knotweed
mile-a-minute
garlic mustard
purple loosestrife
knapweed
whitetop
Genus species
Cirsium arvense
Carduus nutans
Euphorbia esula
Elaeagnus angustifolia
Tamarix spp.
Rhamnus spp.
Phalaris arundinacea
Arundo donax
Bromus tectorum
Dipsacus fullonum
Arctium minus
Polygonum cuspidatum
Polygonum perfoliatum
Alliaria petiolata
Lythrum salicaria
Centaurea spp.
Cardaria draba
USDA
code
CIAR4
CANU4
EUES
ELAN
TAMAR2
RHAMN
PHAR3
ARDO4
BRTE
DIFU2
ARM 12
POCU6
POPE10
ALPE4
LYSA2
CENTA
CADR
Expected occurrence by state
(All = MT, ND, SD, NE, IA, KS, MO, IL, Wl, MN, IN,
KY, OH, WV, PA)
All
All
MT, ND, SD, NE, IA, KS, MO, IL, Wl, MN, IN, OH,
WV, PA
MT, ND, SD, NE, IA, KS, MO, IL, Wl, MN, KY, OH,
PA
MT, ND, SD, NE, KS, MO, IL, OH, PA
All
All
KS, MO, IL, KY, WV
All
MT, SD, NE, IA, KS, MO, IL, Wl, IN, KY, OH, WV, PA
All
MT, ND, SD, NE, IA, KS, MO, IL, Wl, MN, IN, KY, OH,
WV, PA
OH, WV, PA
ND, NE, IA, KS, MO, IL, Wl, MN, IN, KY, OH, WV, PA
All
All
All
136
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7.5 General site assessment
After all bank and riparian data have been collected, an assessment of channel form
and constrainment, and a general site assessment are conducted. Conditions on both
shorelines in a 1000-m reach centered on the cross-channel transect, as well as conditions
upriver that might influence the conditions at the site, are considered for the general
assessment. The photographs in the site dossier (Section 4) are often useful for the general
site assessment. Channel form and constrainment are evaluated using the procedures in Table
7-10. General assessment procedures are presented in Table 7-11.
Table 7-10. Channel form and constrainment.
1. Fill in the site ID and data on both sides of the channel and general assessment form (Figure 7-
7).
2. Classify the channel pattern at the site as single, anastomosing, or braided. Anastomosing
channels have relatively long major and minor channels branching and rejoining in a complex
network. Braided channels also have multiple branching and rejoining channels, but these
sub-channels are generally smaller, shorter, and more numerous, often with no obvious
dominant channel(s) .
3. Characterize channel constraint at the site. Evaluate whether the channel has a broad alluvial
floodplain likely to flood. If not, determine if the channel is constrained by a V-shaped valley, an
incised channel, or a narrow valley that necessarily limits the ability of the channel to migrate at
high flows. For floodplain reaches, determine if the floodplain is protected wholly or partially by
levees.
4. For constrained channels, determine the constraining features. An unconstrained channel such
as a floodplain river, can be locally constrained at non-flood flows by artificial revetment
(riprap).
5. Estimate the percent of the channel constrained (both banks) in the 1000-m reach centered on
the cross channel transect (i.e., the 500 m primary and secondary transects). For
unconstrained channels with no artificial revetments, percent constrained = 0%.
6. Estimate valley width visually, if possible.
Watershed activities and human disturbance at the site are rated as not observed, low,
moderate, or heavy. The distinction is subjective. For example, if there are only one or two
houses visible from the river, the rating for "residences" would be low. If one shoreline is
137
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adjacent to a residential area that encroaches on the riparian zone, the rating for "residences"
would be heavy. Similarly, a small patch of clear-cut logging on a hill overlooking the river would
be rated as low disturbance. Logging activity in the riparian zone would be rated as heavy
disturbance.
A subjective characterization of the level of development at the site and the overall
aesthetic quality is required. Rate each of these attributes on a scale of 1 to 5. For
development, assign a "5" rating if it is pristine, with no signs of any human development;
assign a rating of "1" if the river at the site is totally developed (e.g., the river bank is lined with
houses, or the riparian zone is otherwise completely developed). For aesthetic quality, base
your decision on features of the site that reduce your enjoyment of the site (e.g., trash, foul
odors, rip rapping, development). Also, rate the presence/absence of beaver activity, the
dominant land use at the site, and the dominant age class of the riparian forest. Beaver activity
may include bank lodges, felled trees, cuttings, dams in tributaries or backwaters, or actual
sightings.
138
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Table 7-11. Procedure for general site assessment.
1. Use your perceptions of the site obtained during the course of sampling to make a general
assessment of the 1000-m reach centered on the cross-channel transect (i.e., the 500-m
primary and secondary transects).
2. Rate each type of watershed activity or disturbance listed on the form as either not observed,
low, moderate, or heavy on the channel and general assessment form (Figure 7-7). Ratings
are subjective; extensive effort to quantify the presence and intensity of each disturbance is not
required.
3. Assign a rating of 1 (highly disturbed) to 5 (pristine) based on your general impression of the
intensity of impact from human disturbance. Also, assign a rating to the river based on overall
aesthetic quality based on your opinion of how suitable the river water is for recreation and
aesthetic enjoyment:
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 Trashed; enjoyment nearly impossible.
4. Rate the presence of beaver activity at the site from 5 (intense) to 1 (absent). Beaver activity
may include bank lodges, felled trees, cuttings, dams in tributaries or backwaters, or sightings.
5. Determine the dominant land use at the site. Pick one land use from among forest, agriculture,
range, urban, and suburban/town. If there are other major land uses, make a note of them in
the general assessment section of the form. If forest is the dominant land use, make a guess at
the dominant age class of the forest (0-25, 25-75, or >75 years).
139
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7.6 QA Considerations for riparian habitat sampling
A great variety of conditions among sites, plots, and subplots may be encountered in
riparian sampling. Standardization, training, and crew specialization enhance the QA of riparian
sampling. Some QA considerations for riparian habitat sampling are provided in Table 7-12.
Table 7-12. QA considerations for riparian habitat sampling.
Attempt to expend equal effort for each plot and subplot despite variation in access, vegetation
cover, and general sampling ease.
Practice calibrating visual estimates of cover, distances, stem diameters, plot dimensions, etc.
prior to actual data collection.
Collect specimens of possible invasive species to show to a botanist when identification is
uncertain.
Use the "way-point averaging" function on the handheld GPS to acquire the most accurate site
coordinates.
To minimize intra-crew variability, crew members should specialize in tasks as much as
possible (e.g, cover estimates, invasive species identification)
For the shoreline type and macrohabitat estimates, be sure that all of the intervening shoreline
between bank stations has been observed. Use the shoreline stations at 50-m intervals as a
distance calibration for estimating percentages of shoreline in each type or habitat category.
Conduct the general assessment for the whole site after all other sampling activities to insure a
comprehensive perspective.
140
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7.7 Equipment and supplies
Table 7-13 lists the equipment and supplies required to conduct all the activities
described for characterizing riparian and bank physical habitat. Generic supplies required for all
EMAP-GRE field sampling are listed in Table 2-5.
Table 7-13. Equipment and supplies for sampling riparian habitat.
Qty
1
1
1
1
2 rolls
1
1
1
1
1
1
1 set
1 set
Item
Surveyor's telescoping leveling rod
Clinometer (or Abney level) with percent and degree scales (optional)
Convex spherical canopy densiometer (Lemmon Model B), modified with taped "V"
Bearing compass (backpacking type)
Biodegradable surveyor's plastic flagging (2 colors)
Digital camera
Extra memory card for digital camera
100 m fiberglass tape
Meter stick for bank angle measurements
Laser rangefinder with :>1000-m range for channel width measurements
Laser hypsometer (e.g., Opti-logic LH series) for bank and tree height
measurements
Laminated invasive-species reference guides
Riparian habitat forms
141
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7.8 Literature cited
Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and
Monaco, P. A. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Non-Wadeable
Rivers and Streams, U.S. Environmental Protection Agency, Washington, DC.
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Ringold, P. L., Magee, T. and
Cappaert, M. R. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams,
U.S. Environmental Protection Agency, Office of Research and Development,
Washington, DC.
Lemmon, P.E. 1957. A new instrument for measuring forest overstory density. Journal of
Forestry 55:667-669.
Sappington, L., D. Dieterman, and D. Galat (editors). 1998. Missouri River Benthic Fish
Consortium 1998 Standard Operating Procedures to Evaluate Population Structure and
Habitat Use of Benthic Fishes along the Missouri and Lower Yellowstone Rivers. USGS
Biological Resources Division, Columbia Environmental Research Center, Columbia,
MO.
Ringold, P., S. Cline, M. Kentula, M. Bollman, T. Magee, S. Bryce, and P. Lattin. 2001. Draft
Research Plan, EMAP Western Pilot Riparian Investigations in the John Day and Lower
Deschutes Basins of Eastern Oregon. U.S. Environmental Protection Agency, Corvallis,
OR. Draft.
142
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EMAP-GRE BANK AND SHORELINE MACROHABITAT FORM
Draff
Shoreline Station Location; Bank and Channel Width Characteristics
Station
A
C
E
G
I
K
Latitude Longitude
Wetted
Width (m)
Bank Bank Bank Angle
Height (m) Width (m) (circle one) aa
F G S V 0
F G S V 0
F G S V 0
Shoreline Type Between Stations (% by type - sums to 100)
Stable Bank
Aggrading Bank
Eroding Bank
Riprap
Other revetment (Explain in comments)
Other {Explain in comments)
Flag
0-100
A-C
100-200
C -E
200-300
E-G
300-400
G-!
400-500
-K
Macrohabitat Type Between Stations <% by type, may sum to > 100)
Charme cross-over (straightaway)
Outside bend
Inside bend
Pool
Tributary mouth < 5m wide
Connected secondary channel < 5m wide
Unconnected secondary channel (backwater) < 5m wide
Other {Explain in comments)
Flag
Flag
0-100
A-C
100-200
C-E
200-300
E-G
300-400
O-l
400-500
I-K
Determining Bank Angle
Vs;y SiBfip C^erht Ml
\{TK,n--i Hin/
Steep (30-75*) \ /
^""\ \ /
>«WW"?\ \ /
F = Flat (90°)
Comments
Flag codes: K = No measurement made, U = Suspect measurement, F1 ,F2, etc, = misc. flags assigned by each field crew.
Explain all flags in comment section.
21.
Figure 7-3. Bank and shoreline macrohabitat form (one per site).
143
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RIPARIAN CLASSIFICATION AND HUMAN INFLUENCE
Draft
SITE 10: QRW04448-
OATE:
/ 2 0 0
ANNUAL VISIT
NUMBER:
D1 D2 STATION: DA
Land Use and Land Cover Classification
Subplot
Land Cover circle one)
B D A X HM SM F OT
B D A X HM SM F OT
B D A X HM SM F OT
Flag
Canopy Densiometer (0-1 7)
Landward Downriver Uotwer To. river
F ag
River edge
(Om)
Sytopjot 2: iann-slde edf e
(20m)
Latitude
Subplot 1. river edge (0 m|
Distance to wemed edge |m)
Subplot 2. {20 m)
iand-s cie edge
Longitude
Flao
Human Influence (circle Yor N)
SIKes, riprap,
revetments
By ii dings -
industrial/commercial
Buildings - residential
Roads ©r rails
Outlst pipes
Doeks'marina
Paries or lawns
Row crops
Grazing
Recent logging
Mining
Other (Flag & explain)
Other (FSag 4 explain!
Opposite s
of river
Y N
Land Cover Codes
B - Bare, un-vegetated. non-forest.
non-agriculture, undeveloped
D = Developed, urban
A = Agriculture
X = Xeric (upland) snrubs or
hert>aeeous
HM = Mesle Iripanan) herbaceous
SM - Mesie (riparian) weody
shrubs
F - >10% large tree cover
OT - Other (explain in comments)
Vegetation Coverage Codes
0 - Afcsent
1 = Sparse f<10%)
2 = Moderate (10-40%)_
3 » Heavy (40-75%)
4 = Very heavy (s-TSfo)
Woo5 m tall)
Woody Vefetaton Type
819 trees (>0.3 m DBH)
Smalt trees ^<0.3 m DBH)
D c M N
01234
01234
D C M N
01234
0 1 2 S 4
D C M N
01234
81234
Understory {0.5 to 5 m tall)
Woody Vsgetaton Type
Woody shru&s and
seedlings |<0.1 m DBH)
D C M N
01234
01234
D e t» M
01234
0 I 2 3 4
D C M N
§ 1 2 3 4
91234
Ground Cover |<0.S m tall
Woody Vegetaton Type
Woody shrufes and
seedlings |<0.1 m DBH)
Heroaeeous
Bare ground
D c M N
01234
01234
0 1 2 S 4
D C M N
9 1 2 S 4
0 1 2 S 4
§1294
D C M N
01234
91234
* 1 2 3 4
Figure 7-4. Riparian classification and human influence form (one per station).
144
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• m
10457
SITE ID: GRW04449-
EMAP-GRE RIPARIAN LEGACY TREE AND ««»
INVASIVE PLANTS by
-------
BUBBLE LEVELED'
Figure 7-6. Use of the modified convex spherical canopy densiometer. In this example, 11 of
the 17 intersections show canopy cover, giving a densiometer reading of 11. In
heavy cover, it is easier to count the clear intersections and subtract from 17.
Note proper positioning with the bubble leveled and face reflected at the apex of
the "V."
146
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EMAP-GRE CHANNEL AND GENERAL ASSESSMENT (front)
SITE ID: GRW04448-
DATE:
/ 2 § 0
ViSIT
NUMBER:
D 1 D 2
Reviewed
by (Initials);
Channel Form and Constraint in Reach 500 Meters up and downriver from X-site
Channel Pattern (check one)
H Single Channel
FLAG
Anastomosing Channel Braided Channel Other (Flag and explain in
D long 2i«,»rx channels) d 'short 2i«iv channels) d comments)
Channel Constraint (check
Channel very Channel in broad
U in V-shaped LJ valley but
valley (unlikely to flow (unlikely to flow
overtwnk or cut new overbank or cut new
channels in flood) channels in flood)
Channel In narrow
U valley but very
constrained (valley
width <10xbankfuli
width)
one)
Channel is
LJ unconstrained in
valley (floodplain
present) with fevee
flood control
Constraining Features (check one}
i— 1 Bedrock (Channel in
bedrock dominated
gorge)
FLAG
Channel is
LJ unconstrained in
broad valley
(floodplain present)
with no levee flood
control
FLAG
rj Hillslope (channel In a r-j T /hi D Modified I channel r-j No constraining
narrow V-shaped constetad by/incision) ^f -StltL'f8^ * *?T '*"« ,
' ' other revetment) migration possible)
Percent of channel constrained
(0-100%)
Bankfull width (m)
FL
*G
Channel and Vailey
WiHth
Measurements
Estimated valley width |m)
Estimate of valley width not
possible (check box)
Overall site characteristics in Reach 500 Meters yp and downriver from X-site (check one in
Deveiopment
Aesthetics
Beaver Activity
Dominant Land Use at MCS
fpiek on© only)
Pristine
D5
D 5
Intense
D5
D4 D 3 D
D4 D3 D
D4 n 3 n
D Forest D Agriculture D Rang* D
Dominant Forest Age Class
WATERSHED ACTIVITIES
Residential
N L M H Residences
M L H H Lawn*
N L M H CsrMrycMon
M L II H Pipes.
N L M H Dumping
M L M H
N L M H BrMoeCitherts
N L M H Sewage «t,
N L M H otter (FtaB
aw! explain!
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2 D 1 Highly Disturbed
2 D 1 Unappealing
2 D 1 Absent
Urban D Suburban
D >?5 years
AND DISTURBANCES OBSERVED IN REACH 500 METERS UP AND DOWNRIVER FROM X-SITE FLAG
Recreational
N L M H Hiking Trails
N L H H
M L M H tanipng
N L M H TmMJtter
N L M H
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H L M H Oreilgini
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N L M H Fish Stacking
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N L M H Riprap
N L M H Ottwrffltio
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Section 8
Fish
Erich B. Emery1, Jeff A. Thomas1, Mark Bagley2, and Ted R. Angradi3
EMAP-GRE fish sampling methods are designed to collect all but the rarest fish
inhabiting the near-shore habitat at a site. The sample collected is assumed to accurately
represent the proportional abundance of the littoral fish assemblage at the site. Fish sample
data include species composition, and the size and condition of individual fish. Other measures
of assemblage structure and function can be calculated from the data and combined into
indices of biotic condition potentially useful for assessing the condition of Great Rivers (Simon
and Emery 1995, Emery et al. 2003).
Fish assemblage data are collected by electrofishing with a three-person crew during
the day. A subsample of fish are retained for analysis of tissue contaminants (Section 9). After
electrofishing, the crew collects fish habitat data. The procedures in this section are
substantially revised from previous EMAPfish sampling methods (Peck et al., unpublished
drafts). Habitat sampling methods are based on ORSANCO methods.
8.1 The electrofishing transects
Upon arriving at the site location, the fish-sampling crew flags the primary and
secondary 500-m MCS transect at 100-m intervals (if not already flagged by the river-sampling
crew). Fish are sampled by daytime electrofishing along the two 500-m shoreline transects.
The primary transect extends upriver from the intersection of the cross-channel transect and
the target shoreline (river right or river left) identified in the design file (see Figure 4-1). The
secondary transect extends downriver from the intersection of the cross-channel transect and
Ohio River Valley Water Sanitation Commission (ORSANCO), 5735 Kellogg Avenue, Cincinnati, OH 45228
U.S. Environmental Protection Agency, Office of Research and Development, National Exposure Research
Laboratory, Ecological Research Division, 26 Martin Luther King Dr., Cincinnati, OH 45268
U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
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the target shoreline (unless it has been adjusted as described in Section 4).
The shoreline electrofishing zone extends out from shore to a depth of 6 m (20 ft) or a
distance of 30 m (100 ft), whichever is closer to the shore. Electrofishing is conducted for a
minimum of 1800 seconds (0.5 h) of total shock time to collect fish from the designated zone.
Increased shock time will be necessary to fish shorelines with abundant cover.
8.2 Electrofishing
8.2.1 Boat specifications and electrode configuration (recommended)
The standard EMAP-GRE electrofishing boat is a modified 5.5-m aluminum jon boat
with a welded-aluminum hull. The boat is equipped with a 90-hp outboard motor for river
navigation. An auxiliary 25-hp motor is mounted starboard of the main motor, just behind the
driver. The smaller motor is used to maneuver the boat during electrofishing; it allows operation
at slower speeds and in shallower water than the main motor. Other configurations are
permissible.
A generator supplies power to a control box, which in turn controls the electrical field
configuration. A single boom extends 2.5 - 3.0 m from the front of the boat with a single anode
dropper affixed to the end of the boom. The boat's hull serves as the cathode. There are 3 "kill"
switches on board. There is a kill switch on the control box which shuts off all power coming
from the box, there is a positive-pressure kill switch operated by a foot pedal mounted on the
front deck, and there is a hand-held switch operated by the driver during operation of the
electrofishing equipment. All 3 switches must be "on" in order to activate the electric field. This
ensures redundancy within the electrical safety system: the driver and one crew member can
both kill the electricity from the generator in an unsafe situation.
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8.2.2 Electrofishing procedures
Before starting the electrofishing run, all safety "kill" switches should be tested by
starting the generator, turning all switches to the "on" position, and then throwing each switch to
the "off" position to make sure each is working properly.
Electrofishing should not begin until 1000 h. Beginning at the upriver end of the primary
500-m MCS transect, the driver maneuvers the boat downriver parallel to the shoreline (Figure
8-1). The two other crew members stand in the bow of the boat and net all fish that are
stunned. Stunned fish are placed in an aerated live well. Voltage and amperage adjustments
may be necessary to ensure that a minimum of 3000 watts of output power are maintained at all
times. It may be necessary to adjust power based on sampling effectiveness and incidental fish
mortality. Trained crew members should be able to determine whether insufficient or excessive
power is being used.
During the electrofishing run, the boat is navigated through the shoreline zone at a
speed sufficiently slow to allow the fish netters to recover all stunned fish, including small fish,
such as darters, which are difficult to see and do not always rise up off of the bottom when
stunned. The boat should be moved in a serpentine fashion parallel to the shoreline, ensuring
that the electrical field is passing over the shallow littoral areas as well as over the deeper
channel margin, and ensuring that as much of the zone as possible is transected by the path of
the field. The path of the boat (Figure 8-1) should be analogous to the motion of a person using
a metal detector: a side-to-side path with complete lateral coverage and a slow forward pace.
Care should be taken to thoroughly work the electric field around objects such as snags,
downed trees, piers, boulders, and other potential fish cover until each object yields no more
fish. The field may have to be held over the structure for a few seconds to allow the fish to
wriggle out of the cover and up into the field. The minimum electrofishing time for each transect
is 1800 seconds of shock time. Along shorelines with swift current and/or little cover, it may be
necessary to electrofish the transect twice to achieve the minimum shock time. There is no
upper limit for electrofishing time. Electrofishing procedures are described in Table 8-1.
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A large live well (> 300 L) should be used to ensure adequate holding capacity for all the
fish collected in the 500-m transect. A strong and reliable aerator should be used to maintain
oxygen levels in the tank. If an excessive number offish are captured, it may be necessary to
change the water in the live well during the run. Usually this is done after the electrofishing run
has been completed, just prior to processing the fish. Fish that appear overly stressed as
indicated by loss of righting response should be processed immediately and released.
Individuals returned to the water during the electrofishing run should be released behind the
boat and in deeper water, to ensure that they are not recaptured. At the completion of each
500-m electrofishing run, the crew leader records the end time and the total shock time, in
seconds, on the fish sampling form (Figure 8-2).
Sample point (X-site)
i m
100 in
- -
30i m
400 m
SiOnt
Figure 8-1.
Recommended path of electrofishing boat showing equal coverage of shoreline
and channel lateral margins as well as complete application of the field through,
over or around cover objects. The zone should not extend greater than 30 m
from shore, or to a depth greater than 6 m. Not to scale.
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Table 8-1. Electrofishing procedures.
1. Complete the header information on the fish sampling form (Figure 8-2) including site ID, date,
coordinates of transect starting point at downriver end, transect (primary or secondary), target
shoreline, and transect length. Use a different fish sampling form for the primary and
secondary transects^
2. Obtain the Secchi depth from the river-sampling crew, if possible. Otherwise measure Secchi
depth (see Table 5-3)
3. Navigate to the upriver end of the electrofishing transect and make all necessary electrical
connections. Extend and secure the boom, fill the live well, and turn the aerator on.
4. At a location outside the sampling zone, test the electrofishing unit and kill switches. Using
pulsed DC, adjust voltage and amperage to maintain a minimum power output of 3000 watts.
Make voltage and amperage adjustments to ensure that fish are being rolled easily, that
smaller fish such as darters are effectively stunned, and that fish are not being injured. Record
power output data on the form (volts, watts, amps, pulse rate, pulse width).
5. Record the begin time on the fish sampling form and begin electrofishing. From the top of the
zone, proceed slowly downriver, following a serpentine path parallel to the shoreline (Figure 8-
1). Attempt to net all stunned fish. Avoid netting bias toward larger individuals. Do not attempt
to fish in water deeper than 6 m (20 ft). Stay close to shore and fish the shallower margins. If
the water is generally shallower than 6 m, the path of the boat should extend out into the
channel no more than 30 m (100 ft) from shore. Carefully maneuver the boat around instream
cover, fishing slowly to ensure that the cover is yielding no more fish before moving on.
6. Attempt to fish the transect as thoroughly as possible, but do not place the crew in danger in
order to fish particular habitats. Safety is the first concern. If part of the transect cannot be
fished safely, note this in a comment on the form.
7. At the end of the sample transect turn off the electrofishing gear and record end time and total
shock time (Figure 8-2).
8. The minimum electrofishing time for each transect is 1800 seconds of shock time. Along
shorelines with swift current and/or little cover, it may be necessary to electrofish some or all of
the transect twice to achieve the minimum shock time. There is no upper limit for electrofishing
time.
9. After processing the sample, repeat steps 1-7, as appropriate, for the secondary 500-m
transect.
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8.3 Sample processing
Sample processing includes identifying fish to species, examining them for external
anomalies, measuring and weighing, preserving small specimens for later processing,
photographing voucher specimens, and selecting specimens to be retained for tissue
contaminant analysis. At two sites sampled in each year, fish-fin tissue is collected for DMA
analysis. Fish are recorded by their complete American Fisheries Society (AFS) common name
after Nelson et al. (2004). For processing, one crew member records data on the fish sampling
form (Figure 8-2) while the other crew members sort, identify, weigh, and measure fish. After
they are processed, fish from the primary transect should be released where they will not be
recollected from the secondary transect.
All small specimens (<12 cm) and specimens that cannot be identified with certainty in
the field should be preserved in formalin for later processing by the sampling crew. Preserved
specimens will eventually be deposited in a museum collection. With the exception of small
fish collected for DMA analysis, which must be preserved in 75% ethanol, all fish will be
preserved in 10% buffered formalin. Fish sample processing procedures are described in
Table 8-2.
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Table 8-2. Fish sample processing procedures.
1. Ifthehandling of threatened or endangered fishes is permitted under the collecting permit, they
should be processed first in order to expedite their return to the water. Otherwise they should
be released immediately.
2. For each larger specimen that can be identified to species and accurately weighed in the field,
record the complete AFS common name (Nelson et al. 2004). All small specimens (<12 cm)
and specimens that cannot be identified to species with certainty in the field should be
preserved in one or more jars as a field composite and retained by the crew for later laboratory
processing (fish voucher procedures described in Table 8-3). Do not record data for these
specimens in the field. Potential small voucher specimens should be preserved in the same
containers as the composite fish sample.
3. At the first or second and last site sampled in 2005, fish tissue for DMA analysis will be
collected. These methods are described in Table 8-6.
4. Select and retain fish for tissue analysis (see Section 9).
5. Examine each fish for DELTs (deformities, erosions, lesions, and tumors). Record the
presence of DELTS on an individual fish or among a batch of small fish using the codes in
Table 8-4. Other abnormalities (e.g., blind eyes, pop-eye, fungus) can be recorded using flags.
6. Using a measuring board marked with 3-cm size classes, record fish length by size class (e.g.,
fish < 3 cm long are in size class 1, fish >3 and £ 6 cm are in size class 2, etc.
7. Record the weight of each fish in kg (1g = 0.001kg). Fish too small to be weighed should be
retained for laboratory processing. Be sure to release fish in a location where they will not be
recollected during sampling of the secondary transect.
8. In the laboratory, sort the preserved composite sample to species. Refer unknowns to a
taxonomic expert for identification (no expert is specified by EPA - crews may use any qualified
ichthyologist). Process the specimens as in the field and record the data on the original field
forms. Fish too small to weigh individually may be weighed as a batch. Extract and label
individual species voucher specimens (Table 8-3). Do not discard the processed specimens.
They should be retained in the original jars with fresh formalin as a composite voucher sample
to be deposited in a museum collection.
9. Field forms should be completed by the field crews. All fish should be identified to species and
weighed before the forms are sent to the EMAP data center.
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8.3.1 Unknowns and voucher specimens
Each crew member should be familiar enough with the large-river fish assemblages in
the region to identify most larger specimens. A professional ichthyologist familiar with the fish
species of the region should perform final identifications of unknowns. Obtaining identifications
of unknowns is the responsibility of the crews, but MED can facilitate identifications if
necessary.
Vouchers, in the form of a photograph (Figure 8-6 is an example) or as a preserved
specimen, should be retained as a reference for every species allowable under the collecting
permits. Each fish-sampling crew should photograph or collect one voucher specimen for each
different species encountered each year. All questionable fish and all small fish (<12 cm)
should be preserved at every site as a field composite voucher for later laboratory
identification by the fish-sampling crew. Table 8-3 describes preparation of photo vouchers
and preserved specimen vouchers. Large species can usually be adequately documented by a
digital photograph.
An effort should be made to document with a photo or by collection any known or
suspected non-indigenous exotic or invasive fish species. The collecting permit may specify
that certain species not be returned alive to the water. A spatially-referenced and frequently-
updated database of non-indigenous fish species of the U.S. can be searched at
http://nas.er.usgs.gov/. Fish-sampling crews should be familiar with the potential and reported
non-indigenous species in the river and regions they are sampling. Collections of non-
indigenous fishes made during EMAP-GRE sampling should be submitted by fish-crew leaders
to this database via the above web address.
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Table 8-3. Voucher specimens. Note: these methods are modified at fin-tissue collection
sites.
1. There are three types of voucher samples: 1) preserved field composite samples that include
questionable specimens and all small fish, 2) preserved individual species vouchers that are
extracted from the field composites, and 3) photo vouchers for larger fish. A preserved
individual species voucher or photo voucher should be retained for every species captured by
every fish-sampling crew each year. All fish retained in the field composite are tracked with a
single Sample ID in the header of the fish sampling form (Figure 8-2; multiple jars may be
used).
2. Preserved field composite voucher. Small specimens (<12 cm) and unknowns to be retained
for laboratory processing should be euthanized with a humane method. Euthanized specimens
are placed in a leakproof plastic jar(s) with a screw top with 1 0% buffered formalin (see Table
3.1). Do not cram fish into jars so that they are fixed in a bent position; they should float freely
in the jars. Fish biomass should not exceed 40% of the container contents by weight. Fish
>.12-cm long should be slit open in the lower right abdomen to promote preservation.
3. Fill out a fish voucher label for the jar (Figure 8-7) with EMAP-GRE site number, date, visit
number and jar number. Circle "10% Formalin." Transfer the sample ID number from the label
to the header of field form (Figure 8-2) and the MED tracking form (Sample type CV; Figure 3-
2). If >1 jar is needed, fill out a continuation label including the sample ID from the original fish
voucher label. Add sufficient formalin to each jar to cover the fish. Place the voucher label(s) on
thejar(s)and cover with clear tape.
4. Preserved individual species vouchers (optional). Specimens may be extracted from the
preserved field composite to become individual species vouchers. Fish of the same species
should be placed in a leakproof plastic jar(s) with a screw top. Do not cram fish into jars so that
they are fixed in a bent position; they should float freely in the jars. Fish biomass should not
exceed 40% of the container contents by weight. Fish >_1 5-cm long should be slit open in the
lower right abdomen to promote preservation.
5. Fill out a new fish voucher label for the jar (Figure 8-7) with EMAP-GRE site number, date, visit
number and jar number. Circle "10% Formalin." Transfer the sample ID number from the label
to the MED tracking form (Sample type FS; Figure 3-2) and to the field form. Add sufficient
formalin (See Table 3.1) to each jar to cover the fish. Place the voucher label(s) on jar(s) and
cover with clear tape.
6. Photo vouchers. Place the fish on the measuring board. Place a completed Photo Fish
Voucher Label (Figure 8-7) below the fish with the presumed common name of the fish, EMAP-
GRE site number, transect, and date. Use a digital camera to take a high-resolution picture of
the fish. Check the quality of the image before releasing the fish (Figure 8-6). Record the image
file name on the form as a flag comment after the camera images have been downloaded from
the camera. Save the image files to a folder named with the site number. Back up the image
files as soon as possible. If you rename the image files in the office, be sure to provide the new
name on the form.
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8.3.2. External examination for anomalies
During processing in the field or in the laboratory, both sides of each fish should be
examined for external anomalies. Readily-identified external anomalies include deformities;
erosion of the fins, barbels, and gill covers; lesions; and tumors. Photographs of each type of
anomaly are shown in Figure 8-5 (from Moulton et al. 2002). Smith et al. (2002) has additional
guidance for assessing anomalies. Codes for each type of anomaly are given in Table 8-4.
Table 8-4. External anomaly codes (DELTs).
Category
Deformities
Erosion
Lesions
Tumors
Other
Code
DE
ER
LE
TU
OT
Description
Skeletal anomalies of the head, spine or body shape
Eroded barbels, fins, or gill covers; substantial fraying or
Open sores or exposed tissue; raised warty outgrowths
reduction
Areas of irregular cell growth which are firm and cannot be broken
open easily (masses caused by parasites can be broken open easily)
Flag and describe in a comment
8.3.3 Length and weight measurements
Procedures for recording length and weight measurements are presented in Table 8-2.
Total length (Figure 8-6) is used to determine the 3-cm size class to which each fish belongs.
Weights are taken using a spring-dial or digital scale and recorded to the nearest gram (0.001
kg). Lengths and weights are recorded on individual lines on the fish sampling form (Figure 8-
2). Fish too small to be weighed individually can be grouped into 3-cm size classes and
weighed as a batch.
8.3.4 Fish fin-tissue samples for DMA analysis
At two sites per year, fish-fin tissue is collected by caudal punch or scissor clip (Table 8-
5). DMA will be extracted from this tissue for genetic analysis. The findings will be used to
check identifications of specimens, examine genetic variation within species, and identify
hybrids. At fin-tissue collection sites, the first 20 large specimens of each species are fin
punched/clipped and photographed. All small fish at these sites will be preserved as a
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composite sample in 75% ethanol, and processed (identified, examined for anomalies,
measured, and weighed) in the lab as usual. The specimens are then be sorted into species
and preserved in separate jars (one jar per species). The jars are labeled and shipped to NERL
at the end of the season for fin-tissue sampling and DMA analysis.
In addition to the regular sampling of fin-tissue for DMA analysis, crews may collect
additional fin-tissue samples of unknown or unusual specimens at any site and submit them for
DMA analysis using the same sample tracking procedures. These specimens must be
preserved in ethanol for DMA analysis. For questions specific to the DMA analysis, contact Mark
Bagley (513-569-7455).
Table 8-5. Processing fish for fin-tissue DMA analysis.
1. At two sites each year DMA tissue is check "DMA sample site?" box at the top of the fish
sampling form (Figure 8-2). Do not use the same site (due to revisits) for both samples.
2. Large fish. The first 20 large fish of each species (>12 cm; size class 5 or larger; including
fish retained for fish tissue contaminants - Section 9) across both electrofishing transects at
DMA sample sites should be identified and data should be recorded as usual (Table 8-2). For
each fish, collect a punch or clip of the caudal (tail) fin that is 0.5-1.0 cm2 in area. It may work
best to punch the fish while it is held in the net. Keep a running count so that not more than 20
fish of each species (across transects) are punched. It is acceptable for all DMA samples to
come from the first transect.
3. Place the fin-tissue sample in a #1 paper coin envelope (provided by EPA). Fill out and affix an
adhesive Fish Tissue DMA sample label (Figure 8-7) to the coin envelope. Pre-labeling the coin
envelopes with blank sample labels is recommended.
4. Place the labeled coin envelope next to the fish and take a digital photograph. Release the fish
unless they are to be retained for a fish tissue contaminant sample.
5. Repeat for each large fish of each species in the sample (maximum of 20 per species per site).
Between fish, rinse hands and punch/scissors by vigorous agitation in river water.
6. Transfer the sample ID from the label to the field form and the NERL tracking form for each
sample (sample type DS). Use as many NERL tracking forms as necessary.
7. Download and back up digital images which will be transferred to EPA at the end of the season.
Record the image file name (and common name) on the NERL sample tracking form.
Continued
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Table 8-5. Processing fish for fin-tissue DMA analysis, continued.
8. Place all the sample envelopes fora site in a zip-lock plastic bag labeled with the site ID and
refrigerate. Ship the samples weekly to NERL along with the fish tissue contaminant samples
(Section 9).
9. Small fish. All small fish (<12 cm; size class 4 or smaller; excluding fish retained for fish
tissue contaminants - Section 9) should be preserved as a composite sample in 85% ethanol.
In the lab, the fish should be processed as usual (Table 8-2). After the data are recorded, the
fish should be sorted to species and each species placed in a separate jar with new 75%
ethanol. Note: If more than a week elapses between sample collection and lab
processing, replace the ethanol in the jar(s) with new 75% ethanol.
10. Fill out and affix an adhesive Fish Tissue DMA label on each jar and cover with clear tape. Seal
the jar with plastic electrician's tape. Transfer the sample ID to the fish sampling form (Figure
8-2). Record the ID number next to the first record for that species that does not already have a
sample ID (because it was also included in the large fish DMA sample).
11. Transfer the sample ID and species common name to the NERL sample tracking form (sample
type DS) and retain samples until they can be transferred to NERL.
12. Note: In addition to the sampling of fin-tissue for DNA analysis at two sites in 2005, crews may
collect additional fin-tissue samples of unknown or unusual specimens at any other site and
submit them for DNA analysis using the same sample tracking procedures outlined above.
8.4 Fish habitat
After electrofishing and fish processing, the fish-sampling crew records physical habitat
data for each 500-m shoreline transect. At six of the points marked on the shoreline (0, 100,
200, 300, 400, and 500 m from the downstream end; Figure 8-8), the crew records substrate
composition at 3-m intervals out from the shoreline, or as close as the boat can get to the
shoreline, to a distance of 30 m from shore. In addition to recording depth and substrate
composition, the crew estimates the amount offish cover for each 100-m segment of shoreline
(Figure 8-8). Methods for quantifying fish habitat are described in Table 8-6.
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Table 8-6. Fish habitat data collection.
1. Navigate to the site using GPS. Communicate with the river-sampling crew to obtain specific
site information that may already be available (e.g., new site coordinates, hazards, Secchi
depth, conductivity).
2. If not already marked, locate the downriver end of the 500-m MCS transect using the GPS
coordinates from the design file, and flag it. Use the trip odometer on a hand held GPS or
other method to mark off the transect in 100-m intervals to 500 m. The 0-m point is located at
the downriver end of the transect, but the habit measurements can be made working downriver
as long as the field forms are filled in correctly.
3. Fill in the header information on the fish habitat form (Figure 8-9), including the site ID, date,
transect start coordinates (at downriver end), whether it is the primary (upriver) or secondary
(downriver) transect, and the transect length.
4. Determine the channel morphology at the site and circle the appropriate macrohabitat (see
Table 7-1).
5. At each of the six points located along the shoreline for each zone (at 0, 100, 200, 300, 400,
and 500 m from the transect starting point), one crew member drops the weighted end of a 30-
m floating rope on the shore at the water's edge or as close as the boat can approach the
shoreline (which represents the inside margin of the electrofishing zone). The driver then slowly
backs the boataway from shore in a line perpendicular to the shoreline. The crew member
holding the rope slowly feeds out the line, keeping the rope tight without dislodging the weight
at the shore end.
6. Record the bank substrate (-3 m DFS [distance from shore]) at 3-m DFS intervals, as indicated
by marks along the floating rope. The person operating the pole probes the river bed several
times and announces the depth (to nearest 0.25 m) and substrate(s). Substrate and depth are
recorded to a distance of 30 m from shore (or where the weight on the habitat rope is dropped).
7. In areas of high current velocity, a hand-held GPS or a laser rangefinder may work better than
the habitat rope for locating substrate probe locations.
8. Substrate composition is recorded as boulder, cobble, gravel, sand, fines, or hardpan, or
multiple for each of the 72 points (12 points at 6 sites).
9. For each 100-m zone along the shoreline (e.g. 0-100 m, 100-200 m etc.) determine the
percentage coverage category of each type offish cover in the electrofishing zone. Circle the
percent cover category for each type present on the form. Fish cover is anything that could
provide cover for a fish. A uniform sand or gravel beach with no overhanging vegetation or
undercut banks would provide 0% cover. For linear cover features (e.g., undercut banks)
estimate the percent of the 100 m with the cover feature.
10. Repeat steps 2-9, as appropriate, for the secondary transect.
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8.5 QA considerations for fish sampling
Crew members should be properly trained in techniques for operating the boat and
electrofishing equipment. Proper use of the equipment, including maintaining the electrical field
and maneuvering of the boat to optimize capture of fish, is critical to ensure that a
representative sample is collected. QA offish sample processing depends on correct
identification of specimens. Crew members should have sufficient training to identify most fish
that are collected. Questionable fish should be retained as voucher specimens. Table 8.7
provides some QA considerations for fish sampling.
Table 8-7. QA considerations for fish sampling.
Sampling should probably not take place if Secchi depth is < 15 cm (6 inches) or if river stage
is elevated > 0.5 m (20 inches) above normal levels. The decision to sample or not is up to the
crew leaders.
The transects should be fished before starting habitat data collection so that fish are not
spooked from the shoreline.
Electrofishing should not begin until 1000 h.
Use a digital camera on a high resolution setting for taking photo vouchers.
All small fish (minnows and questionable small specimens) should be retained as a preserved
field composite voucher sample and processed in the lab by the field crew.
Do not cram fish voucher specimens into jars. They should be free floating so they are not
fixed in a bent position.
Replace the ethanol in sample jars with fresh 75% ethanol if more than one week elapses
between initial preservation and lab processing.
Netters should wear polarized sunglasses.
When netting shocked fish, avoid size bias.
• Avoid shorthand or local common names for fish. Nelson et al. (2004) is the standard.
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8.6 Safety considerations for fish sampling
These rivers are large, navigable systems and are often congested by barge and
recreational traffic. Extreme precautions should be taken when electrofishing, crossing the
channel, and navigating to and from sampling locations. Primary responsibility for safety rests
with the crew leader. However, each member of the three-person crew should be alert, aware
of safety considerations (Table 8-89), trained to recognize safety concerns, and trained in first
aid and CPR.
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Table 8-8. Safety considerations for fish sampling.
The electrofishing unit has a high voltage output and is capable of delivering a fatal shock.
Large (>10 kg) silver carp (Hypophthalmichthys molitrix) can jump >2 m out of the water. People have
been seriously injured by carp collisions. Silver carp are present in the lower reaches of all three ORE
rivers. Be alert for jumping fish while running the river and during electrofishing.
Crew members should be able to swim, and should receive CPR, first-aid, and safe boating training.
The rivers sampled for this project are subject to heavy barge and recreational boating traffic. When
navigating at night, running lights and a spotlight are required so that other vessels are aware of the boat
and so the driver can more easily detect obstacles in the water.
If the generator is running, do not touch the anode or cathode (if a cathode other than the boat hull is
used). Do not touch objects outside the boat. Do not reach into the water. If doing so, make sure all
electricity to the water has been turned off by ensuring that all three switches are in the "off" position
(unit, pedal, and hand switch).
Do not electrofish in high waves or other conditions that may cause sudden motions of the boat that can
cause someone to lose their balance.
Do not fish in the rain. Excessive water running from the deck of the boat into the water may create a
path for current to follow from the water, up onto the deck. Prior to each sampling event, all electrical
"kill" switches should be checked to ensure they are working properly.
All members of the electrofishing crew should wear USCG-approved PFDs whenever in the boat.
Good line of sight and communication should be maintained among crew members at all times. The
generator is loud and often drowns out verbal communication. Hand signals should be used to
communicate boat direction, power on/off, and other vital information.
All crew members should know the location of the nearest hospital.
Use caution around onboard gas tanks. Never refill the generator when it is hot. The generator exhaust
gets extremely hot while in use. Caution should be used to ensure that no item is touching the exhaust
and that all items near the exhaust are secured to ensure they do not shift position while underway and
possibly come in contact with the exhaust.
All electrical connections should be checked prior to use to ensure that proper, tight connections are
maintained. Loose connections can cause sparking and fire.
All crew members should know the on-board location of the cell phone, first aid kit, fire extinguisher, and
truck keys.
164
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8.7 Equipment and supplies
Table 8-9 is a checklist of the equipment and supplies necessary for fish sampling.
Generic supplies required for all EMAP-GRE field sampling are listed in Table 2-5.
8.8 Literature cited
Emery, E.B., T.P. Simon, F.H. McCormick, P.L. Angermeir, J.E. DeShon, C.O. Yoder, R.E.
Sanders, W.D. Pearson, G.D. Hickman, R.J. Reash, and J.A. Thomas. 2003.
Development of a multimetric index of assessing the biological condition of the Ohio
River. Transactions of the American Fisheries Society 132:791-808.
Moulton, S.R.II, J.G. Kennon, R.M, Goldstein, and J.A. Hambrook. 2002. Revised protocols for
sampling algal, invertebrate and fish communities as part of the National Water-Quality
Assessment Program. Open-File Report 02-150, U.S. Geological Survey, Reston, VA.
Nelson, J.S, E.J. Grossman, H. Espinosa-Perez. L.T. Findlay, C.R. Gilbert, R.N. Lea, and J.D.
Williams. 2004. Common and scientific names of fishes from the United States, Canada,
and Mexico, Sixth edition. The American Fisheries Society. Bethesda, MD.
Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and
Monaco, P. A. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Non-Wadeable
Rivers and Streams, U.S. Environmental Protection Agency, Washington, DC.
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Ringold, P. L., Magee, T. and
Cappaert, M. R. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams,
U.S. Environmental Protection Agency, Office of Research and Development,
Washington, DC.
165
-------
Simon, T.P. and E.B. Emery. 1995. Modification and assessment of an index of biotic integrity
to quantify water resource quality in Great Rivers. Regulated Rivers: Research and
Management 11:283-298.
Smith, S.B., et al. 2002. Illustrated field guide for assessing external and internal anomalies in
fish. U.S. Geological Survey, Information and Technology Report, 2002-0007, 46 p.
www.cerc.usgs.gov/pubs/center/pdfdocs/ITR_2002_0007.pdf
166
-------
Table 8-9. Equipment and supplies for fish sampling. Standard boat safety gear is not
listed. Generic supplies required for all EMAP-GRE field sampling are listed in
Table 2-5
Qty
1
1
2
1
1
1
2
2
2pr
2
1
2
1
1
1
2
5L
2pr
1
Several
5L
5L
Several
1
1 set
1 set
1 set
Item
Electrofishing anode and boom
Electrofishing control box
Extra boat batteries
Digital camera with macro function, extra memory card
Chainman hip chain with extra string (optional)
Laser rangefinder (optional)
Habitat rope (floating nylon marked in 3 m increments) and anchor (1 L bottle filled with quick-
setting concrete with a loop of rope pushed into the wet concrete)
Habitat pole (2 3-m sections of 3/4" copper pipe with a threaded coupling and caps on
both ends) marked at 0.25 m intervals
Rubber gloves
Non-conducting dip nets with 1/4" mesh
Minnow net for dipping small fish from live well
Measuring board with 3 cm size classes (see Figure 8-6)
1-Kg scale (spring or electronic)
12-Kg scale (spring or electronic)
25-Kg scale (spring or electronic)
Plastic weighing trays
10% borax-buffered formalin
Single hole punch for collecting fin tissue DMA sample (scissors may substitute)
Scalpel for slitting open fish before preservation.
#1 paper coin envelopes (provided by EPA)
75% ethanol for preserving fish after lab processing (2005 DMA sites only)
85% ethanol for preserving fish in field (2005 DMA sites only)
Leak-proof HOPE jars for fish voucher and DMA samples specimens (various sizes from 250 mL -
4L)
Secchi disk and marked line
Fish ID keys
Fish habitat and fish sampling forms
Sample labels
167
-------
•
S1TEID;GRW04449-
EMAP-GRE FISH SAMPLING FORM (front)
„„__ ; f 1 n n ANNUAL VISIT ,-, „ ,-, ,
DATE:. . .'. . .' .2.°.°. . NUMBER: ni D2
Coordinates of Transect Starting Point
Begin time (24h)
Total shock time (sj
End time (24 hr)
Secchi depth
(cm)
Composite Sample ID
Volts
SPECIES
Watts
Number
of Jars
Amps
Revl®w«d
by (Initlafel:
PAGE: •)
DMA SAMPLE SITE?: YesfJ
MCS transect
B Primary
Secondary
Pylse rate
QPPS
. O Hz
1DELTS2 COUNT c^|s WEIGHT (kg)
Reach Length
(m)
Pulse WidtB
(ms|
SAMPLE ID
OF
NO n
Flag
Flag
FLAG
COMMENTS
FJag codes; K = No measurement made, U = Suspect measurement.* F1SF2, etc, = misc. flags assigned by each —::::::::::::::::::::===„
• field crew. Explain all flags in comment section, ^L I ^^m
28. U_^J •
Figure 8-2. Fish sampling form (front).
168
-------
• EMAP-GRE FISH SAMPLING FORM (back)
1 linn ANNUAL VISIT ._. „
SITE ID; GRW04449- DATE; / / 2 0 0 NUMBER- >-> 1
Reviewed ^^_
by (initial^: ^^|
MH
D 2 PASE: OF
MCS transect
Sampto ID; „ _ „
....... D Primary O Secondary
DELTS^ COUHIT WlIGHT(kg) FLA6
FLAG
COMMENTS
Flag codes: K ~ No measurement made, U = Suspect measurement., F1,F2, etc, = misc. flags assigned by each
field crew. Explain all flags in comment section. Draft
•
i~^m
•
Figure 8-3. Fish sampling form (back).
169
-------
• EMAP-GRE FISH SAMPLING FORM (cont.)
SiTEID;GRW0444i- DATE: / / 2 0 0 ™«S? D 1
Reviewed ^^_
by M vi ; ^^
D 2 PAGE: OF
MCS transect
Sampl8lD: D Primary D Secondary
^-^^ COPNT WEIGHT (kfl) SAMPLIID FLAG
FLAS
COIIIIiNTS
Flag codes: K - No measurement made, U = Suspect measurement^ F1,F2, etc. = misc. flags assigned by each
field crew. Explain all flags in comment section. Draft
• so, GSM •
Figure 8-4. Fish sampling continuation form.
170
-------
a, of IDE) a
and
I, of
Ins (ill
c, tf (HI
aiea h of
d tl (TU) ti the
Figure 8-5. DELT anomalies (reprinted from Moulton et al. 2002).
171
-------
FISH
Figure 8-6. Example of a voucher photo. The fish shown is in size class 4 (9-12 cm total
length).
172
-------
FISH
75%
GRW0444i-
Transect number 1
visit number 1
3001??
of
2
2
75% 10% Formalin
/
1 2
1 2
ID#
of
/ / 200_
1° 2°
Common name
FIN DNA
GRW04449-
/ 1200
311255
Common
FIN DNA
GRW04449-
I
Common
Figure 8-7. Fish voucher labels, photo voucher label, fish voucher tag, and fin tissue DNA
sample label. The fish voucher label at upper left is placed on the jar holding the
field composite of preserved specimens retained by the sampling crew for
laboratory identification and individual species voucher samples. If multiple jars
are needed, the continuation label at upper right is used. The fish voucher label
is filled out and placed next to the fish specimen when it is photographed. The
fish voucher tag label is placed in the jar of preserved specimens. The fin-tissue
DNA label is placed on the coin envelope for larger fish (and photographed with
the fish) and placed on the jar for smaller specimens. Use continuation label if
multiple jars are needed for DNA sample. Not actual size.
173
-------
Om
Sample
in
100m
200m
3iOm *
30 m out or
6 m
400m
§00 m
Figure 8-8. Layout of physical habitat measurement locations. At each transect into the
channel, substrate is recorded every 3 m to a distance of 30 m or a depth of 6 m,
whichever is closer to the bank. Depth is recorded out to a distance 30 m from
shore (or wherever the weight on habitat rope is dropped) using the habitat pole
or depth sounder. Each transect starts as close as the boat can get to the
shoreline. Fish cover is quantified for the 100-m segments between substrate
transects. Not to scale.
174
-------
Reyiewec
jit ry"3 FISH HABITAT FORM (front)
Draft SITE ID: GRW04449- DATE: / / 2 0 0 ANNUA
uy
L VISIT
MB£R:
D1
•
D2
Coordinates Of Transect Starting Point (500 m of primary, 0 m of secondary transect)
Latitude
Channel Morphology {check one)
Transect
0
A
(Down
•stream
end
100
c
200
E
DFS
-3
0
3
S
s
12
1S
11
21
24
27
30
-3
0
3
8
9
12
16
IB
21
24
2?
38
-3
0
3
6
9
12
18
18
21
24
2?
3§
Depth (m)
Longitude MCS transect
D Prlmiry Q
Secondary
D D D inside D Pool D Tribytiry
D Connected Sec. Channel D Unconnected Sec. Channel D Other (ti«a and «p!»m ,n conw
entsj
Flag
Substrate Flag Fish Cover Percent Cover Flag
BO
ED
BD
BO
BD
BD
BD
BD
BD
BD
BD
BD
BD
BO
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
BD
CB BR SA FN HP
CB GR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB 6R SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB BR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB BR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB SR SA FN HP
CB BR SA FN HP
CB GR SA FN HP
OB OR SA FN HP
CB OR SA FN HP
CB OR SA FN HP
CB GR SA FN HP
CB GR SA FN HP
CB GR SA FN HP
CB OR SA FN HP
CB OR SA FN HP
CB OR SA FN HP
CB GR SA FN HP
CB GR SA FN HP
0-100m
LWD
Macrophytes
Algae
Overhanging vegetation
Riprap
Natural boulders/cobbles
Boats.'docks
Undercut banks
Live trees^roots
fFlag and explain m
Other comments)
100 - 200 m
LWD
Macrophytes
Algae
Overhanging vegetation
Riprap
Natural boyiders/cobtoies
Boats.'docks
Undercut banks
Live trees/roots
(Flag and explain m
Other comments!
200 - 300 m
LWD
Macrophytes
Algae
Overhanging vegetation
Riprap
Natural bouidere/cobbles
Boats.'docks
Undercut banks
Live trees/roots
(Flag and explain in
Other comments)
o
0
0
0
0
0
0
0
0
0
0
o
0
0
0
0
0
0
0
0
0
0
o
0
0
0
0
0
0
0
0
0
0
o
1
1
1
1
1
1
1
1
1
1
O
1
1
1
1
1
1
1
1
1
1
o
1
\
1
1
1
\
\
1
1
1
•et
£
2
2
2
2
2
2
2
2
2
2
**
0
2
2
2
2
2
2
2
2
2
2
°
o
2
2
2
2
2
2
2
2
2
2
*>-
*r
3
3
3
3
3
3
3
3
3
3
r»
*t
3
3
3
3
3
3
3
3
3
3
r—
3
3
3
3
3
3
3
3
3
3
Substrate codes: 1D= boulder |>2§0 mm), CB= cobble (i4-2§0 mm), BR= gravel (2-64 mm), 8A= sand, FN= fines, HP= hardpan.
• Flag codes: K = No measurement made, U = Syspect measurement, F1, F2, etc, = misc. flags assigned by each field crew.
Explain all flags in comment section.
K
4
4
4
4
4
4
4
4
4
4
E
4
4
4
4
4
4
4
4
4
4
K
4
4
4
4
4
4
4
4
4
4
, H
Figure 8-9. Fish habitat sampling form (front).
175
-------
•
2
Draft
Substation DPS
300
G
400
I
500
K
Upstream
end
•
-3
0
3
S
S
12
IS
IS
21
24
2?
30
-3
0
3
6
i
12
IS
1«
21
24
2?
30
-3
0
3
6
S
12
IS
18
21
24
2?
30
•^ EMAP-GRE HABITAT FORM (back) SIS?* _
™ SITE ID: GRWQ4449- DATE: / / 2 0 0 *NN NUMB
Depth (m)
ER: D1 D2
Substrate Flag Fish Cover Percent Cover Fia§
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD OB OR SA FN HP
BD OB OR SA FN HP
BD CB OR SA FN HP
BD CB GR SA FN HP
BD OB OR SA FN HP
BD CB QR SA FN HP
BD OB GR Sft FN HP
BD CB GR SA FN HP
BD GB GR Sft FN HP
BD CB GR SA FN HP
BD CB GR SA FN HP
BD CB GR Sft FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB GR SA FN HP
BD CB GR SA FN HP
BD CB GR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
BD CB OR SA FN HP
300 - 400 m
LWD
Macrophytes
Algae
Overhanging vegetation
Riprap
Natural boulders/cobbies
Boats/docks
Undercut banks
Live trees/roots
{Flag and explain in
Other comments!
400 - 500 m
LWD
Macrophytes
Algae
Overhanging vegetation
Riprap
Natural boulders/cobbles
Soats^doeks
Undercut banks
Live trees/roots
(Flag and Mcplam m
Other canimnts]
o
0 f
O V **
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
o |
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
0 1 2
•S ?
3 4
3 4
3 4
3 4
3 4
3 4
3 4
3 4
3 4
3 4
? ?
3 4
3 4
3 4
3 4
3 4
3 4
3 4
3 4
3 4
3 4
rlag codes: K = Ns measurement made, U ~ Suspect measurement, FlfF2, etc. = misc. flags assigned by each field crew
Explain all in comment section.
27
m
Figure 8-10. Fish habitat sampling form (back).
176
-------
Section 9
Fish Tissue Contaminants
James M. Lazorchak1, Erich B. Emery2, David M. Walters1, and Spence A. Peterson3
Fish tissue contaminants are an indicator of bioaccumulation of persistent toxic
substances in the environment (Table 9-1), and can be used to estimate exposure to
contaminants associated with fish consumption for higher trophic levels, including humans.
Various studies have been done on fish tissue contaminants that have focused on different
parts of the fish (whole fish, fillets, livers). EMAP-GRE will focus on whole fish because of its
emphasis on the health of the ecosystem. Although whole-fish contamination is primarily an
indicator of risk to piscivorous wildlife, whole-fish data are still relevant for estimating human
exposure to contaminants through fish consumption. Use of whole fish reduces sample
processing effort in the field because no gutting, skinning, or filleting offish are necessary.
At every EMAP-GRE site, two composite fish samples are collected: a small-fish sample
and a large-fish sample. The small-fish sample includes individuals of one species (if possible)
whose adults are small. The large-fish sample includes individuals of one species (if possible)
whose adults are larger. Both sizes offish have advantages. Small fish are more ubiquitous
than the larger fish, and therefore are more likely to be present in sufficient numbers at more
sites. With small species, it may be possible to get a more representative sample of the
contaminant load at the site by combining 20 - 200 individual small fish in a composite sample
than by combining only a few larger fish. Small fish may be a more appropriate indicator for
assessing ecological risk to wildlife because they are more likely to be prey for piscivores than
are larger fish. Large fish are more mobile than small fish, and are more likely to partly reflect
contaminant exposure away from the site. Larger, longer-lived fish may exhibit greater
bioaccumulation and may be more sensitive indicators of contaminants in the environment.
1 U.S. Environmental Protection Agency, Office of Research and Development, National Exposure Research
Laboratory, Ecological Research Division, 26 W. Martin Luther King Dr., Cincinnati, OH 45268
2 Ohio River Valley Sanitation Commission, 5735 Kellogg Avenue, Cincinnati, OH 45228
3 U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Western Ecology Division, 200 SW 35th St., Corvallis, OR 97333
177
-------
9.1 Integration with fish sampling
Fish tissue samples are collected during electro-fishing (Section 8). At each site, two
separate 500-m reaches are electrofished. Composite fish samples for tissue sample analysis
may be retained from either or both of the 500-m electrofishing reaches at the discretion of the
crew leader. The origin of the fish from within the site is recorded on the fish tissue form
(Figure 9-1).
9.2 Selecting fish tissue specimens
The fish-sampling crew should attempt to collect a sample of small fish, and, if possible,
a sample of larger fish. Procedures for selecting fish for tissue specimens are described in
Table 9-2.
9.3 Preparing composite samples
After the fish comprising the small-fish and large-fish composite have been selected
(Table 9-3), they are recorded on the fish tissue form (Figure 9-1) and packaged for shipment
to the laboratory. Procedures for preparing composite samples are described in Table 9-4.
178
-------
Table 9-1. Target analytes for composite fish tissue samples. Detection limit for
mercury is 0.01 ppm. Detection limit for all other analytes is 0.001 ppm. Number
in parentheses is the CAS number. Number followed by a # is the Ballschmitter-
Zell number.
Mercury (7439-97-6)
Aldrin (309-00-2)
Chlordane-cis (5103-71-9)
Chlordane-trans (5103-74-2)
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 I (959-98-8)
Endosulfan II (33213-65-9)
Endrin (72-20-8)
Heptachlor (76-44-8)
Heptachlor Epoxide (1024-57-3)
Hexachlorobenzene (118-74-1)
Hexachlorocyclohaxane [Gamma-BHC/Lindane] (58-89-9)
Mirex (2385-85-5)
trans-Nonachlor (3765-80-5)
cis-Nonachlor (5103-73-1)
Oxychlordane (27304-13-8)
PCS Congeners
2,4-Dichlorobiphenyl, #8 (34883-43-7)
2,2',5-Trichlorobiphenyl, #18 (37680-65-2)
2,4,4'-Trichlorobiphenyl, #28 (7012-37-5)
2,2',5,51-Tetrachlorobiphenyl, #52 (35693-99-3)
2,2',3,51-Tetrachlorobiphenyl, #44 (41464-39-5)
2,3',4,41-Tetrachlorobiphenyl, #66 (32598-10-0)
2,2',4,5,5'-Pentachlorobiphenyl, #101 (37680-73-2)
3,3',4,4' Tetrachlorobiphenyl, #77 (32598-13-3) (coplaner)
2,3',4,4',5-Pentachlorobiphenyl, #118 (31508-00-6)
2,2',4,4',5,51-Hexachlorobiphenyl, #153 (35065-27-1)
2,3,3',4,4'-Pentachlorobiphenyl, #105 (32598-14-4)
2,2',3,4,4',5-Hexachlorobiphenyl, #138 (35065-28-2)
2,2',3,4',5,5',6-Heptachlorobiphenyl, #187 (52663-68-0)
2,2',3,3',4,41-Hexachlorobiphenyl, #128 (38380-07-3)
2,2',3,4,4',5,51-Heptachlorobiphenyl, #180 (35065-29-3)
2,2',3,3',4,41,5-Heptachlorobiphenyl, #170 (35065-30-6)
2,2',3,3',4,41,5,6-Octachlorobiphenyl, #195 (52663-78-2)
2,2',3,3',4,41,5,51,6-Nonachlorobiphenyl, #206 (40186-72-9)
Decachlorobiphenyl, #209 (2051-24-3)
3,3',4,4',5 Pentachlorobiphenyl, #126 (coplaner)
Polybrominated Diphenyl Ethers (PBDE) congeners 47, 99, 100, 153 and 154
Percent Moisture and Lipid content
179
-------
Table 9-2. Procedure for selecting fish for tissue analysis.
1. Small-fish-species com posite. Retain similar-sized specimens (smallest individual >. 75% of
the total length of the largest individual) of a single species from the list in Table 9-3 in priority
order. Select the first species on the list for which a sample of at least 50 g can be collected.
Retain a total sample of the selected species of up to 400 g. The "similar-size" rule can be
violated if necessary to obtain ^. 50 g.
2. Large-fish-species composite. Retain at least three and preferably five similar-sized
specimens (smallest individual >_ 75% of the total length of the largest individual) of a single
species from the list in Table 9-3 in priority order. Try not to retain fish larger than 2 kg. Large
specimens present problems for storage and lab processing.
3. If at least three similar-sized or same-size-class specimens are not available, move down the
list (Table 9-3) to the next large species. Collecting three individuals of the same species is
preferable to collecting a mixture of species, even in they are higher priority species.
4. If at least three similar-sized or same-size-class specimens are not available for any large
species, return to the top of the list (Table 9-3) and composite individuals across size classes
to obtain at least three specimens of a single species.
5. If these criteria still cannot be met, use best judgement to collect small- and large-fish species
composite samples comprised of as many specimens as possible. For small fish, at least 8 g of
fish are needed for analysis. Collect a multispecies composite sample as a last resort.
180
-------
Table 9-3. Fish target species list for tissue analysis. 3-cm size classes refer to
demarcations on the measuring board. The same information appears on the
back of the fish tissue form (Figure 9-1).
Priority
3-cm
SMALL target species
1
2
3
4
5
6
7
emerald shiner
river shiner
spotfin shiner
bullhead minnow
silver chub
another minnow
gizzard shad
€120
<120
<12i
<120
<120
<150
1 -4
1 -4
1 -4
1 -4
1 -4
1 -4
1 -5
LARGE target species
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
sayger
sauger
sauger
iargemouth bass
largemoyth bass
iargemouth bass
other black bass
brown troyt
rainbow trout
channel catfish
channel catfish
channel catfish
freshwater drum
shorthead redhorse
other redhorse
bluegill
iongear sunfish
other sunfish species
common carp
smailmouth buffalo
river earpsucker
flathead catfish
white bassfwiper
quillback
120 -180
180 - 240
>240
180 -240
240 -300
>300
> 180
> 120
> 120
120 - 180
450 - 510
180 - 450
>120
>120
>120
>120
>120
>120
>180
>120
>120
>120
>120
>120
5-6
7-8
>9
7-8
8-10
>11
>7
>5
>5
5-6
16-17
7-15
^5
>5
>5
>5
>6
>5
>7
>5
> 5
>5
>6
>5
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Table 9-4. Procedures for preparing composite fish tissue samples.
1. Fill in site ID and date on the fish tissue form (Figure 9-1).
2. Small-fish-species composite. Record the AFS common name (from Nelson et al. 2004) and
count of individuals in the sample. From the composite collection location choices on the form,
indicate where the fish were collected at the site. Fish may be collected from beyond the end of
the transect if needed (mark "other" for collection location).
3. Euthanize fish with a cervical/cranial blow or other humane method. Use clean hands to
transfer specimens to aluminum foil. Keep hands, work surfaces, and foil clean and free of
potential contaminants (mud, fuel, slime, formalin, sun screen, insect repellant, etc.)
4. On a clean work surface, wrap all the fish in a single piece of aluminum foil, making sure that
the dull side of the foil is in contact with the fish. Place the wrapped sam pie in a 1 gallon self-
sealing plastic bag. Expel excess air from each bag and wrap each bag with clear tape to seal
the sample. Go to step 8.
5. Large-fish-species composite. Record the AFS common name (from Nelson et al. 2004) and
size class of each individual in the sample. From composite collection location choices on the
form, indicate where the fish were collected at the site. Fish may be collected from beyond the
end of the transect if needed (mark "other" for collection location).
6. Euthanize fish with a cranial blow or other humane method. Use clean hands to transfer
specimens to aluminum foil. Keep hands, work surfaces, and foil clean and free of potential
contaminants (mud, fuel, slime, formalin, sun screen, insect repellant, etc.).
7. On a clean work surface, wrap the fish in a single piece of aluminum foil, if possible, making
sure that the dull side of the foil is in contact with the fish. Fish may be wrapped individually if
necessary. Place the wrapped sample in a 2 gallon self-sealing plastic bag (use a separate bag
from the small fish sample). Expel excess air from each bag and wrap each bag with clear
tape to seal the sample.
8. Prepare a fish tissue label for each sample (Figure 9-2). Fill in the site ID and date and circle
the sample type (small or large fish). Record the sample ID (from the labels) on the form.
9. Affix the appropriate label on each bag and cover with clear tape. Place each sample in a
second self-sealing plastic bag.
10. Prepare a continuation label (Figure 9-2) for the outside of each bag. Transfer the sample ID
from the inner label to the outer label. Affix the label to the outside of the outer bag and cover
with clear tape.
11. Place the double-bagged samples in a cooler with bagged ice. If possible, freeze the samples
at the base location prior to shipment.
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9.4 Equipment and supplies
Table 9-5 is a checklist of equipment and supplies required for collecting fish tissue
samples. Generic supplies required for all EMAP-GRE field sampling are listed in Table 2-5.
Table 9-5. Equipment and supplies for collecting fish tissue samples. Generic supplies
required for all EMAP-GRE field sampling are listed in Table 2-5.
Qty
1
1 roll
2
2
1
1
1 sets
Item
Measuring board with 3 cm size classes (see Figure 8-6)
Heavy duty aluminum foil (1 8" size)
1 -gallon self-sealing plastic bags
2-gallon self-sealing plastic bags
Cooler with ice.
Fish tissue form
Fish tissue sample labels
9.5 Literature cited
Nelson, J.S, E.J. Grossman, H. Espinosa-Perez. L.T. Findlay, C.R. Gilbert, R.N. Lea., and J.D.
Williams. 2004. Common and scientific names of fishes from the United States, Canada,
and Mexico, Sixth edition. The American Fisheries Society. Bethesda, MD.
183
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| N^U EMAP-GRE FISH TISSUE FORM
Draft
SITE ID: DATE: / / 2 0 0 AMNUAl
Sample ID Composite Collection Location Dp Dg
(Circle One)
Common Name Size Class
Fish
Sample ID Composite Collection Location DP DS
,,,,,,, (Circle OneJ
Common
tewed tsy ^^H
(Initials): ^H
^H
VISIT n 1 n 2
LJ I LJ £
DB OT
Coynt Flag
DB OT
Count Flag
Composite Collection Location Codes
DP Distributed through the primary SOOm transect
DS Distributed the secondary 600m
DB Distributed through both transects
OT Other (describe in comments)
Flag
Flag codes: K = Mo made, U = Syspecl measurement, F1 ,F2, ete. « misc. by each field crew.
EMpiaint nil flags in comment section.
31.
Figure 9-1. Fish tissue form.
184
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Large
/
visit 1 2
300211
Large
/
Site visit number 1 2
ID
Figure 9-2. Sample labels for fish tissue contaminants. The number at the bottom of the label
on the left is the unique sample ID. The label on the left is affixed to the inner bag
holding the sample. The continuation label on the right is affixed to the outer bag
or can be used for additional bags if needed. Not actual size.
185
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Blank Page
186
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Section 10
Benthic Macroinvertebrates
Ted R. Angradi1, Donald J. Klemm2, Jim M. Lazorchak2, and Brent R. Johnson2
Benthic macroinvertebrates inhabit river bed sediments and adhere to hard substrates in
the water column. Macroinvertebrates have several advantages as indicators of ecological
condition (Barbour et al. 1999, Klemm et al. 1990). They are ubiquitous in all GRE aquatic
habitats and are relatively easy to collect in large numbers in most habitats. Macroinvertebrate
assemblages are typically very diverse and sensitive to a variety of stressors. In some cases,
macroinvertebrate assemblage composition can reveal the nature of the anthropogenic stress
to which the assemblage has been exposed (Barbour et al. 1999).
In EMAP-GRE, benthic macroinvertebrates are collected by the river-sampling crew in
two habitats: shallow (<1 m), near-shore littoral areas, and the surface of large woody debris
(LWD) or "snags" in the main channel. In near-shore littoral areas, benthos samples are
collected by kick sampling. Snags are sampled by boat using a modified kick net. The kick
sampling and sample-processing procedures described herein are adapted, with significant
modification, from Peck et al. (Unpublished drafts).
10.1 Near-shore kick sampling
At each site, two 500-m main channel shoreline (MCS) transects, starting at the
intersection of the cross-channel transect and the MCS (Figure 4-1), are located and flagged by
either the fish- or river-sampling crew, depending on which crew arrives at the site first. The
primary transect is initially flagged at 100-m intervals; intermediate littoral stations at 50-m
intervals are located and flagged using a handheld GPS or by visual estimation during littoral
sampling (a different flag color than at the 100-m stations may be used). At each of the
resulting 11 evenly-spaced points along the 500-m MCS transect (stations A-K; Figure 4-2), two
1 U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
2 U.S. Environmental Protection Agency, Office of Research and Development, National Exposure Research
Laboratory, Ecological Research Division, 26 W. Martin Luther King Dr., Cincinnati, OH 45268
187
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30-second, 0.26-m2 (2.8 ft2) kick samples are collected in the near-shore littoral zone using a
standard rectangular-frame kick net (335 x508-mm frame; 500-um mesh). The 22 kick
samples at each site are combined into a single composite sample representing a total bottom
area of 5.7 m2. Sampling is restricted to the littoral habitat because deeper benthic habitats of
the channel are much more difficult to sample, and benthic organisms are often present in very
low abundance in non-littoral channel areas- especially in large, sand-bed rivers. Table 10-1
describes the kick sampling procedures in detail.
In some reaches that are extensively modified with wing-dams, spur-dikes, or other
channel-training structures, vertical shorelines may prohibit safe kick sampling at some littoral
stations. When this situation is encountered, search 5 m up- and down-river from the station for
safe kick sample locations. If no safe location is available, do not collect kick samples at that
station and note the missing samples with a flag on the form. Littoral sample locations in dike
fields are located along the wetted edge of the natural shoreline contour unless the littoral
station occurs opposite of the base of a dike (see Figure 10-1).
10.2 Snag sampling
In alluvial floodplain rivers, snags are massive pieces of large woody debris (LWD) that
are imbedded in or resting on the river bottom (Angradi et al., 2004). The snag sample (Figure
10-2) is collected from the snag that is nearest to the intersection of the cross-channel and the
MCS and which meets the suitability criteria. A specialized "snag net" is used to collect a 1-m-
long sample from the up-current side of the snag. The snag net resembles a standard
rectangular-frame kick net, but with the frame constructed so that the net fits over half the
circumference of a snag (Figure 10-3). Two sizes of snag nets with mouth widths ("diameters")
of 0.2 m (8 inches) and 0.33 m (13 inches) will be used. For larger snags, a standard
rectangular-frame kick net (33 x 51-cm frame; 500-um mesh) can be used to sample the
surface of the snag. Snag sampling methods are described in Table 10-2.
188
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River let bankline
Flow
I
Littoral location
Actual sample
Dike
Vertical bank
likely
Figure 10-1. Guidance on sample location for littoral stations in dike fields. Littoral stations
should follow the wetted edge of the natural contour of the shoreline unless the
station occurs at the base of a dike. In this case, the sample is collected on the
face of the dike. Scalloped shorelines behind some dikes may have
unsampleable vertical banks.
Suitable snags for sampling are in water at least 0.6 m deep, are exposed to some
current, and are at least 5 m long, the minimum length for a piece of LWD established for
EMAP (Peck et al., unpublished drafts). Suitable snags are at least 15 cm (6 inches) in
diameter where the snag breaks the surface or comes within 30 cm (12 inches) of the water
surface. This definition of a suitable snag is restrictive to minimize among-sample variability and
maximize the efficiency of the snag nets. Finding the nearest suitable snag may require some
searching up- and down-river from the X-site. Surrogate snag samples may be collected
189
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from man-made structures (pilings, navigation markers, etc.) if no natural snags are
present. Ancillary data collected for each snag include depth, snag diameter at the water
surface, snag surface characteristics, water velocity, and the distance of the snag from the
shore. Take a picture of the sampled snag if possible.
Figure 10-2. Snag sampling on the Missouri River. The captain is holding the boat in position
against the snag using the motors. The biologist at left is brushing the surface of
the snag in front of the snag net.
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10.3 Large woody debris abundance
At each site, large woody debris (LWD) meeting the minimum size criteria is tallied for a
500 m x 10 m near-shore quadrat adjacent to the MCS transect. This procedure is included in
this section of the manual because the LWD is most efficiently tallied while searching for a
suitable snag sampling location. Nearshore LWD is categorized by type, after Angradi et al.
(2004), as below the wetted perimeter ("wet"), between the wetted perimeter and the bankfull
stage but not having originated a its present location ("beached"), and partly below the wetted
perimeter (providing aquatic habitat) but originating at the present location ("anchored"). A
piece of LWD that is anchored may also be wet because it provides aquatic habitat, but it
should be counted as "anchored." Anchored LWD that is not providing aquatic habitat is not
counted. Density of LWD (number per hectare of channel) in the main channel including the 10-
m-wide near-shore quadrat is visually estimated and recorded by density category. EMAP-GRE
includes fewer LWD size categories than previous EMAP methods (Kaufmann 2003). Our
experience on the Upper Missouri River in 2000-2003 showed that most LWD was < 1 m
diameter at the large end and £ 20 m in length. Furthermore, accurate estimation of piece
length is difficult for LWD in the channel because usually only a portion of the LWD is visible.
Procedures for quantifying LWD are described in Table 10-3.
10.4 Sample processing
After the composite kick or snag samples have each been distributed into one or more
jars (with no jar filled more than 1/3 full with sample), the jars should be filled almost completely
with 10% carbonate-buffered formalin. Each jar is then topped of with a concentrated rose
bengal solution (Table 3-1). A label is filled out (Figure 10-6) and placed inside each jar and on
the outside of each jar. If extra jars are needed, transfer the sample ID to a continuation label
and place it on the outside of each jar. Each sample composite (not each jar) gets a unique
sample ID. Details of the sample processing procedure are provided in Table 10-4.
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Table 10-1. Procedures for collecting near-shore littoral kick samples (adapted, in part,
from Klemm et al. 2003).
1. Fill in the site ID and date on the littoral and snag sampling form (Figure 10-4). Go to MCS
station A at the intersection of the cross-channel transect and the MCS (Figure 4-1 and 4-2)
and record the latitude and longitude at the intersection using a hand-held GPS unit.
2. At each of 11 evenly-spaced stations on the MCS transect (i.e., station A at Om, station B at
50 m, etc.; Figure 4-2) collect two kick samples. Locate stations that have not been pre-
flagged (50 m, 150 m, 250 m, etc.) using a hand-held GPS or by estimating the halfway point
between flagged stations.
3. Locate kick samples in a zone bounded on the shore side by the apparent low-water mark from
daily flow fluctuations (most relevant on the regulated Upper Missouri River) and bounded on
the river side by the 0.6-m depth contour (recommended maximum sample depth; deeper
sampling may be possible). The low-water mark at a site can often be detected by the
presence of periphyton or attached filamentous algae just below the low water mark. If
samples cannot be safely collected at a station due to vertical banks or other reason, search 5
m up- and down-river for a safe location. If a safe location is still not available, do not collect
kick samples at that station and note the missing sample for the station with a flag on the form.
4. If there is sufficient current to extend the net, go to step 5; if not go to step 16.
5. Method for kick sampling in current. At each sample location, hold the net opening facing
up-current, and position the net securely on the stream bottom. Avoid large rocks or debris that
prevent the net from seating properly on the bottom.
6. Visually define a square quadrat on the bottom just up-current from the net that is one net-width
on every side (0.26 m2). Check the quadrat for heavy organisms
the bottom is visible). Place these organisms in the net by hand.
on every side (0.26 m2). Check the quadrat for heavy organisms such as mussels and snails (if
7. Holding the net in place with your knees, pick up any loose cobbles or large pieces of gravel or
debris from the quadrat and rub them with your hands or a small brush so that organisms wash
into the net. Discard the rocks outside the quadrat. In Waterloo deep to brush rocks in front of
the net, place the rocks in the sample composite bucket.
8. After scrubbing and removing the larger substrate particles, hold the net securely in position
while stirring the substrate remaining within the quadrat to a depth of 10 cm for 30 seconds
using either a gloved hand, or by kicking the substrate vigorously for 30 seconds.
9. Remove the net from the water with a quick upstream motion to wash the organisms to the
back of the net.
Continued
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Table 10-1. Procedures for collecting near-shore littoral kick samples, continued.
10. Kicking in a sand bottom can result in a very full net. Most of the sand will pass through the
mesh if the net is gently agitated while the net mouth is held out of the water with one hand and
the collecting bucket supported with the other hand.
11. Sweep the net through clean water several times to consolidate net contents in the screened
bucket at the cod end. Inspect the net for clinging organisms; using forceps, place any
organisms found into the Dolphin bucket. Dump the contents of the Dolphin bucket into a
sample composite bucket. In some cases it may be faster and easier to hold the first sample
from a station in the net while the second kick sample is being taken.
12. On the form, circle the appropriate dominant substrate size/type at the sample location.
13. Repeat steps 5-11 for a second kick sample location at the station. Be sure to move upstream
at least 1 m before collecting the second kick sample.
14. Repeat steps 5-13 for the remaining littoral sample stations.
15. Go to step 23.
16. Method for sweep sampling in slack water or pools. At each sample location, hold the net
opening facing upstream, position the net securely on the stream bottom. Avoid rocks or
debris that prevent the net from seating properly on the bottom.
17. Visually define a square quadrat on the bottom just up-current from the net that is one net width
on every side (0.26 m2). Check the quadrat for heavy organisms
the bottom is visible). Place these organisms in the net by hand.
on every side (0.26 m2). Check the quadrat for heavy organisms such as mussels and snails (if
18. Holding the net in place with your knees, pick up any loose cobbles or large pieces of gravel
and debris and place them in the sample composite bucket.
19. Vigorously kick the substrate remaining within the quadrat and then drag net through the
disturbed area just above the bottom. Continue this for 30 seconds (counting "one Missouri,
two Missouri, etc." is sufficiently accurate). Keep the net moving so captured organisms cannot
escape.
20. Remove the net from the water with a quick upstream motion to wash the organisms to the
back of the net.
21. Sweep the net through clean water several times to consolidate net contents in the Dolphin
bucket at the cod end. Inspect the net for clinging organisms; using forceps, place any
organisms found into the Dolphin bucket. Dump the contents of the Dolphin bucket into the
composite sample bucket.
22. Repeat steps 16-21 for a second kick sample location at the station. Be sure to move
upstream at least 1 m before collecting the second kick sample.
Continued
193
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Table 10-1. Procedures for collecting near-shore littoral kick samples, continued.
23. On the form, circle the appropriate dominant substrate size/type at the sample location.
24. Repeat steps 15-23 for the remaining sweep sample replicates in slack water areas.
25. Dump the contents of the composite bucket, including both "current" and "slack water"
samples, into a 0.3-m diameter 500-um mesh sieve and wash the sample gently (no nozzle)
using the onboard washdown hose (the 0.3-m diameter sieve will fit over a 20-L bucket to catch
wash water). Gravel, large organic particles, and macrophytes should be thoroughly washed,
inspected for clinging organisms and discarded. If there is a large amount of coarse sand or
small gravel in the sample, use a second 20-L bucket to elutriate the sample before sieving.
26. Transfer the washed composite sample into a 500-mL jar using the wide-bore funnel and the
wash bottle with minimal water. Use two or more jars if necessary. Do not fill any jar more
than 1/3 full with sample.
27. For sample jars that are not pre-labeled, the site number and sample type should be written
directly on the jar(s) with a Sharpie (e.g., "kick 045") to avoid mixing up jars. This notation
should be covered later with the sample label. Place the samples in a cooler.
28. Go to Table 10-4 for procedures for labeling and preserving samples.
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10.5 QA considerations for macroinvertebrate sampling
Standardization of effort and attention to detail are important for maintaining a high QA
standard for field sampling. Several QA considerations for macroinvertebrate sampling are
presented in Table 10-5.
10.6 Safety considerations for macroinvertebrate sampling
Safety is paramount. General and boat-related safety guidance is presented in Section
2. Safety considerations relevant to macroinvertebrate sampling are presented in Table 10-6.
10.7 Equipment and supplies
Table 10-7 is a checklist of equipment and supplies required for collecting
macroinvertebrate samples. Generic supplies required for all EMAP-GRE field sampling are
listed in Table 2-5.
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Table 10-2. Procedure for collecting snag macroinvertebrate samples.
1. Find the natural snag nearest to the intersection of the cross-channel transect and the MCS
(littoral sample station A, Figure 4-2) which meets the snag suitability criteria. Search both
shorelines 1 km up-river and down-river from the transect and on the way back to the ram p. If
no suitable natural snag is found, a man-made snag substitute should be sampled (e.g., piling).
If a snag is sampled that does not meet all the natural snag criteria in step 2, note this in the
comments on the littoral and snag sampling form (Figure 10-4).
2. The snag must be in flowing waters at least 0.6 m (2 ft) deep, must be >j5 m (16.5 ft) long, and
with a diameter of >. 0.15 m (6 inches) where the snag breaks the water surface or comes
within 0.3 m (1 ft) of the water surface. Select the proper snag net for the snag (small [0.20 m
"diameter"] or large [0.35 m "diameter"]). For snags too large for the large snag net, use the
rectangular kick net (Figure 10-3).
3. Navigate up to the snag. Approach the snag slowly. The boat driver should be able hold the
boat in position using the motor(s) (Figure 10-2).
4. Place the net against the snag facing up-current just below where the snag breaks the surface
or where the snag comes closest to the water surface. If debris is wrapped around the snag at
the water surface, sample further down (up current) on the snag.
5. A second crew member should use a long-handled brush to scrub the snag to wash organisms
into the net in a traveling sample down the snag (Figure 10-2). Attempt to sample M m of
snag. Be sure to scrub the sides of the snag. This is a qualitative method; not all the organisms
on the snag will be captured in the net.
6. Sweep the net through clean water to consolidate net contents in the screened bucket at the
cod end. Inspect the net for clinging organisms; place any found into the screened bucket using
a forceps.
7. Transfer the contents of the Dolphin bucket directly into a 250-mL jar and rinse organisms from
the Dolphin bucket into the jar using a wash bottle and a minimum of water. Sieving the sample
will probably be unnecessary. If sieving is necessary to reduce sample volume, use the
procedures in Table 10-1, step 25. Use two jars if the sample fills the jar more than 1/3 full.
8. Record the depth under the sampled part of the snag using the boat's sonar (Figure 10-5).
Estimate the snag diameter size class in cm at the water surface (or where the snag comes
closest to the surface). Record the snag surface characteristics (e.g., smooth, rough, algae
present). Determine the distance from the snag to the nearest shoreline using a laser
rangefinder or other method (visual estimation is acceptable). Record the net used and the
approximate length of the snag sample (1 m is the goal). Record the coordinates of the snag.
Continued
196
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Table 10-2. Procedure for collecting snag macroinvertebrate samples, continued.
9. Use a velocity meter to measure surface water-velocity (m/s) above the sampled area.
10. Alternatively, navigate several boat lengths up-river of the snag, shift to neutral and allow the
boat to drift past the snag as closely as possible and record the speed-over-ground from the
boat-mounted GPS unit as the boat passes the snag. A speed-over-ground of 1 km/h ~ 0.28
m/s (1 mile/h = 0.45 m/s). This method does not work if it is windy.
11. For sample jars that are not pre-labeled, the site number and sample type should be written
directly on the jar(s) with a Sharpie (e.g., "snag 045") to avoid confusion later. This notation
should be covered later with the sample label. Place the samples in a cooler.
12. Go to Table 10-4 for procedures for labeling and preserving samples.
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Table 10-3. Procedures for quantifying large woody debris (LWD).
1. LWD is defined as pieces >. 5 m long and >. 0.3 m in diameter at the large end. Pieces may or
may not break the water surface. For LWD in deep water (also called snags), the diameter of
the large end of the piece will have to be estimated. In most cases snags in the main channel
that are exposed to current are necessarily massive (or they would not be there) and will have
a large-end diameter > 0.3 m. For pieces that are not cylindrical at the large end, visually
estimate what the diameter would be for a cylindrical piece of the same volume.
2. Near shore LWD tally. While cruising along the target shoreline in the boat (or while on foot if
there is a lot of LWD), tally each piece of LWD by type: "wet," "beached," or "anchored" in the
10 x 500 m littoral quadrat on the littoral and snag sampling form. Wet LWD is in the channel
below the wetted perimeter. Beached LWD is between the wetted perimeter and the visually-
estimated bankfull level. Anchored LWD is partly below the wetted perimeter, but is anchored in
the bank where it originated. Anchored LWD is typically a tree that has been undermined by
bank erosion and has toppled over into the river but which has not been washed away. Beaver-
felled trees often become anchored LWD. LWD that is anchored but does not provide
aquatic habitat is not counted. For log jams in which some pieces may not be visible,
attempt to estimate the number of pieces. Sum the tally and record the total for each type on
the form (Figure 10-5).
3. Channel LWD. Note the presence of LWD in the channel outside the littoral quadrat. Estimate
the density of LWD as number/hectare in the channel between the main channel banks for the
500-m MCS transect (including the wet LWD from the littoral transect). A hectare is equal to an
area of 100 x 100 m. Mark the appropriate category on the form.
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Table 10-4. Procedures for labeling and preserving macro in vertebrate samples.
1. To avoid clutter in the boat, benthos samples can be transported to the ramp or base location
(if it is close to the ramp) in a cooler to be preserved.
2. Fill each jar almost to the top with 10% carbonate-buffered formalin. Top off each jar with
concentrated rose bengal solution (Table 3-1). Prepare a label(s) (Figure 10-6) for inside each
jar(s). Circle the sample type (kick or snag); fill in the site number; enter the sample date, and
print the collectors name. Place the label(s) inside the jar(s).
3. Cap the jar and gently invert and rotate the jar to distribute the preservative. The preservative in
the sample jar should be pink.
4. Prepare a label (Figure 10-6) for the outside of the jar. Circle the sample type (kick or snag);
fill in the site number from the design file; enter the sample date, visit number, jar number and
total number of jars. Place the label on the jar and cover it with clear tape. Record the total
number of jars and the sample ID from the label on the littoral and snag sampling form.
5. If the sample requires more than one jar use a continuation label (Figure 10-6). Use the sample
ID number from step 4.
6. Seal each jar with plastic electrician's tape by wrapping with the threads (clockwise). Store the
preserved sample upright in a secondary container until transport or shipment to the laboratory.
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Table 10-5. QA considerations for macroinvertebrate sampling.
Attempt to expend equal effort for each kick sample replicate despite variation in current and
substrate.
Strive to avoid any bias when locating kick samples.
A few drops of non-permanent "thread-lock" on the kick-net ferule threads will prevent the
handle from twisting during sampling.
Use a gentle wash (no nozzle) when sieving the sample to avoid damaging fragile organisms
(e.g., worms).
Avoid sampling when a barge is approaching.
Always inspect the kick or snag net for invertebrates clinging to the mesh.
Try to get a natural snag or snag substitute (e.g., piling, channel marker, floating dock) sample
at every site.
Remove large organic debris from samples and drain excessive water from jars before adding
preservative to insure that final preservative strength is sufficient to preserve organisms.
Table 10-6. Safety considerations for macroinvertebrate sampling.
Use extreme care walking on rip rap. Rocks can shift unexpectedly and serious falls are
possible.
Use caution when kick sampling in swift or deep water. Wear a suitable PFD and consider
using a safety tether held by an assistant. For most people, conditions are rarely suitable for
collecting a good kick sample in water deeper than 0.6 m.
Do not attempt to kick sample vertical or near-vertical banks.
Professional-quality breathable waders with a belt are recommended for kick sampling.
Neoprene booties are an alternative, but should have sturdy, puncture-resistant soles.
Avoid wet-wading in areas down-river from effluent discharge points.
Use caution approaching and sampling snags. Good communication between the crew and
captain is essential to avoid grounding and injury.
• Use safety glasses and gloves when handling formalin.
200
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Table 10-7. Equipment and supply checklist for macroinvertebrate sampling. Generic
supplies required for all EMAP-GRE field sampling are listed in Table 2-6.
Qty
1
2
1
1
1
1
2-3
1 roll
1
2pr.
1
1
1
1
at least 6
at least 4
1 roll
1 set
at least 3 L
1 L
1
Item
Modified rectangular kick net ("also called Slack Sampler") with 500-|jm mesh
and handle (e.g., Wildco 425-M53 or equivalent)
300-mL Dolphin plankton bucket with 500-um mesh (e,g,. Wildco 47-D60 or
equivalent)
0.20-m diameter snag net with 500-um mesh and 200-mL Dolphin bucket
(Wildco 424-C56 or equivalent)
0.33-m diameter snag net with 500-um mesh and 200-mL Dolphin bucket
(Wildco 424-A56 or equivalent)
US standard 35 sieve (500-um mesh) 30-cm diameter, stainless-steel mesh
Wash bucket with (500-um mesh) (e.g., Wildco 190-E25) (optional)
20-L plastic bucket for transporting composite between stations and catching
waste water during when washing the sample
Biodegradable flagging (different color from tape use to lay out the site)
Velocity meter
Forceps for removing invertebrates from nets
1-L wash bottle
Long-handled scrub brush for snag sampling ("deck brush" style works best)
Small brush for kick sampling
Large bore funnel for transferring samples from sieve to jar
HOPE sample jars, wide mouth, leakproof, screw top, 1 L capacity (for kick
samples)
HOPE sample jars, wide mouth, leakproof, screw top, 250-mL capacity (for
snag samples) (Fisher Scientific 03-31 1-3D or equivalent)
Plastic electrician's tape for sealing sample jars
Labels
10% carbon ate -buffered formalin
Concentrated rose bengal solution (see Table 3-1)
Littoral and snag sampling form
201
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10.8 Literature cited
Angradi, T.R., E.W. Schweiger, D.W. Bolgrien, P. Ismert, and T. Selle. 2004. Bank stabilization,
riparian land-use, and the distribution of large woody debris in a regulated reach of the
Upper Missouri River, North Dakota, USA. River Research and Application 20:1-18.
Barbour, M.T., J. Gerritsen, B.D. Snyder, and J.B. Stribling. 1999. Rapid Bioassessment
Protocols for Use in Streams and Wadeable Rivers: Periphyton, Benthic
Macroinvertebrates and Fish, Second Edition. EPA/841/B-99/002. U.S. Environmental
Protection Agency, Office of Water, Washington, DC.
Http://epa.gov/OWOW/monitoring/techmon.html
Klemm, D.J., P.A. Lewis, F. Fulk, and 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, Washington, DC.
Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and
Monaco, P. A. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Non-Wadeable
Rivers and Streams, U.S. Environmental Protection Agency, Washington, DC.
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Ringold, P. L., Magee, T. and
Cappaert, M. R. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams,
U.S. Environmental Protection Agency, Office of Research and Development,
Washington, DC.
202
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Figure 10-3. Snag and kick nets. The two sizes of snag nets are 0.20 m (left) and 0.33 m
diameter (middle). For snags >0.33 m in diameter, the standard D-frame kick net
(right) is used.
203
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LITTORAL AND SNAG SAMPLING FORM
Draft
SITE 10: GRW04449-
DATE:
/ 2 0 0
ANNUAL
NUMBER;
Dominant Substrate at Kick Sample Location (Distance from start of transect in parentheses)
Substrate
Flag
Substrate
Flao.
A
(0)
XB SB CB OC OF SA FM WD HP OT
G
(300)
XB SB CB OC OF SA FN WD HP OT
B
(50)
XB 88 OB OC Of SA Ft) WD HP OT
XB SB CB SC OF SA FN WO HP OT
C
(100)
JCB SB CB OO OF SA FM WD HP OT
(440)
XB SB CB OC OF SA FN WD HP OT
XB SB CB SC 6F SA Ft) WO HP OT
J
|4iO|
XB SB CB GC QF SA FN WO HP OT
£
(200)
XB SB CB OC OF SA FN WD HP OT
(5001
XB SB CB GC OF SA FN WD HP OT
F
(258)
XB SB CB OC OF SA FN WO HP OT
Total numter of kick samples:
Flag:
CODE
SEE CLASS
SIZE RANGE (mm)
DESCRIPTION/COMMENTS
XB
SB
CB
GC
GF
SA
FN
WO
HP
OT
Large boulders
Small boulders
Cobbles
Gravel (coarse)
Gravel (fine)
Sand
Fines
Wood
Hard pan
Other
>1000
>250to1000
>64 to 250
>16to64
>2to16
>0.06 to 2
<0.06
Any size
Includes some riprap
Basketball size and larger; Includes some riprap
Tennis ball to basketball size; Includes some rlpra
Marble to tennis ball size
Ladybug to marble size
Gritty between fingers
Silt. day. myck, not gritty between fingers
Describe in comments
Firm, consolidated fine sybstrate. packed clay
Describe in comments
Composite Kick Sample
Sample ID
No. of
Jars
Comment
Periphyton
Composite
Sample
Sample ID
Comp.
Vol. ImL)
n Stations
Sampled
No. of
Jars
Comment
Sediment
Composite
Sample
Sample ID
Comp.
Vol. (L)
* Stations
Sampled
Commenl
LWD Tally (Record total in small box)
LWD - > 5 m long >. G.3 m diameter at larger end
Wet LWD
Beached LWD
Anchored LWD
Flag
0.3 • 0.6 m
(12-24")
> 0.6 B)
(> 24")
Channel LWD
Density
(pieces/ha)
None
Q2-S
Flag
Ftafl codes: K=no made, U=syspect measyrement; F1. F2, etc=mlsc by field crew. Explain In comments.
32.
Figure 10-4. Littoral and snag sampling form (front).
204
-------
EMAP-GRE LITTORAL AND SNAG SAMPLING
Draft
SITE ID: GRW04449-
DATE:
1200
ANNUAL VISIT n 1 n 7
NUMBER:
GPS Coordinates of Snag
Snag Characteristics
Latitude
Net Used
Flag
Length of
Sample jm)
Flag
Snag
Surface
Flag
Longitude
SS LS KN
Depth
Flag
Flag
Dist, from
Shore (m)
Flag
Velocity
m.'s
Flag
Snag Diameter
D 18 • m cm D -15 • m cm
n 2S - 3i em n > 5* cnt
D 35 - 45 cm
Flag
Snag Surface Codes
SB s Srftooth, no algae
SA = Smooth, algae
RB ~ Rough, no algae
RA = Roygh, algae
BB = Bark on. no algae
BA - Bark on. algae
Velocity Conversions
t K/h = 6.28 m/s
! fttlto/ft K 0.45 m/s
Snag nets
SS = Smail snag
LS * Large Snag
KM = Kick Net
Snag Sample
Sample ID
No. of Jars
Comment
Flag
Comment
Flag codes: K=no measurement made, U=suspect measurement; F1, F2, ete=miise flags assigned by field crew. Explain in comments.
33.
Figure 10-5. Littoral and snag sampling form (back).
205
-------
BENTHOS
BK BS
/
1 2
300011
Jar of
BENTHOS
BK BS
/
Site visit number 1 2
ID
Jar of
BENTHOS
IK 3S
/
Collector
Figure 10-6. Labels for benthic macro!nvertebrate samples. BK = kick sample; BS = snag
sample. The label at upper left would be affixed to the outside of a jar. The
continuation label at upper right is used if more than one jar is needed. The
bottom label would be placed inside each jar. Not actual size.
206
-------
Section 11
Periphyton and Sediment
Brian H. Hill1 and James M. Lazorchak2
Periphyton includes algae, fungi, bacteria, protozoa, and associated organic matter on
the surface of aquatic substrata. Periphyton assemblage composition is a useful indicator of
environmental condition because it responds rapidly to a number of anthropogenic
disturbances, including habitat alteration, excess nutrients, metals, herbicides, hydrocarbons,
and acids (Pan et al. 1996, Hill et al. 2003).
Benthic organisms are in intimate contact with river sediments. Benthic assemblages
are influenced by the physical and chemical properties of sediment. Sediment characteristics
serve as exposure indicators for benthos, fish, and other wildlife (e.g., sediment toxicity) and as
functional indicators of key ecosystem processes (e.g., sediment enzyme activity) (Sinsabaugh
and Foreman 2001, Hill et al. 2002). Periphyton and sediment collection methods described
herein are adapted from previous EMAP methods for wadeable and non-wadeable streams
(Peck et al., unpublished drafts).
11.1 Periphyton sample collection
At each site, a 500-m main channel shoreline (MCS) transect, starting at the intersection
of the cross-channel transect and the MCS (Figure 4-1), is laid out by either the fish- or river-
sampling crew, depending on which crew arrives at the site first. The primary transect is initially
flagged at 100-m intervals; intermediate littoral stations at 50-m intervals are located and
flagged using a handheld GPS or by visual estimation during littoral sampling (use a different
flag color than at the 100-m stations). At each of 11 evenly-spaced stations along the 500-m
MCS transect (every 50 m; Figure 4-2), a 25-cm2 littoral periphyton sample is collected from
the dominant hard substrate at the station. If no hard substrates are present, fine substrates
1 U.S. Environmental Protection Agency, Office of Research and Development, National Health and
Environmental Effects Laboratory, Mid-Continent Ecology Division, 6201 Congdon Blvd, Duluth, MN 55804
2 U.S. Environmental Protection Agency, Office of Research and Development, National Exposure Research
Laboratory, Ecological Research Division, 26 W. Martin Luther King Dr., Cincinnati, OH 45268
207
-------
are sampled. For guidance on where to collect littoral samples in a dike field, see Figure 10-1.
Table 11-1 describes the periphyton sample collection procedures in detail.
11.2 Sediment sample collection
At or near each of the 11 periphyton sampling locations, a fine-sediment sample is
collected using either a hand scoop or a "petite Ponar" grab sampler. The objective is to collect
a 4-L composite sample that is representative of MCS depositional areas at the site. The
composite sample will be subsampled in the lab for multiple analyses. Table 11-2 describes the
sediment sample collection procedures in detail.
11.3 QA Considerations for periphyton and sediment sampling
Standardization of effort and attention to detail are important for maintaining a high QA
standard for field sampling. Several QA considerations for periphyton and sediment sampling
are presented in Table 11-4.
11.4 Safety considerations for periphyton and sediment sampling
Safety is paramount. General and boat-related safety guidance is presented in Section
2. Safety considerations relevant to periphyton and sediment sampling are presented in Table
11-5.
11.5 Equipment and supplies
Table 11.6 is a checklist of equipment and supplies required for collecting periphyton
and sediment samples. Generic supplies required for all EMAP-GRE field sampling are listed in
Table 2-5.
208
-------
Table 11-1. Procedure for collecting periphyton samples.
1. Go to MCS station A at the intersection of the cross-channel transect and the main channel
shoreline (Figure 4-2). Periphyton is collected at locations that correspond to
macroinvertebrate kick sample locations (see Section 10).
2. Locate periphyton samples in a zone bounded on the shore side by the apparent low-water
mark from daily flow fluctuations (most relevant on the regulated Upper Missouri River) and
bounded on the river side by the 0.3-m (about mid-biceps) depth contour. This is the
recommended maximum sample depth. The low-water mark at a site can often be detected by
the presence of periphyton or attached filamentous algae just below the low-water mark. If
samples cannot be safely collected at a station due to vertical banks or other reason, search 5
m up- and down-river for a safe location. If a safe location is still not available, do not collect a
periphyton sample at that station and note the missing sample for the station with a flag on the
form.
3. At each station, select a piece of hard substrate (coarse gravel, cobble, wood) that can be
easily removed from the river bottom (usually <15 cm diameter). If substrate is heterogenous,
select an example of the dominant substrate type at the station. Be sure to avoid the area that
has just been kick sampled. Sampling just upriver from the kick sample location is
recommended.
4. Place the substrate in a large funnel draining into a 500-mL sample composite bottle. Brush a 5
x 5 cm (2 x 2 inch) area on the upper surface of the substrate using a stiff tooth brush (supplied
by EPA). Use of a delimiter or template is not required. Rinse the loosened periphyton and
toothbrush bristles into the funnel using filtered river water in a wash bottle. Discard the
brushed substrate.
5. If the substrate can be removed from the water but not held over the funnel (e.g., LWD), use a
knife blade to scrape a 5 x 5 cm (2 x 2 inch) area from the upper surface and wash the knife
blade into the funnel.
6. If there are no hard substrates at the sample station, use a 60-mL syringe to vacuum up 25 cm2
of fine substrate (silty sand, silt, clay, muck) to a depth of approximately 2 cm. Alternatively,
press a small petri plate into the substrate and slip a spatula under it and remove it from the
substrate. In deep areas, this type of sampling might not be feasible. If no periphyton sample
can be collected at a station, flag the data and note the reason in a comment on the littoral and
snag sampling form (Figure 10-4).
7. If there are no sufficiently large substrates for a brush sample, or sufficiently fine substrates for
a syringe sample (e.g., if the station is all hardpan or fine gravel), search 5 m up- and downriver
for the nearest suitable substrate (coarse or fine) and sample it. If there are still no suitable
substrates, do not collect a periphyton sample at that station and flag the data on the and note
the reason in a comment on the littoral and snag sampling form.
Continued
209
-------
Table 11-1. Procedure for collecting periphyton samples, continued.
8. Repeat steps 2-7 at each of the 11 MCS stations. Record the total number of replicates
(stations sampled) included in the composite.
9. Fill the composite jar 2/3 full with river water, cap it, and place it in a cooler with ice (keep the
sample dark until it is preserved). For sample jars that are not pre-labeled, the site number and
sample type should be written directly on the jar with a Sharpie (e.g., "peri 045") to avoid
confusion later. This notation should be covered later with the outside of the jar sample label.
10. Go to Table 11-3 for procedures for labeling and preserving samples.
210
-------
Table 11 -2. Procedure for collecting sediment samples.
1. At each of 11 evenly-spaced stations on the MCS (i.e., 0, 50, 100m from the down-river end of
the transect; Figure 4-2), that correspond to macroinvertebrate kick sample locations (see
Section 10), collect a sediment sample.
2. Locate sediment samples in areas or patches of fine substrate (silty sand, silt, clay, muck) in a
zone bounded on the shore side by the apparent low-water mark from daily flow fluctuations
(most relevant on regulated Upper Missouri River) and bounded on the river side by the 0.3-m
(usually about mid biceps) depth contour (recommended maximum sample depth; deeper
sampling may be possible). The low-water mark at a site can often be detected by the
presence of periphyton or attached filamentous algae just below the low-water mark. If samples
cannot be safely collected by wading at a station due to vertical banks or other reason go to
step 5.
3. Be sure to avoid the area that has just been kick sampled. Sampling up-river from the kick
sample location is recommended. If fine substrates are not present within 5 m up- or downriver
from the station, do not collect sediment at that station and flag the station on the form.
4. If fine substrate is present, use a small scoop to collect a sample of about 225 cm2 (= 15 x 15
cm [6x6 inches] ) of the top 2 cm of substrate (this volume is approximately equal to six
scoops). Place the sample in a clean bucket. Use gloves for handling sediment. Do not
assume rip rapped shorelines lack fine sediment. Look for fines between the large rocks
5. If wading is not possible, use a petite Ponar sampler or similar device deployed from the boat
to collect a sediment sample adjacent to the station. Release the petite Ponar sample onto a
tub and use the scoop to collect about 225 cm2 (=15 x 15 cm [6 x 6 inches] ) of the top 2 cm of
the sample. Estimate sample area visually. Place the subsample in the sediment composite
bucket and discard the rest of the Ponar sample.
6. Repeat steps 2-5 at each of the 11 littoral stations. Record the total number of replicates
(stations) included in the composite. Note in a comment the stations at which sediment was
collected using a non-wading method.
7. It is important that a sufficient sediment (not less than 4 L) sample for analysis be collected. If
multiple stations have no fine sediment, it is permissible to collect extra sample at stations that
do have fine sediment or between stations. Be sure to note this in a comment.
8. Using a large stainless steel spoon, thoroughly mix the composite sample in the bucket and
transfer 4 L of the composite in a 30 x 50-cm 3-mil thick polyethylene bag. Try to limit the
amount of sediment adhering to the inside of the bag near the top. Grasp the bag just above
the sediment to express the air. Twist and knot the bag to seal it. Write the site number and
date directly on the bag with a permanent marker and place it in a cooler with ice.
9^ Go to Table 11-3 for procedures for labeling and preserving samples.
211
-------
Table 11-3. Procedures for labeling and preserving periphyton and sediment samples.
1. To avoid clutter in the boat, periphyton and sediment samples may be transported to the ramp
or base location (if it is close to the ramp) in a cooler with ice for final labeling and preservation.
2. Periphyton. Add 20 ml_ of 100% borax-buffered formalin to the 500-mL periphyton composite,
and top off the bottle with river or tap water (final formalin concentration of 4%). Prepare a label
(Figure 11-1) for outside the jar. Using a fine-point permanent marker, fill in the site number,
the sample date, and total preserved sample volume. Place the label on the jar and cover it
with clear tape. Record the sample ID and other data on the littoral and snag sampling form
(Figure 10-4)
3. Seal each periphyton jar with plastic electrician's tape by wrapping with the threads (clockwise).
Store the preserved samples upright in a secondary container to await transport or shipment to
the laboratory.
4. Sediment. Place the sediment sample inside a second 3-mil polyethylene bag, twist the top,
and knot to seal. Prepare a label (Figure 11-1) for outside the outer bag. Using a fine-point
permanent marker, fill in the site number and sample date. Place the label on the outer bag
and cover it with clear tape. Record the sample ID and other data on the littoral and snag
sampling form (Figure 10-4). Place the sample on ice or in a refrigerator. Do not freeze
sediment samples.
Table 11 -4. QA considerations for periphyton and sediment sampling.
Try to make sure each of the 11 periphyton and sediment subsamples comprises an
approximately equal portion of the total composite.
It is permissible to collect sediment between stations to insure a composite volume of at least
4L. Note deviations from standard procedure in a comment.
Do not assume rip rapped shorelines lack fine sediment. Look for fines between the large
rocks.
Mix the composite sediment sample thoroughly before extracting the final 4L composite.
When sampling LWD pulled from the river, be sure to sample the upper surface of the LWD.
• Monitor sediment samples in your possession to insure they do not warm up or freeze.
212
-------
Table 11-5. Safety considerations for periphyton and sediment sampling.
Use extreme care walking on riprap. Rocks can shift unexpectedly and serious falls are
possible.
Use caution when sampling in swift or deep water. Wear a suitable PFD and consider using a
safety tether held by an assistant. For most people, conditions are rarely suitable for collecting
a periphyton or sediment sample in water deeper than 0.6 m.
Do not attempt to collect periphyton or sediment from vertical or near vertical banks.
Professional-quality breathable waders with a belt are recommended for littoral sampling.
Neoprene boots are an alternative, but should have sturdy, puncture-resistant soles.
Use caution using the Ponar-type samplers. The jaws are sharp and may close unexpectedly.
Replace frayed lines and worn parts.
Raise the Ponar sampler from and into a plastic tub rather than from the boat deck. This
prevents feet from getting under the sampler.
Don't try to remove large pieces of LWD from the river by yourself.
Use safety glasses and gloves when handling formalin.
Avoid contact with sediment samples. Use gloves if necessary.
213
-------
Table 11 -6. Equipment and supply checklist for periphyton and sediment sampling.
Generic supplies required for all EMAP-GRE field sampling are listed in Table 2-
5.
Qty
1
1
1
1
1
1
1
1
1
1
2
1 L
1
1 roll
1 set
1
Item
Petite Ponar sampler (Wildco 1728-G30 or equivalent) with plastic tub, drop line, and
spare pinch pin. A standard Ponar or similar device may substitute.
Stiff-bristle tooth brush for brushing periphyton (Wildco 1 56-F40 or equivalent)
Knife for scraping LWD
60-mL syringe with 1-cm (3/8") hole bored into the end for sampling fine substrate
Scoop for collecting sediment
Graduated plastic bucket
Large stainless steel spoon for mixing sediment composite
1-L wash bottle
Large bore funnel with > 20-cm wide-opening
HOPE sample jars, screw top, 500-mL capacity for periphyton samples (Fisher
Scientific 03-31 1-3E or equivalent)
30 x 50-cm, 3-mil polyethylene bags for sediment samples (Aquatic EcoSystems FSB5-
10 "x20" or equivalent)
Borax-buffered formalin (100%)
Pipette with 10-mL capacity or small plastic beaker for adding formalin to periphyton
samples
Plastic electrician's tape for sealing sample jars
Sample labels
Littoral and snag sampling form
214
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11.6 Literature cited
Hill, B.H., AT. Herlihy, and P.R. Kaufmann. 2002. Dehydrogenase activity in sediments from
Appalachian Mountain, Piedmont, and Coastal Plains streams of the eastern USA.
Freshwater Biology 47:185-194.
Hill, B.H., AT. Herlihy, P.R. Kaufmann, M.A. Vander Borgh, and S.J. DeCelles. 2003.
Assessment of streams of the eastern United States using a periphyton index of biotic
integrity. Ecological Indicators 2:325-338.
Pan, Y., R.J. Stevenson, B.H. Hill, A. Herlihy, and G.B. Collins. 1996. Using diatoms as
indicators of ecological conditions in lotic systems: A regional assessment. Journal of
the North American Benthological Society 15:481-495.
Peck, D. V., Averill, D. K., Herlihy, A. T., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Cappaert, M. R., Magee, T. and
Monaco, P. A. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Non-Wadeable
Rivers and Streams, U.S. Environmental Protection Agency, Washington, DC.
Peck, D. V., Herlihy, A. T., Hill, B. H., Hughes, R. M., Kaufmann, P. R., Klemm, D. J.,
Lazorchak, J. M., McCormick, F. H., Peterson, S. A., Ringold, P. L., Magee, T. and
Cappaert, M. R. Unpublished draft. Environmental Monitoring and Assessment Program
- Surface Waters Western Pilot Study: Field Operations Manual for Wadeable Streams,
U.S. Environmental Protection Agency, Office of Research and Development,
Washington, DC.
Sinsabaugh, R.L. and C.M. Foreman. 2001. Activity profiles of bacterioplankton in a eutrophic
river. Freshwater Biology 46:1239-1249.
215
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(SG)
GRW04449-
/
L
1 2
300255
(PA)
(4%
GRW0444S-
/
Composite volume ml_
visit number 1 2
300211
Figure 11-1. Set of sample labels for the site. Labels are affixed to the outside of the jar or
bag (sediment). Bottom number is the unique sample ID. Not actual size.
216
-------
217
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