&EPA
UnitBd Statn
Environmental Protection
Agency
Office of Water
Refutations and Standards (WH-553)
Washington DC 20460
1982
Final Draft Report
Water
SAMPLING PROTOCOLS
FOR COLLECTING SURFACE WATER,
BED SEDIMENT, BIVALVES, AND FISH
FOR PRIORITY POLLUTANT ANALYSIS
Final Draft Report
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FINAL DRAFT REPORT
SAMPLING PROTOCOLS FOR COLLECTING SURFACE WATER,
BED SEDIMENT, BIVALVES, AND FISH FOR
PRIORITY POLLUTANT ANALYSIS
EPA Contract 68-01-6195
Work Assignment No. 4
U.S. Environmental Protection Agency
Office of Water Regulations and Standards
Monitoring and Data Support Division
401 M Street, S.W.
Washington, D.C. 20460
Project Officer: Mr. Rod Frederick
Work Assignment Manager: Mr. Michael Slinak
Prepared by:
VERSAR INC.
6621 Electronic Drive
Springfield, Virginia 22151
IUS 5029
Prepared for:
Dalton-Dalton-Newport
3605 Warrensvllle Center Road
Cleveland, Ohio 44122
May 28, 1982
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DISCLAIMER
This report has been reviewed by the Office of Water Regulations and
Standards, U.S. Environmental Protection Agency, and approved for publica-
tion. Approval does not signify that the contents necessarily reflect the
views and policies of the U.S. Environmental Protection Agency, nor does
mention of trade names or commercial products constitute endorsement or
recommendation for use.
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FOREWORD
In a previous effort for the U.S. Environmental Protection Agency,
Monitoring and Data Support Division (MDSD), Versar Inc. described sam-
pling protocols for fish, sediment, and water in shallow water environ-
ments. Those protocols are not applicable to many deep water situations,
such as estuaries and the Great Lakes. Deep water systems are the primary
source of commercial fisheries, and many other factors such as water
supply, waste assimilation, navigation, and recreation are affected by the
quality of deep water systems. This manual incorporates the content of the
previous sampling protocol document and includes methods for sampling deep
water environments. It also includes methods for sampling estuarlne
bivalves, since in estuarine systems bivalves may provide more information
on toxics than fish. Analytical methods (including quality control/quality
assurance) are being developed by EFA's Office of Research and Development
(EMSL/Cincinnati) and therefore are not described in this manual.
Versar Inc. of Springfield, Virginia, has been subcontracted by
Dalton-Dalton-Newport (DDK) of Cleveland, Ohio, under EPA Contract
68-01-6195, to develop the necessary sampling protocols for MDSD. This
document is the final draft report for the Work Assignment.
Responsible Personnel
Program Manager: Bruno Maestri
Work Assignment Manager: J. Randall Freed
Work Assignment Staff: Phil Abell, Douglas Dixon
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ACKNOWLEDGEMENTS
The authors gratefully acknowledge the management and technical
guidance provided by Michael Slimaic of the U.S. EPA, Monitoring and Data
Support Division; Gayaneh Contos and Bruno Maestri of Versar Inc.; and Dr.
Robert G. Rolan of Dalton-Daiton-Newport (the prime contractor under which
this task was performed). We wish to acknowledge Ralph Huddleston for his
contributions to the fish sampling chapter, and the many reviewers and
scientists within EPA, the U.S. Geological Survey, U.S. Fish and Wildlife
Service, and State agencies who provided ideas and comments for this
manual. Also, the efforts of Teresa Halsey, Thompson Chambers, Juliet
Crumrlne, and the Versar secretarial staff are gratefully acknowledged.
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TABLE OF CONTENTS
Page No.
1.0 INTRODUCTION 1-L
1.1 Background 1~L
1.2 Recommendation for Use 1~2
2.0 SAMPLING AMBIENT WATER 2~l
2.1 Site Selection 2~2
2.2 Sample Equipment and Use 2-3
2.2.1 Cylindrical Samplers 2-8
2.2.2 Bottles (opening at one end) 2-12
2.2.3 US-Series Integrating Samplers 2-17
2.2.4 Bag Samplers 2-20
2.2.5 Pump Samplers 2-20
2.3 Sampling Procedures. ...» 2-22
2.3.1 Rivers, Streams, and Creeks 2-22
2.3.2 Lakes, Ponds, and Impoundments 2-24
2.3.3 Estuaries 2-25
2.3.4 General Procedures 2-26
2.4 Container Selection and Cleaning 2-27
2.4.1 Container Selection 2-27
2.4.1.1 Metals and Inorganics 2-28
2.4.1.2 Cyanide 2-28
2.4.1.3 Asbesto 2-28
2.4.1.4 Volatile Organics 2-28
2.4.1.5 Extractable Organics 2-29
2.4.1.6 Total Phenolics 2-29
2.4.2 Container Washing 2-29
2.4.2.1 Metals Containers 2-30
2.4.2.2 Cyanide Containers 2-31
2.4.2.3 Asbestos Containers 2-31
2.4.2.4 Volatile Organics Containers 2-31
2.4.2.5 Extractable Organics Containers 2-32
2.4.2.6 Total Phenolics Containers 2-32
2.5 Sample Handling, Preservation, and Shipment 2-32
2.5.1 Sample Handling 2-33
2.5.2 Sample Preservation 2-37
2.5.2.1 Metals 2-37
2.5.2.2 Cyanide 2-38
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TABLE OF CONTESTS (Cont.)
Page No.
2.5.2.3 Asbestos 2*38
2.5.2.4 Volatile Organic* 2-39
2.5.2.5 attractable Organics 2-40
2.5.2.6 Total Phenolics 2-40
2.5.3 Sample Transport 2-40
2.6 Quality Assurance/Quality Control Procedures 2-42
2.7 References .' 2-45
3.0 SAMPLING BED SEDIMENT 3-1
3.1 Site Selection 3-2
3.2 Sampling Equipment and Use 3-6
3.2.1 Corers 3-3
3.2.1.1 Teflon or Glass Tube 3-9
3.2.1.2 Gravity Corers 3-11
3.2.1.3 Free Fall or Boomerang Corers 3-16
3.2.1.4 Piston Corers 3-16
3.2.1.5 Multiple Tube Corers 3-18
3.2.2 Mechanical Grabs 3-20
3.2.2.1 Ekman Grab or Box Dredge 3-21
3.2.2.2 Petersen Grab 3-22
3.2.2.3 Ponar and VanVeen Grabs 3-24
3.2.2.4 Smlch-Mclntyre Grab 3-25
3.2.2.5 Jawed Grab Samplers 3-26
3.2.2.6 Pole-Operated Grabs 3-26
3.2.2.7 Shipek Grab Sampler 3-29
3.2.3 Scoops and Buckets 3-29
3.2.3.1 Rotating Bucket Sampler BMB-60 3-29
3.2.3.2 Scoops and Drag Buckets 3-31
3.3 Sample Handling, Preservation, and Shipment 3-31
3.4 References « « 3-39
4.0 SHELLFISH SAMPLING 4-1
4.1 Site Selection 4-2
4.2 Target Species 4-4
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TABLE OF CONTENTS (Cont.)
~~~"""~""~"~""~~"Page No.
4.3 Sampling Equipment and Use 4-6
4.3.1 Mechanical Grabs 4-7
4.3.1.1 Pole Operated Grab Buckets and Tongs... 4-7
4.3*1.2 Line or Cable Operated Grab Buckets.... 4-8
4.3.2 Biological Dredge 4-9
4.3.3 Coring Device 4-10
4.3.4 Miscellaneous Devices 4-10
4.3.4.1 Scoops or Shovels 4-10
4.3.4.2 Hakes 4-11
4.3.4.3 Dip Nets and Other Assorted Devices.... 4-11
4.3.5 Purchasing Specimens/Coordinated Sampling 4-11
4.3.6 Summary 4-12
4.4 Sample Handling, Preservation, and Shipment 4-13
4.5 References • 4-16
5.0 SAMPLING FISH 5-1
5.1 Site Selection 5-2
5.2 Target Species 5-6
5.2.1 Freshwater Target Species.... 5-8
5.2.2 Estuarine Species Selection 5-10
5.3 Sampling Equipment and Use • • 5-11
5.3.1 Active Collection 5-12
5.3.1.1 Electrofishing 5-12
5.3.1.2 Seines 5-14
5.3.1.3 Trawls 5-15
5.3.1.4 Angling 5-17
5.3.1.5 Poisoning 5-17
5.3.2 Passive Collection 5-13
5.3.2.1 Gill Nets 5-18
5.3.2.2 Trammel Nets 5-19
5.3.2.3 Hoop, Fyke, and Pound Nets 5-19
5.3.2.4 D-Traps 5-23
5.3.2.5 Purchasing Specimen.. 5-26
5.3.3 Summary 5-26
5.4 When to Collect 5-29
5.5 Container Selection and Cleaning 5-30
5.6 Sample Handling, Preservation, and Shipment 5-32
5.7 References 5-35
APPENDIX A
APPENDIX B
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LIST OF TABLES
.Page No.
Table 2-1 Summary of Water Sampling Equipment 2-5
Table 2-2 Container Type, Sample Volume, and Preservation 2-34
Table 3-1 Summary of Bottom Sampling Equipment. 3-32
Table 4-1 Target Species for Bivalve Shellfish 4-6
Table 5-1 Target Species for Warm Water, Cold Water and The
Great Lakes, and Other Cold Water Lentic Systems 5-9
Table 5-2 Target Species in Estuaries 5-11
Table 5-3 Summary of Fish Sampling Equipment 5-27
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LIST OF FIGURES
Page No.
Figure 2-1 Structural Features of the Kemmerer Water Sampler.... 2-9
Figure 2-2 Structural Features of the Van Oorn Water Sampler.... 2-10
Figure 2-3 Glass Lined Niskin Bottle 2-13
Figure 2-4 Sampling Device for Collecting Water to be Analyzed
for Priority Pollutants 2-15
Figure 2-5 Pole Operated Bottle Sampler 2-16
Figure 2-6 Depth-Integrating Hand Line Sampler, US DH-59 2-19
Figure 2-7 Bag Sampler 2-21
Figure 2-8 Sample Labels 2-36
Figure 2-9 Chain-of-Cuatody Tag 2-43
Figure 3-1 Coring Tube Adapted with Handle 3-12
Figure 3-2 Two Types of Coring Tubes with Handles. 3-13
Figure 3-3 Hand Corer 3-14
Figure 3-4 Phleger Corer 3-15
Figure 3-5 BMB-53 Piston Corer 3-17
Figure 3-6 Multiple Coring Tube 3-19
A & B
Figure 3-7 Ekman or Box Dredge 3-22
Figure 3-8 Petersen Grab 3-23
Figure 3-9 Ponar Grab Sampler 3-25
Figure 3-10 Jawed Grab 3-27
Figure 3-11 Controlled Depth Volumetric Bottom Sampler 3-28
Figure 3-12 Shipek Grab Sampler 3-30
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LIST OF FIGURES (Cont.)
Page No.
Figure 5-1 Beam Trawl 5-16
Figure 5-2 Otter Trawl 5-16
Figure 5-3 Gill Net 5-18
Figure 5-4 Trammel Net 5-20
Figure 5-5 Hoop Nets 5-21
Figure 5-6 Fyke Net . 5-22
Figure 5-7 Modifications of the D-Trap 5-25
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1.0 INTRODUCTION
1.1 Background
The U.S. Environmental Protection Agency, Office of Water Re-
gulations and Standards, is conducting a program to evaluate exposure and
associated risk from the presence of toxic pollutants in our nation's
aquatic environment. This program addresses Che goals of the Clean Water
Act of 1977 by developing exposure profiles for the 129 priority pollu-
tants. An exposure profile identifies subpopulations (geographic, de-
mographic, etc*) and environmental levels and forms of pollutants that come
in contact with these subpopulations. This sampling document describes
procedures for collecting, preserving, and shipping samples that will be
analyzed to provide information on ambient levels of the priority pollu-
tants, which is an essential component of the data base necessary for
characterizing exposure.
EPA's Monitoring and Data Support Division has responsibility
for providing technical guidance and coordinating the "Basic State Water
Monitoring Program" and the U.S. EPA "Regional Toxicant Monitoring Pro-
gram." A common goal of these programs is to collect and analyze ambient
data on toxic pollutants. The "Nationwide Urban Runoff Program" is focused
on determining the contribution of urban runoff to ambient levels of toxic
pollutants. These programs are now collecting, or are beginning to col-
lect, ambient samples of surface water, sediment, bivalves, and fish. Data
from these samples will be entered into EPA's water quality data base
(STORET) and toxics data base (TOXET) to be used in evaluating exposure and
risk.
The goal of this sampling document is to describe concise and
comprehensive field methods for collecting, preserving, and shipping sam-
ples of ambient water, bed sediment, bivalves, and fish that can be ana-
lyzed for toxic pollutants. Specific objectives of the document are to:
1. Evaluate the collection methods used in taking surface
water, sediment, bivalve, and fish samples.
2. Recommend appropriate sample devices, containers, and
preservatives.
l-l
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3. Recommend quantities required for analyses.
4. Recommend appropriate "target" species of fish and
bivalves.
5. Evaluate temporal considerations in sampling.
6* Prepare a concise field manual of sampling protocols.
The procedures outlined in this document emphasize methods that
are contamination-free and cost-effective. Since the objectives of the
programs using this document are to provide information on general trends
in water quality and presence/absence of toxic pollutants, many of the more
rigorous and more expensive quantitative techniques have been omitted.
Although highly sophisticated techniques are not considered vital for gen-
erating the "first-cut" data used in exposure profiles, they are useful in
verifying data or performing more intensive programs.
1.2 Recommendation for Use
These protocols were developed after a careful review of avail-
able literature, discussions with major EPA program and research offices,
and review by personnel involved in sampling and analytical studies within
EPA Regional Offices, the U.S. Geological Survey, and the U.S. Fish and
Wildlife Service. These protocols provide general guidance in the collec-
tion, preservation and transporation of ambient samples; no guidance is
provided for analytical protocols. Analytical protocols (including Quality
Assurance/Quality Control) are presently available from respective EPA pro-
gram offices and the EPA Office of Research and Development (Environmental
Monitoring and Support Laboratory, Clncinnatti, Ohio).
Sampling protocols for each of the different media (surface
water, bed sediment, bivalves, and fish) are considered in separate chap-
ters. These chapters are written to be independent of each other, and as a
result, there is some repetition of information.
Some general considerations apply to all field sampling pro-
grams, the most Important of which is the safety of the sampling crew.
Aquatic sampling should never be done by one individual since ropes, nets,
and other sampling equipment present an increased risk in the event of an
accident. The presense of at least two people provides the needed safety
1-2
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margin and usually results in a more efficient sampling effort. A one man
sampling crew is seldom cost-effective and is always unsafe.
Although the procedures presented herein emphasize methods to
avoid contamination, it is recognized that field conditions may sometimes
make it impossible or impractical to strictly adhere to the recommended
methods. In such cases, the data generated will not necessarily be inval-
id, but confidence in the data will be reduced. Because the purpose of the
programs that will use these methods is primarily to determine presence or
absence of priority pollutants, and follow-up monitoring of samples may be
performed where positive results are obtained, reasonable caution is the
key to successful sampling. All deviations from the recommended procedures
should be noted; effective quality controls can demonstrate the signifi-
cance of these deviations.
Although this manual does provide guidelines for sample handling
and site selection, it does not deal directly with designing a program.
The statistical and logistic considerations that affect program design are
in the province of the EPA and State offices that administer specific pro-
grams and are beyond the scope of this more general effort.
The procedures described in this manual are recommended for the
following specific EPA programs:
1. Ambient sampling under the "Nationwide Urban Runoff
Program."
2. Ambient sampling by EPA Regional Offices (and con-
tractor laboratories) for EPA'a "Exposure/Risk" pro-
gram, managed and coordinated by the Monitoring and
Data Support Division.
3. Ambient sampling by states under the "Basic State Water
Monitoring Program."
Recommendations, suggestions, and criticisms about these proto-
cols should be forwarded to:
Monitoring and Data Support Division (WH-553)
U.S. Environmental Protection Agency
401 M Street, S.W.
Washington, D.C. 20460
L-3
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2.0 SAMPLING AMBIENT WATER
Analysis of water samples provides information on Che nature and
quantity of constituents present in the water column. Whereas sedlnent,
bivalves, and fish concentrate some pollutants and can indicate general,
long term trends in water quality, water samples give a precise picture of
instantaneous conditions.
Laboratory methods used to analyze the dissolved and suspended
materials in water have advanced enormously in recent years and are con-
tinuously refined. However, a chain is only as strong as its weakest link,
and poor sampling technique has been the weak link in many water quality
studies because of problems with contamination or obtaining a representa-
tive sample. The objective of this section is to provide protocols for
collecting uncontaminated, representative water samples in the most cost-
effective manner.
In most natural waters, there is considerable heterogeneity, in both
time and space, in the concentrations of constituents in the water column.
Well-known examples of spatial heterogeneity are the thermal stratifica-
tion of lakes during the summer and winter and the chemical stratification
of estuaries. In rivers and streams, the greatest percentage of suspended
solids tends to be near the bottom. Because of the tendency of many pol-
lutants to be adsorbed by solids, a higher concentration of these pollut-
ants may exist in this area. Furthermore, there is also lateral stratifi-
cation in that discharges or tributaries may travel downstream as a plume
for quite a distance before becoming well-mixed with the receiving stream.
Temporal heterogeneity is related to variation in both natural
hydrologic conditions and point and non-point discharges of pollutants.
There are two basic ways to approach the problem of heterogeneity of
water in water quality studies:
I. Take numerous grab samples (a grab sample is a sample
taken at a particular location in the water column at a
single time) and analyze each.
2. Take composite samples (a composite sample is a mixture
of discrete samples (subsamples) which can be taken
from different places at different times).
2-1
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Since the costs associated with laboratory analysis are usually much
greater than the costs of sampling, composite samples are often more cost-
effective. For the programs using the protocols set forth here, composite
samples formed by mixing water from several sites in close proximity from a
single stream, river, estuary, or lake are the best approach (grab samples
are sufficient in very small, well-mixed streams)' The more intensive
labor and equipment requirements for time-composited samples are not neces-
sary under most circumstances.
Due to the fact that the priority pollutants are usually found in ex-
tremely low concentrations in ambient waters, the spectre of contamination
is more threatening with water than with sedimen-t, bivalves, or fish.
Precautions must be taken with sampling equipment, containers, and pre-
servatives to ensure that no contamination occurs* Furthermore, adequate
preservation methods must be employed to prevent changes in the nature of
the sample between the actual sampling and the analysis in the lab.
Quality Assurance and Quality Control (QA/QC) procedures should also
be Incorporated in the sampling protocol. These procedures improve the re-
liability of the data.
2.1 Site Selection
Selection of appropriate sampling sites is of utmost importance
in any sampling program. To the extent possible, water sampling sites
should coincide with the sites where bed sediment and fish samples are
taken. Other important factors include:
1. Locations where quality-based uses such as water supply,
fish and wildlife, and recreation may be threatened or where
good quality needs to be verified.
2. Easy access.
3. Location in a well-mixed zone where a minimal number of sub-
samples can be taken to yield a truly representative com-
posite sample.
4. Availability of historical or supplemental data (data on
priority pollutants are rare or absent in aost cases).
If there is interest in the effects of certain discharges on
ambient water quality, sites should be located both upstream and downstream
2-2
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of the discharges* When sampling below discharges or tributaries, sam-
plers should remember chat there is often a pronounced lateral variation in
water quality due to lack of mixing; plumes may extend for several*miles in
slow moving streams and rivers. Water quality also varies vertically.
Before any sampling is performed, an initial reconnaissance
should be done to locate suitable sampling sites. Check the distribution
and magnitude of currents during the reconnaissance so that sample sites
can be located in a representative manner. Bridges are often a good choice
as a site since they provide ready access and permit sampling at any point
across the width of the water body. Wading for samples is not recommended
in lakes, ponds, estuaries, and slow-moving rivers and streams because bot-
tom deposits are easily disturbed thereby resulting in Increased sediment
in the overlying water column. In slow-moving or deep water, a boat is
usually required for sampling.
If the study objectives are more rigorous than to describe the
general distribution and occurrence of pollutants, it is recommended that a
statistician be consulted for advice on site selection and number of sam-
ples needed. The statistical considerations for site selection are beyond
the scope of this manual, but are summarized in the EMSL "Handbook for
Sampling and Sample Preservation of Water and Wastewater" (EPA, 1981).
Other sources of information on site selection include works by Sanders
(1979), Kittrell (1969), and Mackenthun (1969).
2.2 Sample Equipment and Use .
A variety of devices have been developed for sampling water.
These range from a milk jug tied to the end of a stick to elaborate deep
ocean samplers costing thousands of dollars. Neither of these extreme ex-
amples is suitable for priority pollutant sampling. The milk jug sampler
is obviously capable of gross contamination, and the elaborate oceanograph-
ic samplers are usually far too expensive for routine sampling programs.
Samplers falling between these two extremes are discussed in the following
paragraphs.
Any device selected for priority pollutant sampling should have
the following features:
2-3
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1. Glass or Teflon* construction (or any other material which
does not contaminate the samples).
2. Operating ease at various depths.
3. Capability to be quickly and thoroughly cleaned.
4. Relatively inexpensive.
Regardless of the type of sampler chosen, it is a good quality
control measure to test any new sampling device (even if it is identical to
one previously used) by analyzing blank samples for all parameters for
which the device will be used. This can be done by filling the sampler
with distilled (not delonized) vater of known purity, allowing the water to
remain in the sample device for an amount of time equivalent to the time
required to take a sample, and then emptying it into a clean sample storage
bottle whose purity has been previously verified. Subsequent analysis of
this "blank" sample will reveal whether or not the device is capable of
contaminating samples.
In addition to Introducing contaminants to the sample, some mate-
rials (including certain types of glass) are capable of adsorbing metals to
the container walls. This property of a sampler can be tested by filling
the sampler with dilute (in the range frequently encountered in natural
waters) standard solutions of the metal ions to be sampled. After these
standard solutions have been left in the sampler for an amount of time
equivalent to the sample collection time, they are carefully analyzed to
determine if any metal ions have been adsorbed by the sampler. This test
should also be performed on the containers used for sample storage with the
necessary time modification and the addition of any preservative which will
be used. The same procedures are recommended to check sampling equipment
for organic pollutant contamination. Obviously, the sampling devices and
storage containers used to perform this quality control check should be
thoroughly cleaned to remove any traces of contamination before being used
for any actual sampling. Cleaning procedures are described in Section 2.4.
There are five basic designs of surface water sampling equip*
ment: cylindrical samplers (usually with openings at both ends), bottles
(opening at one end), the US-series integrating samplers, bag samplers, and
pumps. Each of these is discussed in the following pages. Table 2-1 is a
summary comparison of the principal types of equipment.
2-4
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Table 2-1
iry of Water Soap I Ing Equipment
Device
Kawmerer Hater
Sample Bottle
Kemmerer Mater
Sample Bottle
Van Dorn Bottle
Nansen Bottle
FJarlle Hater
Bottle
Nlskln Sampling
Bottle-Teflon
Coated Internal
Closure
Construction
Brass or nickel
plated brass.
Acrylic plastic.
ura thane end
seals.
Acrylic plastic.
PVC. other
materials.
Brass tubing,
Teflon-lined.
Heoprene gaskets.
Brass tubing.
Teflon-coated.
Rigid PVC with
optional Teflon
coating.
Closure Size Depth Range
Mechanism Range Maters (feet)
Internal rod or wire. 0.4 to 8L 1(3) to 90
Messenger activated. (300)
Internal rod or wire. 1.2 to BL 1(3) to 90
Messenger activated. (300)
Internal elastic 2 to SOL Surface to
tubing. Messenger (0.3 to 90 (300)
activated. 7.9 gal.)
Rotary valves close I.25L Surface to
when bottle turns (0.35 gal.) deep ocean.
end-over-end. and 1.51
Messenger activated. (0.4 gal.)
Spring-loaded metal I.3L Surface to
cover plates with (0.39 gal.) deep ocean.
rubber discs to
seal ends. Messenger
activated.
Internal latex 1.7 to 30L 1.6(6) to deep
tubing. Messenger (0.4 to 7.6 ocean.
activated gal.)
Possibility of
Contamination
Metals from bottle
and from closure
mechanism.
Ftithalate esters
and possibly
other organlcs.
Metals possible
from closure
mechanism.
Phthalate esters
and other
organlcs.
Minimal If Teflon-
lined.
Passible organlcs
or metals contami-
nation from rubber
seal.
Trace metals and
organlcs from
latex closure
mechanism.
Organlcs from
PVC If not Teflon
coated.
Advantages
Ease of usej
good depth
range.
Ease of use;
good depth
range.
Ease of use;
good depth
range.
Can be used
In series
for deep
water.
Flow-through
design.
Can sample
near bottom.
Multiple
bottles can
be triggered
remotely by
use of
appropriate
accessory.
Disadvantages
Separate sampler
needed to sample
for metals.
Separate sampler
needed to sample
for phthalate
esters. Possible
metals contamination
from closure
mechanism.
High probability
of contamination.
Reversing operation
prevents sampling
close to bottom.
Possible contamina-
tion.
Not convenient for
very she! low low
water. Some con-
tamination possi-
bilities.
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Tabla 2-1 (Cont.)
Summary of Mater Sampling Equipment
Device
Flow-through
NUMn Typa
Bottle Tetlon-
Coatod -Exter-
nal Closure.
Glass-lined
Contamlnatlon-
fr»« Modifi-
cation of
Ml skin Bottle
Grab Sampler-
G|«ts Battlo
On Una
Grab Samplar-
61 ass Bottle
on Pale
US 011-59 Depth-
Intagratlng
Sampler
Construction
Rigid PVC with
optional Taflon
coating.
Body-anodlzed
aluminum alloy*
Pyrex glass Inner
sample chamber.
Flow restrlctor.
cliack valve.
rupture disc
ot stain lass
st«al.
Glass bottle
support ad by
•atal fraaa-
Taflon cap.
Aluiclnu* polo
wltfc glass
SBMpla bottla-
Taflon cap.
Bronza (avall-
abla with apoxy-
palntad body.
nylon nozzles.
and slllcone~
rubber gaskets
for trace avtals
soapllng).
Closure
Hochanlso
Close-open-c lose
valvas. No Intarnal
•achanls*.
Stainless steel disc
ruptures at preset
depth. Stainless
steal check valve
prevents contamina-
tion after bottle
Is filled.
Spring closure.
Pulled open by
auxiliary line
at depth.
External lever on
pole.
N/A I see text for
operational da-
tails}.
Size Depth Range
Rang* Maters (feet)
1.7 to 60L t.B<6) to deep
(0.4 to 15.6 ocean.
gal.)
3 to 901. Moderate depth
(1.3 to to 4000 maters
23.3 gal.) (13.200 feet).
I3L (3.9
gal.)
Standard
size.
3.7L Surface to 9
(1 gal.) (30).
l2SaL to IL Surface to
(3.73 to approximately
30 oz.l l.a (6).
473mL 0 to 5.9 (19)
(14 oi.}
Possibility of
Contamination
Minimal If T*f Ion-
coated. Otherwise
good chance of
organ Ics con-
tamination from
PWC.
Minimal
Minimal
Minimal
Minimal organ Ics
with bronze body.
Minimal matals
Kith painted
body.
Advantages
Can penetrate
surface slicks
with minimal
chance of con-
tamination*
Lack ot con-
tamination.
especial ly
organ Ics.
Can penetrate
surface slicks*
Easily con-
structed.
Can penetrate
surface slicks.
Interchangeable
bottles.
Depth Integrat-
ing. Can be
operated from
brldgas or
boats. Hand-
line operation.
Disadvantages
Hot convenient for
shallow water.
Very expansive.
Vary heavy and
cumbersome.
Limited to rel-
atively shallow
waters.
Limited to shallow
waters.
Depth limit at ton.
Intake tube may
become plugged by
trash. Primarily
designed tor
((lowing water.
-------
Table 2-1 (Con*.)
Summary of Mater Sampling Equipment
Device
US 0)1-76 Duplh-
InlfrjroHng
Sampler
IS f-n Potnl-
Integratlng
Sampler
Baj Sampler
Construction
Bronze (available
Uth epoxy-pelnted
body, nylon
nozzles, and slll-
cone-rubber gas-
kets lor trace
metals sampling).
Aluminum (avail-
able utth epoxy-
palnled body.
nylon nozzles.
and slllcone-
rubbar gaskets
for trace matals
sampling).
Aluminum side
plates Kith
polyethylene
bttJS.
Closure
Mechanism
N/A (see tent for
operational de-
tails).
N/A (see tart for
operational de-
tails).
Messenger activated
knife cuts open
sealed fill tube.
Size Depth- Range
Range Meters (feet)
946ml 0 to J.fl (19)
<2B ox.)
473 or 946ml. 0 to 22 (73)
(14 or 28 with 473ml.
os.) conta Inert
0 to 19.9 (91)
tHth 946mL
container.
I.9L Unlimited.
40.4 gal.)
fosslblllty ol
Contamination
Minimal organlcs
tilth bronze body.
Minimal metals
irfth pa In tad
body.
Minimal organlcs
irfth aluminum
body. Minimal
metals Ml tit
painted body.
Fhthalate ester*
and other organlcs
from bag.
Advantages
Depth Integrat-
ing. Can be
operated from
bridges or
boats.
folnt-lnta-
gratlng. Can
•ample con-
tinuously
over a range
In depth.
Can penetrate
surface slicks j
may be useful
for metals.
Disadvantages
Depth limitation.
Intake tube may
become plugged by
trash. Primarily
designed for
sampling Hotting
wter.
Depth limitation.
Valvlng system
requires a DC
pou»r supply.
Patent lei lor
organlcs con-
tamination.
t •>
-I
-------
2.2.1 Cylindrical Samplers
One of the most frequently used designs employs a cylinder with
stoppers at each end and a closing device which can be operated remotely by
either a messenger weight or an electrical switch coupled to a solenoid de-
vice. Samplers of this design are the best type for sampling very deep
water. Depending on their specific valve and closure features, they may be
most susceptible to contamination by surface slicks, however. Included in
this family are the Kemnerer (Figure 2-1) and Van Dorn (Figure 2-2) bottle
samplers. The Kemmerer sampler uses a rigid rod or wire along which the
stoppers slide whereas the Van Dorn samplers typically utilize a flexible
piece of elastic tubing which pulls the stoppers into each end. Samplers
of this type are usually made of PVG (polyvinyl chloride), acrylic plastic,
or brass. Those made of PVC or acrylic plastic may contaminate samples
with phthalate esters whereas those made of brass may introduce contaminat-
ing metals. One possible solution to this problem is to use a metal sam-
pler for those samples to be analyzed for organics and a FVC sampler for
samples to be analyzed for metals. Some Van Dorn type samplers are now
available which are Teflon* coated to reduce contamination. Even this may
not ensure contaminant-free samples, however, because as Glam etc al. (1975)
report, Teflon* sheet material may contain up to 400 ppb of di-2-ethylbutyl
phthalate.
The Kemmerer Bottle is designed to be used primarily in depths
ranging from 1 to 90 meters (3-300 feet). Several versions are available,
including brass, nickel plated brass, and acrylic plastic with urethane end
seals. Sizes range up to about 8 liters. The acrylic model has a high
risk of contaminating organics samples while the brass model is probably
unsuitable for metals sampling. Again, before this sampler Is used, blanks
should be run to verify the absence of.any contamination. The Kemmerer is
also suitable for sampling from bridges and piers. If this (or any other
messenger operated sampler) is used from a bridge, care should be used in
dropping the messenger weight. If the weight is dropped from any con-
siderable height, 1C may severely damage the triggering mechanism. This
problem can be avoided by suspending the messenger a few feet above the
water by means of a string and then allowing it to free-fall from this re-
2-8
-------
9
Figure 2-1. Structural Features of the Kemmerer Water
Sampler (From APHA 1976).
2-9
-------
Figure 2-2. Structural Features of the Van Dora
Water Sampler (From APHA 1976).
2-10
-------
ducad height. Most Kemmerer bottles come from the manufacturer with a
small hole drilled in the side of the messenger to accomodate a string.
Modifications of the basic Kemmerer and Van Dorn samplers are
discussed below.
Nansen Bottle
This popular design has been used for many years in oceanographic
sampling. The bottle essentially consists of a brass tube with rotary
valves at each end. A messenger weight is dropped to release the catch
mechanism allowing the bottle to turn end-over-end, thus closing the
valves. Recent improvements include a baked Teflon* lining and simplified
wire clamp release assemblies. Since the bottle must turn end-over-end
(reverse) to operate the valves, sampling cannot be performed in proximity
to the bottom without danger of disturbing the sediment fines. The head
and valve assemblies of the newer bottles are removable to facilitate
cleaning. There is some chance of contamination of organics and metals
samples by the neoprene gaskets. This bottle is available in 1.25-liter
(1.3-quart) and 1.5-liter (1.6-quart) capacities.
Pjarlie Water Bottle
In this modification of the Van Dorn design, two spring-loaded
metal cover plates with rubber discs are used to seal each end of the brass
tube. This is also a flow through design, and the messenger activated
closure mechanism is external. The internal surfaces of the bottle are
Teflon* coated to prevent contamination. The capacity of this sampler is
1.3 liters (1.4 quarts). All operating parts are made of non-corrosive
brass or stainless steel.
Niskin Bottle
This sampler is available in sizes ranging from 1.7 to 30 liters
(0.4 to 7.8 gallons). It is constructed of PVC and is available with an op-
tional Teflon* coating. An internal closure device is used which consists
of latex tubing. This bottle is designed primarily for deep water sam-
pling, and multiple bottles may be used at different depths with triggering
at the operator's command. The Teflon* coated version may be well suited
2-11
-------
for priority pollutant sampling, but blanks (as described above) should be
run first to ensure that no contamination occurs. The latex closure device
nay possibly contaminate samples with zinc and iron (Segar and Berberian,
1975). There is also a distinct possibility of organics contamination from
the latex tubing.
Floy-Through Niskin Bottle
This sampler is an adaption of the Van Dorn design which uses no
internal closure springs or mechanisms. The sampler is constructed of
rigid PVC with an optional Teflon* coating available. A close-open-close
valve system opens automatically at a pre-set depth. This provides the
advantage of allowing the sampler to penetrate surface contamination (such
as oil slicks) with minimal risk of contaminating the internal sample area.
It is a good practice, however, to avoid lowering any sampler through
obvious surface pollution (unless the purpose is to sample directly beneath
such pollution), as oils and other substances may adhere to the external
surface of the sampler and be flushed inside when the sample is collected.
This sampler is available in [1.7- to 60-liter] (0.4- to 15.6-gallon)
capacities. The absence of any internal closure mechanism greatly reduces
the risk of sample contamination. Complete assurance that contamination is
not occurring, however, can only be achieved if sample blanks are run.
Glass-lined Niskin Bottle
This sampler is yet another modification of the Van Dorn design
and is manufactured specifically for oceanographic work. It consists of an
anodized aluminum alloy body with a Pyrex glass inner chamber (see Figure
2-3) and is available in sizes ranging from 5 to 90 liters (1.3 to 23.4
gallons). The 15-liter (3.9-gallon) size is standard. This is a special-
ized piece of equipment which nay be prohibitively expensive (over $1,000
for the smallest version) for general use. As a result of the inert
materials used in its construction, there is minimal chance of contamina-
tion from this sampler.
2.2.2 Bottles (opening at one end)
A popular type of sampling device, useful only in shallow water,
consists of a glass bottle attached to a pole or a line with some provision
2-12
-------
FILTER
FLOW aeSTRICTOR
RUPTURE DISC
CHECK VALVE
Figure 2-3. Glass Lined Niskin Bottle
2-13
-------
for opening the cap after submersion. Commercial devices of this design
are available or they may be readily constructed. Construction of a
typical sampler of this design (Figure 2-4) involves fastening a removable
glass bottle with 0.5-liter to 4-liter (0.1 to 1 gallon) capacity to a
stainless steel frame which holds a hinged, collar-type Teflon* lid (Gump
£t al. 1975). The frame has a Teflon* bumper to protect against impact.
The lid is held closed by a spring; when the bottle is dropped to the
desired depth, the spring is released by a tug on the attached string, air
rushes out of the bottle, and the bottle fills. Use of such a bottle
obviously requires ballast; a concrete block or other non-metal object is
preferred. The weight, tied to a rope attached to the bottle, should be as
far below the bottle as practical to prevent contamination. When sampling
is performed near the bottom, sufficient time should elapse for any sedi-
ments disturbed by the weight to be carried downstream before the sample is
taken. The weight should not be allowed to touch bottom in lakes or ponds
because the fine sediments kicked up by the weight will not settle out in
any reasonable period of time and will significantly bias the sample. The
same will often be true in estuaries.
The principal disadvantage of these homemade bottles is that the
spring holding the cap shut cannot be relied upon to keep the bottle closed
so that water is taken at the desired depth. Even if a very strong spring
is used, a great deal of ballast may be required to sink the bottle to the
preferred depth because of the buoyancy of the bottle. The ballast poses a
potential for contaminating the sample.
A type of bottle sampler which is commercially available consists
of a 1-llter (0.26 gallon) borosilicate glass sampling bottle with a
Teflon*-lined cap (Figure 2-5). The bottle is attached to a 1.9-cm (3/4-
inch) square aluminum tube 1.3 meters (6 feet) in length. The cleaned,
capped sample bottle is clamped onto the sampler and submerged. The oper-
ator can then remove the cap by means of a handle at the other end of the
tube, collect the sample from the desired depth, and recap the bottle.
This system has several advantages for shallow water sampling. The bottles
are relatively inexpensive, and a different bottle can be used for each
site which will minimize the chance of cross-contamination. This sampler
is also capped until the operator opens it which means it can penetrate
2-14
-------
PULL LINE
TEFLON GASKET
SPRING
WING NUT
SNAP SCHACKLS ATTACHED
TO SUPPORTING CHAIN
TEFLON BUMPER RING
GLASS BOTTLE
Figure 2-4.
Sampling Device for Collecting Water to
be Analyzed for Priority Pollutants
(After Gump at. al. 1975)
2-15
-------
I
Screw Cap
Sanple Bottle
Pack
Figure 2-5. Pole Operated Bottle Sampler
2-16
-------
surface slicks without contamination of the sample containing area* As
mentioned previously, a great deal of caution should be exercised if the
sampler penetrates surface contamination to ensure that contaminating mate-
rial from the outer surfaces is not flushed into the container when it is
opened. Obviously the depth to which these devices can be used is limited
by the length of the pole. Bottles are available for this sampler in capa-
cities ranging from 125 ml to 1000 mL.
2.2.3 US-Series Integrating Samplers
This is a group of samplers designed to acquire integrated (both
depth- and point-integrated) samples for suspended sediment analysis.
These samplers are useful for obtaining depth- or point-integrated water
samples for priority pollutant analysis as well.
Depth-integrated samples are taken continuously in a sampler that
moves vertically at an approximately constant rate between the surface and
the bottom. The sample enters the container at a velocity roughly equal to
the instantaneous stream velocity at each point in the vertical transit.
Point-integrated samples, on the other hand, are accumulated continuously
by a sampler held at a relatively fixed point in the stream. Again, water
enters the sampler at a velocity about equal to the Instantaneous stream
velocity at the point being sampled. The capability of the US-series
samplers to obtain samples isokinetically (filling rate proportional to
stream velocity) makes these samplers the beat choice when taking flow-
weighted samples in rivers and streams (using the EDI and EWI methods de-
scribed in Section 2.3). Most of these samplers use a glass bottle within
a metal (aluminum, bronze, or plated steel) body, and can be provided with
an optional Teflon* nozzle and medical-grade silicone gasket. The optional
epoxy-coated bodies available for trace metal sampling applications are not
recommended because the coating can chip off and contaminate the sample
with organic pollutants* Since the sample itself does not contact anything
but the Teflon* nozzle, the sample bottle, and a minute area of the sili-
cone gasket, the potential for contamination using the optional nozzle and
gasket and the standard body is minimal. Naturally, blanks should be run,
as described earlier in this section, to verify the absence of contamina-
tion.
2-17
-------
Although the U.S.-series integrating samplers are excellent for
use in streams and shallow rivers, they are not suitable for sampling deep-
er waters such as lakes and larger estuaries. Depth limits and other char-
•
acteristics are discussed below for three models operable with a hand
line.
US DH-59 Depth-Integrating Sampler
This sampler (Figure 2-6) is designed for hand-line operation.
The standard version is made of bronze, and optional Teflon* nozzles and
silicon*-rubber gaskets are available. Three different nozzles are avail-
able in sizes of 6.4 mm, 4.3 mm, and 3.2 mm. This enables the sampler to
operate at maximum depths of 2.7 meters (9 feet), 4.9 meters (16 feet), and
5.8 meters (19 feet), respectively.
US DH-76 Depth-Integrating Sampler
This is essentially a larger version of the DH-59 sampler with a
946-mL (1-quart) sample container. This sampler is also designed to be
uaed on a cable and the construction materials are the same as the DH-59,
including the availability of optional components for metals sampling. The
sampler is limited to a depth of 4.9 meters (16 feet) with all three noz-
zles.
US P-72 Point-Integrating Sampler
This is a point-integrating sampler designed for hand-line oper-
ation. The sampler is available with either a 473 mL (1 pint) or 946-mL (1
quart) sample container. The body of the standard version is constructed
of aluminum, and the optional nozzle and gasket are available. The only
nozzle size available is 4.8 mm, which limits this sampler to a depth of
2.7 meters (9 feet) when the 473-mL sample container is used or 4.9 meters
(16 feet) when the 946-mL sample container is used.
Other integrating samplers in the US-series are available for
fast water and other special applications. Additional details regarding
the US-series samplers and their use may be obtained by contacting:
Engineer-in-Charge
Federal Inter-Agency Sedimentation Project
St. Anthony Falls Hydraulic Laboratory
Hennepin Island and Third Avenue, SE
Minneapolis, Minn. 55414
2-18
-------
Figure 2-6. Depth-Integrating Hand Line Sampler, US DH-59.
2-19
-------
2.2.4 Bag Samplers
At least one manufacturer is currently offering a bag type sam-
pler for oceanographic work. This sampling device consists of a polyethy-
lene bag held by aluminum side plates (see Figure 2-7). The fill tube is
covered by a plastic sheath which is cut away upon opening. This feature
enables the sampler to penetrate oil slicks and other surface contamina-
tion. Because the actual sample container is constructed of polyethylene,
this device is not recommended for organic priority pollutant sampling. It
nay be useful for sampling metals and inorganics since the sampler is de-
signed Co orient itself against the current in such a manner as to place
the aluminum plates downstream of the sample opening. If the sampler is to
be used for metals sampling, the polyethylene bags should be tested by
filling with dilute standard concentrations of the metals to be analyzed.
These should be left in the bag for a time period equivalent to the resi-
dence time of the actual sample. The standard solutions should then be an-
alyzed to determine whether any adsorption has occurred. This is a good
procedure to use with any type of sampler which will be used for trace
metals.
If non-contaminating bags become available, or if it is proven
that the standard bags do not cause contamination, bag samplers may be used
for priority pollutant sampling.
2.2.5 Pump Sampler
A number of pump samplers have been developed, especially for
wastewater sampling. While the automated versions of these samplers are
unsurpassed for taking time- or flow-composited samples from point-source
discharges or small flow-controlling structures (e.g., weirs, flumes), they
are generally not practical for sampling most surface waters. Most have
neither the mobility nor the capability to take deep samples necessary for
sampling most natural surface waters. Also, they are quite expensive.
For streams and shallow rivers, pump samplers can be used to
obtain a composite sample by raising and lowering the intake line while
moving laterally across the water body (such a sample would not be
flow-proportional unless flow measurements and appropriate calculations
2-20
-------
Intake
Sample Bag
Bellows
Figure 2-7. Bag Sampler.
2-21
-------
were done beforehand). Several automatic sampler manufacturers have
introduced sample collection systems specifically designed for sampling
toxic pollutants, and "toxic sampler" replacement parts for older model
samplers. If an automatic sampler is used for sampling priority pollutants,
it should be equipped with these noncontaminating parts.
The variety of automatic samplers presently available is too
great to permit adequate discussion within the scope of this manual. Good
discussions of the types of such samplers available and their use are given
in EPA (1981), Shelley (1977), and Harris and Keffer (1974).
2.3 Sampling Procedures
The sampling regimes which may be used to collect priority pol-
lutant samples are nearly as variable as the types of available equip-
ment. Sampling regimes range from simple grab samples to multipoint,
flow-integrated composite samples. The level of sophistication required in
sampling technique is obviously related to the goals of the study, which
are, in this case, to determine whether measurable quantities of priority
pollutants occur in natural waters and to determine approximate concentra-
tions and distributions for those pollutants that are measurable. In view
of these goals, sophisticated time-compositing techniques are not generally
necessary, although they may be desirable at times (e.g., for verifying
unusual data).
Aquatic environments vary considerably in their physical charac-
teristics, and sampling procedures vary accordingly. Three basic types of
aquatic environments are covered by this manual: lotic (rivers, streams,
creeks), lentlc (lakes, ponds, Impoundments), and estuaries. Procedures
applicable to each are discussed below.
2.3.1 Rivers. Streams, and Creeks
In free-flowing bodies of water, the preferred sampling proce-
dure is to take a discharge-weighted sample. The advantage of discharge-
weighted sampling is that the samples will represent the average concen-
trations of pollutants transported by the water body. The spatial hetero-
geneity in pollutant distribution can lead to anomalous concentration and
loading values if discharge-weighted sampling is not performed.
2-22
-------
The most effective means of discharge-weighted sampling in shal-
low to moderate depth rivers and streams is to use the US-series integrat-
ing samplers in conjunction with depth integrating methods. As discussed
in Section 2.2, the US-series samplers take samples isokinetically, that
is, at a filling rate proportional to water velocity. Two different depth
integrating methods are commonly used to obtain a mean discharge-weighted
concentration for a cross section. These two methods are the equal-dis-
charge-increment (EDI) method and the equal-width-increment (EWI) method.
These two methods are explained in the National Handbook of Recommended
Methods for Water-Data Acquisition (USGS, 1980a), as follows.
"In the EDI method, the cross-sectional area is divided
laterally into a series of subsections, each of which conveys the
same water discharge. Depth integration is then carried out at
the vertical in each subsection where half of the subsection dis-
charge is on one side and half is on the other side. In each
individual subsection, a vertical transit rate is used that will
provide a sample volume for the vertical which Is equal to the
sample volumes for every other vertical. ... Generally, if more
than five verticals (more than five subsections) are sampled, an
accurate mean discharge-weighted concentration will be obtained."
"In the EWI method, depth integration is performed at a
series of verticals in the flow section that are equally spaced
across the transect to obtain a series of subsamples. Unlike the
EDI method, however, the vertical transit rate used at each ver-
tical is exactly the same as that used at every other vertical,
and the subsamples are composited even though they are of differ-
ent volumes. This procedure provides a transect sample whose
concentration is discharge weighted both vertically and laterally
and whose volume is proportional to the water discharge in the
sampled zone. An advantage of the EWI method is that a knowledge
of the lateral distribution of discharge is not required."
"The primary disadvantage of the EDI method is that the
lateral distribution of water discharge must be known or measured
each time prior to sampling. With the EWI method, on the other
hand, (1) it is sometimes difficult to maintain the same vertical
transit rate at all verticals, (2) more verticals must be sampled
for a given accuracy than with the EDI method, and (3) wherever
the flow is not essentially perpendicular to the transect, the
width increment between sampling verticals must be adjusted by
dividing it by the sine of the angle between the flow lines and
the transect. Generally, 10 to 20 verticals will provide an
accurate mean discharge-weighted concentration by the EWI
method."
2-23
-------
If sampling equipment other than the US-series samplers is used,
the EDI method can be adapted for use with other samplers. Flow measure-
ments would need to be taken first, and subsamples would be taken through-
out the cross-section corresponding to equal increments of flow.
If it is not practical to take a discharge-weighted sample be*
cause of equipment, time, or difficulty in making flow measurements, take
subsamples at several verticals along a transect. Locate the verticals in
a manner that is proportional to estimated flow, i.e., verticals should be
closer together at mid-channel where most of the flow travels, than toward
the banks, where the proportion of total flow is smaller. Take subsamples
at several depths for each vertical. The locations of all verticals, and
the depths at which subsamples are taken should be recorded in the field
notebook.
Aa previously mentioned, it is preferable to select sites that
are located in areas where the water is well-mixed. Certain river or
stream characteristics can point to well-mixed areas. Since the extent to
which mixing occurs is principally governed by turbulence and water velo-
city, the selection of a site Immediately below a riffle area will ensure
good vertical mixing. Horizontal (cross-channel) mixing occurs in con-
strictions in the channel. A method of verifying the presence or absence
of stratification or plumes is to traverse the river or stream taking
specific conductance measurements. Uniform specific conductance usually
indicates good mixing. Dissolved oxygen measurements can also indicate
stratification.
The number of verticals sampled along a transect is usually de-
termined in the field by the sampling crew. This determination is based on
mixing characteristics of the water body (the more homogenous the water
body, the fewer verticals needed).
2.3.2 Lakes, Ponds, and Impoundments
Lakes, ponds, and impoundments have a much greater tendency to
stratify than rivers and streams. The relative lack of mixing in lentic
systems requires that more subsamples be obtained.
2-24
-------
The number of sampling sites on a lake, pond, or impoundment
will vary with the size and shape of the basin. In small ponds, a single
vertical at the deepest point may be sufficient. In naturally-formed
ponds, the deepest point is usually near the center; in impoundments, the
deepest point is usually near the dam.
In lakes and larger impoundments, several verticals should be
composited to form a single sample* These verticals are often taken along
a transect or grid. Again, the number of verticals and the depths at which
samples are taken are usually at the discretion of the sampling crew. In
most cases, if the lake is stratified, it is recommended that separate
composites be made of epilimnetic and hypolimnetic zones. In unstratified
lakes, a composite consists of several verticals with subsamples collected
at various depths.
In lakes with irregular shape and with several bays and coves
that are protected from the wind, additional separate composite samples may
be needed to adequately represent water quality (USGS, 1980a). Similarly,
additional samples should be taken where discharges, tributaries, land use
characteristics, and other such factors are suspected of influencing water
quality. The number of separate composite samples that can be taken is ob-
viously a function of the need for spatial (or temporal) resolution in the
data and of the availability of manpower and funds.
The locations of all sampling sites, as well as field data on
parameters such as pH, dissolved oxygen, temperature, and other parameters
measured _in_ situ, should be entered into the field notebook during the
actual sampling.
2.3.3 Estuaries
The physical characteristics of estuaries are extremely complex
and variable. Thermal and chemical stratification, tidal action, and large
size make it very difficult to accurately measure flow patterns in estua-
ries. In most cases, it is recommended that subsamples be taken at various
depths along a transect or grid, with the verticals concentrated dear the
center of water mass. It may be advisable to take separate composites of
the layers above and below thermal and saline gradients because the water
2-25
-------
quality of these layers Is often considerably different. Always take tidal
stages into account when sampling estuaries and be sure to note exact date
and time of sampling.
AB with freshwater systems, a good method of determining layer-
ing Is to take measurements of temperature and specific conductance (or
salinity) at numerous depths and verticals along a transect. The number of
verticals and the depths at which subsamples are taken can be planned based
on the homogeneity of the water in the estuary.
2.3.4 General Procedures
Regardless of the actual sampling device used, mix the sub-
samples in a large compositing jug before pouring samples into the various
containers. A total of 8 liters is required for each composited priority
pollutant sample, including provision for spillage of 1 liter during trans-
fer.
Use a widemouth, glass compositing jug with a handle and
Teflon«-lined lid. Keep the lid on except when pouring samples in or out.
Volume of the jug should be about 15 liters to allow for collection of
duplicate samples for quality control. A commensurately larger jug will
be required if non-priority pollutant samples (e.g., solids, nutrients,
total organic carbon, oxygen demand) are also collected.
Different compositing jugs should be used for each composite
sample. The compositing jugs should be cleaned according to the same basic
procedure used for the sample containers (discussed in Section 2.4).
Before each subsample is taken, rinse the line connected to the
collecting device with site water to remove oil and other residues. Such
residues often accumulate In boat bottoms and similar places where line is
stored. Nylon line is preferred since it is durable and tends to absorb
less than cotton or polypropylene.
Grab samples should be taken for cyanide and volatile organ-
ics. If using a depth-integrating sampler, take the sample for cyanide and
volatile organics at the eentermost vertical; if using other sampling
equipment, take the sample at mid-depth at the eentermost vertical. If
separate composites are being taken for various layers (e.g., epilimnetic,
2-26
-------
hypolianstie) eaire the grab sample for cyanide and volatile organics at the
aid-point of the layer in the centernost vertical.
2.4 Container Selection and Cleaning
Water samples obtained for priority pollutant analysis require
careful handling to avoid the possibility of introducing Interferences,
both positive and negative, in the sampling process, in the containers used
for storage, or in transport to the lab for analysis. Possible routes of
positive interference or contamination include residues on the sampling
equipment (such as rust and corrosion products), leaching of materials from
containers, paint leached from the hulls of ships and boats, and dust and
other micro-particles in the sampling environment. Negative interferences
may arise as a result of the adsorption of chemicals to surfaces of con-
tainers or from the breakdown of samples due to improper preservation
procedures. The risks of contamination, adsorption, and desorption have
been reviewed by a number of investigators (Cooper, 1958; Robertson, 1968;
Tolg, 1972; National Bureau of Standards, 1974a, 1974b, and 1976). The
purpose of this section is to summarize the currently accepted practices
with regard to container selection and the cleaning procedures required to
reduce the risk of sample contamination. The container and cleaning
requirements specified below are summarized from Methods for Chemical
Analysis of Water and Wastes (EPA, 1979b) and the Office of Research and
Development proposed analytical procedures for priority pollutant analysis
(EPA, 1980).
2.4.1 Container Selection
The choice of containers and cap material is Influenced by a
variety of factors which include resistance to breakage, size, weight,
interference with sample constituents, cost, and availability. The most
important factor to consider, however, when choosing the type of con-
tainer is the possibility of interference with constituents for which the
sample is to be analyzed. This interference may result from adsorption of
a constituent by the walls of the container or from the release of an
interfering substance by the container. Suitable containers will be
described below according to the type of toxic pollutants for which the
sample is to be analyzed. These include metals and inorganics, cyanide,
asbestos, volatile organics, extractable organics, and total phenolics.
2-2:
-------
2.4.1.1 Metals and Inorganics
Over the last several years, considerable research has been con-
ducted on the sampling and analysis procedures for the determination of
trace metal concentrations in water samples. As a result of this research,
much information is available concerning the interferences that may be in-
troduced by the walla of the sample containers. Proper bottle selection
and preparation techniques can reduce such interferences.
Samples to be analyzed for toxic metals and Inorganics can be
stored in 1-liter polyethylene or high quality borosilicate glass bottles
with polypropylene caps. Teflon* lid liners should be purchased or cut
from sheet Teflon* and Inserted in the cap to prevent possible contamina-
tion from the caps normally supplied with the bottles. Metal bearing
materials, such as aluminum foil, must never be allowed to directly contact
the sample either in the sampling process or in sample storage.
2.4.1.2 Cyanide
Samples collected for cyanide analysis should be stored In con-
tainers exactly like those used for metals and inorganics.
2.4.1,3 Asbestos
Samples collected for asbestos analysis should be stored in 1-
liter plastic bottles.
2.4.1.4 Volatile Orsanies
Samples to be analyzed for volatile organics should be stored in
40-mL or 125-mL screw cap septum vials with a Teflon®-silicone disc in the
cap to prevent contamination of the sample by the cap. The discs should be
placed in the caps (Teflon* side down) in the laboratory prior to beginning
the sampling program. The discs should not be touched by hands at any time
but should be inserted with clean tweezers or some similar instrument. In
addition, extra discs stored in aluminum foil should be carried during
field sampling in case some of the discs previously placed in the caps are
lost.
2-28
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2.4.1.5 Extractabla Organ!es
At no time should plastic containers, such as polyethylene or
polypropylene, be used to store samples to be analyzed for the extfactable
organlcs (pesticides, base/neutrals, and acids). This is because samples
may be contaminated by leaching of substances (such as phthalate esters)
from the plastic container walls and because plastic containers are known
to adsorb many organic compounds such as pesticides (EPA, 1979b and .1980).
Samples to be analyzed for extractable organlcs should be stored in 3.8-
liter (1-gallon) amber bottles made of high quality borosillcate glass.
The amber glass reduces the possibility of photolytic reactions which could
alter the constituents of the sample. Nevertheless, the bottles should not
be left in direct sunlight for any extended period of time. All caps must
be lined with Teflon* to avoid sample contamination by the cap material.
Teflon* cap liners may be purchased or cut from Teflon* sheeting by the
laboratory personnel.
2.4.1.6 Total Phenolies
One-liter, high quality borosillcate glass bottles with a
Teflon«-lined screw cap should be used. Amber glass bottles are preferred
(but not required) since they inhibit photolytic reactions. If clear bot-
tles are used, care should be taken to keep the bottles in the dark to the
maximum extent possible.
2.A.2 Container Washing
Various procedures are used to clean the containers, depending
on the parameters being tested. In addition to the different sample bot-
tles, all sampling equipment containers and intermediate reservoirs should
*
be cleaned. The sampling equipment must be cleaned with regard to all
parameters on the priority pollutant list using the following procedure
(EPA, 1979b and 1980):
1. Wash all sampling equipment containers, caps and
Intermediate vessels with a non-phosphate clinical
laboratory grade detergent and hot water.
2. Triple rinse with tap water.
3. Rinse with 1:1 nitric acid (HN03 - Reagent grade).
2-29
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4. Rinse with deionized-distilled water.
S. Rinse with 1:1 hydrochloric acid (HC1 - Reagent grade).
6. Triple rinse with deionized-distilled water.
7. Rinse with acetone and finally with pesticide grade hexane.
8. Dry in a contaminant free area such as a laminar flow hood.
The aitric acid and hydrochloric acid washes are designed to remove trace
metals, and the acetone-hexane rinses are to remove organic impurities
which may interfere with the subsequent priority pollutant analyses. The
above cleaning procedure should be rigorous and should immediately precede
all sampling events. After they are dried, the equipment and containers
should be sealed and stored in a contaminant free area. The practice of
taking a "blank" sample before starting actual sampling (as described In
Section 2.5) helps to identify any contaminants that were not removed by
washing, or that were the result of the handling and transport process.
Methods for cleaning the various sample bottles are discussed
below. These methods apply to both new and used sample containers.
2.4.2". 1'" Metals Containers
Because of the sensitivity of the tests examining waterborne
trace metals, particular attention must be given to the thorough cleaning
of sample containers. The following schedule should be followed for the
preparation of all sample bottles and accessories, whether glass, poly-
ethylene, polypropylene, or Teflon*.
1. Wash with non-phosphate laboratory grade detergent and tap
water.
2. Rinse with 1:1 nitric acid (HN03 - Reagent grade).
3. Rinse with tap water.
4. Rinse with 1:1 hydrochloric acid (HC1 - Reagent grade),
5. Rinse with tap water.
6. Triple rinse with distilled-deionized water.
2-30
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2.4.2.2 Cyanide Containers
The following procedure should be used to wash bottles for
cyanide samples:
1. Wash containers and caps with a clinical, laboratory grade
non-phosphate detergent; scrub thoroughly with a brush (if
possible, wash liners and caps separately).
2. Rinse three times with tap water, then three tines with
distilled-deionzied water.
3. Invert to drain dry.
4. Visually inspect for any contamination prior to storage.
2.4.2.3 Asbestos Containers
Containers for asbestos samples should be washed as follows
(Anderson and Long, 1980):
1. Wash with clinical laboratory-grade non-phosphate detergent
and hot water.
2. Rinse with tap water.
3. Rinse three times with distilled water.
4. Rinse container twice with site water just prior to actual
sampling.
2.4.2.4 Volatile Organics Containers
The following procedure should be used to wash the septum vials,
Teflons-silicone septa, and caps used to contain volatile organics samples:
1. Wash vials, septa, and caps with a clinical laboratory grade
non-phosphate detergent and hot water.
2. Rinse three times with tap water, then three times with
distilled-deionized water.
3. Heat vials and septa at 105° for one hour in a clean oven or
muffle-furnace.
4. Allow the vials and septa to cool at room temperature In an
enclosed contaminant-free area, such as a laminar flow hood.
5. After cooling, seal the vials with the septa (Teflon* side
down) and screw on the caps. The vials should remain sealed
until the samples are taken.
2-31
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2.4.2*5 Extractable Organics Containars
Because of the prevalence of organic solvents and plasticizers
in the natural environment and the resulting risk of sample contamination,
the sample containers, Teflon* sheeting, and caps must be scrupulously
cleaned before use* The following procedure is recommended:
1. Wash the bottles, Teflon* liners, and caps in hot water with
a clinical laboratory grade non-phosphate detergent.
2. Rinse three times with tap water and three times with
distilled-deionzied water.
3. Rinse bottles, Teflon* liners and caps with acetone and
finally with pesticide grade hexane.
4. Air-dry at room temperature la an enclosed contaminant free
area such as a laminar flow hood. After they are dried,
the containers should be sealed until ready for use.
2.4.2.6 Total Phenolics Containers
Bottles for total phenolics samples should be prepared according
to the instructions given for cyanide samples.
2.5 Sample Handling, Preservation and Shipment
The handling, preservation, and shipment of water samples to be
analyzed for priority pollutants presents a significant challenge to the
Investigator because of the risks of sample contamination and the fact that
some samples start undergoing changes the instant they are sampled. The
risk of sample contamination, briefly discussed in Section 2.4, Container
Selection and Cleaning, cannot be underestimated. Many of the EPA-desig-
nated priority pollutant compounds are contained in common household items
such as disinfectants, paints, cleaners, and preservatives. Many others are
commonly present as a result of anthropogenic activities such as combus-
tion, vehicular waste, and pest control. In the laboratory, even more of
the listed compounds are used as common chemical solvents. Since these
compounds are encountered in the everyday life of the sampling and analysis
personnel, the likelihood of contamination of the sample is great unless
the proper sample handling and (previously discussed) cleaning procedures
are carefully followed. Besides the risk of sample contamination, the risk
of sample breakdown due to chemical or biological activity after the sample
2-32
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has been collected is also great. In general, the complete and unequivocal
preservation of samples is a practical impossibility, and complete stabi-
lity can never be obtained (EPA, 1979b). These obstacles, however, can be
overcome if the proper sample handling and preservation precautions are
followed. The purpose of this section is to discuss the recommended
methods for handling and preservation that are intended to retard biolo-
gical action, retard hydrolysis of chemical compounds, and reduce the
volatility of the components.
2.5.1 Sample Handling
. Once a water sample has been obtained, it must be carefully
mixed and poured into separate sample containers for metals and inorganics,
asbestos, extractable organics, total phenollcs, and cyanide (as previously
mentioned, grab samples should be taken for cyanide and volatile organics}.
The specific type and size of each container was discussed in Section 2.4
and is summarized in Table 2-2. Samples aay be nixed before splitting by
various methods including hand stirring with clean glass or Teflon* rods,
shaking or agitating sample containers, and magnetic mixing with Teflon*
coated stirring bars. None of the these methods is truly effective for at-
taining a homogeneous distribution of suspended solids throughout the con-
tainer. As a result, the last samples poured from the compositing jug usu-
ally have the greatest concentration of suspended solids. The recently de-
veloped USGS cone splitter may offer the best results for sample split-
ting; however, Judgment should await results of current EPA testing to
evaluate the splitter as a possible source of priority pollutant sample
contamination because of its plastic construction. The USGS splitter allows
the sampling team to obtain different subsample volumes from a sample while
still maintaining the same basic chemical and physical properties of the
original sample. Studies conducted by the USGS and EPA have shown that the
cone splitter can split samples as small as 250 mL into 10 equal subsamples
within + 3 percent of the correct volume and sediment concentration (USGS,
1980b). Unless it is proved that the USGS cone splitter does not cause
contamination, the recommended method of sample splitting is to stir or
shake the contents of the jug before pouring each sample, or to siphon
water while mixing the compositing jug.
2-33
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Table 2-2
Container Type, Sample Volume, and Preservation
(EPA, 1979 and 1980; Anderson and Long, 1979)
Saopla
?araMtar
Cyanide
Metals
Aa baa toe
Volatile Orjenica
Zxtr actable
Total Phenolic*
Sit* and Type
of Container
I liter polyethylene
or glaaa bottle
I UCK glaaa or poly-
ethylene bottle
1 liter plaatlc boccle
40 ml or 123 «1 glaaa
glue screw eop Tial
1 gallon amber flint
glaee battle
1 11 ear glaaa bottle
NuBbar
of
Sa«plea
1
1
1
2
1
1
Preaerration
Technique
tTaOB(l:10 dilucioa) eo Pa>l2;
raJrigarata eo 4"C
Nitric acid (1:1 dilution) eo
to pB<2
Koo* raqoirad* BaeoaaMnd
rafrlgaratlon or stora.|*
ia dark
Balriiaraca eo 4*C
tafrlgaraca eo *"C
H230* eo Pa<2
r*frig*r*ta eo 4*C
Holdiaf
TiM*
14
-------
The volatile organlcs samples require special treatment because
of the very volatile nature of the compounds to be measured* The volatile
organics vials should be completely filled to prevent volatilization, and
extreme caution should be exercised when filling the vials to avoid any
turbulence which could also result in sample loss. The sample should be
carefully poured down the side of the vial to minimize turbulence. A two-
fold or threefold displacement of water in the vial provides further assur-
ance of a representative sample. As a rule, it is best co gently pour the
last few drops into the vial so that surface tension holds the water in a
sort of "reverse miniscus." The Teflon^-silicone septa, Teflon* side down,
is placed over the convex miniscus and some overflow is lost, but airspace
in the bottle is eliminated. After capping, turn the bottle over and tap
it to check for bubbles; if any are present, remove the lid, top off the
bottle, and repeat the sealing process. It is also important to avoid the
risk of contaminating the volatile organics sample during handling and
transport. Samples can be contaminated by diffusion of volatile organlcs
(particularly fluorocarbons and methylene chloride) through the septum seal
into the sample. A field reagent blank, or "trip blank," prepared from
organic free water and carried through the sampling and handling protocol
can serve as a check on such contamination (EPA, 1980).
Samples should be clearly labelled with the date, time, sample
code, station location, sampling team member's initials, analysis request-
ed, and preservation treatment (See Figure 2-3). Duplicate samples should
also be prepared in order to provide a quality control check on laboratory
analysis. The number of duplicates prepared per sample batch should be
determined in consultation with the laboratory performing the analyses and
in accordance with the laboratory's quality assurance plan. The investi-
gator should note, however, that in order to meet duplicate needs, the
sample volumes collected will have to be adjusted. Duplicate samples
prepared for analysis should be labelled with an individual sample code
similar to other samples and not identified as a duplicate. The volume
level should be marked on all sample containers. This will indicate to che
laboratory personnel whether any sample has been lost in the shipment pro-
cess or whether the sample ha:i been contaminated by fluid entering the
2-35
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<•
•
^
<
at
u
•>
01
9
m
TOTAL METALS HN03 Added
OgiealSammNo.
i
i
Dviti* Tin*
SamfttrsSivmaui Offlct
Volume
Cammtim:
to
U
•.
a
CYANIDE NiOH A4dtd
Otffa-i^-«.
i
a
j—r—
JMlMrt^M* r«i»rv«tiv»:
•
•
«
^
<
«.
u
^
«
a
•
PHSNOLICS HjS04 AddM
OgMttSem&tNo.
i
a
OattaMTIm*
Hamtltfj Stpatvn Offlet
Volumt:
CammcnTi:
Figure 2-3. Sample Labels.
2-36
-------
container. The field blank collected should also accompany all samples and
duplicates as they are prepared for laboratory shipment.
Documentation during sample collection should include a cumula-
tive listing of sample times, methods, locations, and sample codes. Other
pertinent information, such as time the sample was taken, depth, and any in
situ measurements, should be recorded in a field notebook and cross-refer-
enced to the sample by using the identification number.
2.5.2 Sample Preservation
Refrigeration of samples to 4"C is a common technique used in
field work and helps stabilize samples by reducing biological and chemical
activity. All samples except metals must be refrigerated. In addition to
refrigeration as a general means of sample preservation, specific tech-
niques are required for certain parameters as discussed in the following
subsections. Table 2-2 lists preservation techniques aa veil as container
types, sample quantity, and maximum holding times.
2.5.2.1 Metals
Nitric acid (diluted 1:1 from concentrated reagent) should be
added to the sample to adjust the pH to less than 2. Again, the pH may be
tested by pouring several drops onto a piece of pH test paper. Adjusting
the sample to pH less than 2 will stabilize the sample for up to 6 months.
Neither nitric acid nor the preserved samples can be shipped by air freight
If the sampling trip involves air travel, the sample may be initially pre-
served by refrigerating and should be shipped to the laboratory immediate-
ly. Upon receipt in the laboratory, the sample must be acidified to pH <2
with HN03. At the time of analysis, the sample container should be
thoroughly rinsed with 1:1 HN03 and the washings added to the sample
(volume correction may be required).
When it is desired to determine concentrations of dissolved
metals in the water sample, the sample must be filtered through a 0.45
micron average pore diameter membrane filter immediately after collection.
It is advisable to discard the first ISO to 200 ml of filtrate in order to
rinse the filter and filtering apparatus of any contaminating substances.
After filtration, the sample (minimum volume 200 ml) should be preserved as
2-37
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discussed above. It is recommended that all-plastic filtering equipment
and pol7carbonace filters be used when taking dissolved metals samples
since glass filtering apparatus and cellulose membrane filters may cause
metal ion filtration losses (Trultt and Weber, 1979).
2.5.2.2 Cyanide
Oxidizing agents such as chlorine decompose many cyanides. If
the sample site is near a sewage treatment facility, power plant, or other
industry which may be using chlorine to treat discharge water, the sample
should first be tested to determine if residual chlorine is present. This
may be done by placing a drop of the sample on a piece of potassium iodide-
starch test paper. A blue color Indicates the need for treatment. If
treatment is required, add ascorbic acid, a few crystals at a time, until a
drop of the sample shows no color on the indicator paper. Then add an ad-
ditional 0.6 g of ascorbic acid for each liter of sample volume and mix
well. The ascorbic acid may be prepared in 0.6 g packets at the laboratory
prior to beginning the sampling. This test does not always give positive
results with small quantities of chlorine. This concentration, however,
will not significantly interfere with analysis procedures.
A HaOH solution (1:10 dilution previously prepared) should be ad-
ded to all cyanide samples to adjust the pH to a value greater than 12.
The pH may be adequately measured using pH test paper (the paper should not
be placed in the sample, but rather a few drops of sample can be poured
onto the paper). After the pH is adjusted, the sample should be refriger-
ated to 4*C immediately. By adjusting the pE and refrigerating the sample,
loss of hydrogen cyanide may be prevented. Samples should be analyzed for
cyanide within 14 days (EPA, 1980).
If air travel is involved in the sampling trip, no more than 0.95
liter (1 quart) of sodium hydroxide can be taken on a commercial flight.
This volume of NaOH should be more than adequate for normal sampling trips.
2.5.2.3 Asbestos
No preservation methods are required for asbestos samples. It is
recommended that the sample be refrigerated to 4°C or kept in the dark to
reduce biological activity.
2-38
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2*5.2.4 Volatile Organic*
All samples collected for volatile organics analysis oust be
Iced or refrigerated to 4°C from the time of collection until extraction.
If the sample contains residual chlorine, add sodium thiosulfate
preservative, (10 mg/40 mL will suffice for up to 5 ppm Cl2) to empty
sample bottles just prior to collection or shipment to the sample site.
Samples must be collected in duplicate with one sample preserved
with acid if more than 7 days will elapse before analysis. The acid
preservative, as described below, can degrade the alkyl compounds;
therefore, the acid preserved samples are to be used qnly for the analysis
of the aromatic hydrocarbons. The specific procedures to be followed when
7 days will elapse before analysis are aa follows (EPA, 1980):
1. In the first duplicate, do not use acid preservative. Fill
a sample bottle so that no air bubbles pass through the sam-
ple as the bottle is being filled. Seal the bottle so that
no air bubbles are entrapped in the bottle. Shake
vigorously for 1 minute if sodium thiosulfate is being used.
Label as "no acid sample." Maintain the hermetic seal on
the sample bottle until analysis.
2. Collect about 500 mL of sample in a clean container. Adjust
the pH of the sample to about pH 2 by adding HC1 (1:1) while
stirring vigorously. Fill a sample bottle so that no air
bubbles pass through the sample as the bottle is being
filled. Seal the bottle so that no air bubbles are
entrapped in the bottle. Shake vigorously for 1 minute if
sodium thiosulfate is being used. Label as "acid
preserved." Maintain the hermetic seal on the bottle until
analysis*
Before filling either of the duplicates with water, add a few clean boil-
ing beads to facilitate mixing.
Finally, all volatile organics samples oust be analyzed within
14 days of collection. In summary, if the sample is to be analyzed within
7 days of collection, no duplicate is required and the single sample may be
analyzed for both alkyl and aromatic compounds. If more than 7 days will
elapse before analysis, duplicate samples are required with the no-acid
sample analyzed for the alkyl compounds and the acid preserved sample
analysed for the aromatic compounds. ' ,
2-39
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2.5.2.5 Extractable Organic 3
All samples collected for extractable organics analysis must be
iced or refrigerated to 4°C from the time of collection until extraction.
Immediate delivery (within 24 hours) to the laboratory avoids the need for
the addition of chemical preservatives. However, if the samples will not
be extracted within 43 hours of collection, they must be preserved as fol-
lows: if the sample contains residual chlorine, add 35 tag of sodium thio-
sulfate per 1 ppm of free chlorine per liter of sample. All samples must
be extracted within 7 days and completely analyzed within 40 days of
collection (EPA, 1980).
2.5.2.6 Total Phenolics
Preserve samples for total phenols analysis by adjusting the pH
to less than 2 with reagent grade sulfuric acid (12804). Samples should
be stored at 4*C until analysis. The maximum holding time is 28 days (EPA,
1980).
2.53 Sample Transort
Samples should be directly transported to the laboratory the
same day they are collected. All sample containers should be placed in a
strong shipping container such as a metal picnic cooler. All lids should
be tightened before containers are placed in the shipping container. Glass
bottles should be separated in the shipping container by cushioning (e.g.,
styrofoam) or absorbent material (e.g., blotting paper) to prevent contact
with other objects and to eliminate breakage. For example, a 3.8-liter (1-
gallon) glass bottle (extractable organics samples) can be placed in tvo
carved out styrofoam sheets which secure the bottle at the top and bottom.
Small glass bottles (volatile organics samples) can be placed inside 0. 95-
liter (1-quart) plastic cubic containers with screw-type lids to minimize
breakage and contain any leakage.
Polyethylene bottles or plastic cubic containers do not require
cushioning material to prevent breakage but do need to be protected from
punctures by sharp objects.
All samples should be maintained at 4*C during transport. Ice
or synthetic "blue ice" can be placed in separate plastic bags and sealed,
2-40
-------
or in large mouthed cubic containers with lids. As an alternative, sample
bottles and ice can be placed together in a large sturdy plastic bag which
will provide an additional waterproof lining to the shipping container.
After all sample containers have been carefully arranged and ice has been
added, then samples should be delivered to the laboratory for analysis.
If the laboratory selected to analyze toxic pollutants is more
than 150 miles away, it may be necessary to use a commercial carrier that
provides overnight delivery to ensure that sample holding time constraints
are met. Regardless of the carrier, it is essential that arrangements for
sample pickup be made prior to sampling period termination. Most carriers
have a deadline for package pickup but are willing to make special arrange-
ments if notified in advance. It is advisable to establish an account
with the carrier and to obtain a supply of appropriate air bills to expe-
dite sample shipment. It'is imperative that the laboratory be given max-
imum time possible to coordinate sample delivery and analysis. Before
sampling is initiated, the laboratory should be contacted and informed of
Intended sampling activities.
Another shipping option is that of using a commercial airline.
However, this option requires making arrangements for laboratory personnel
to pick up samples when they arrive. Under normal circumstances, water
samples will not meet DOT criteria of hazardous materials as described in
49 CRF 170-179 and, therefore, may be shipped as non-hazardous. Samples
can be shipped in coolers by procedures similar to those described for lo-
cal transport. The shipping container must be marked "THIS END UP," and
arrows which Indicate the proper upward position of the container should be
affixed. A sticker containing the laboratory's name and address must be
placed on the outside container. Care must be taken to secure the drainage
hole at the bottom of the cooler so that if a sample container leaks or if
Ice water leaks through the ice bag, the contents cannot escape through the
drainage hole. The container should be taped shut in order to obtain as
much of a seal as possible around the lid to prevent any leakage.
As soon as sampling is initiated, the toxic pollutant analysis
laboratory should be notified so that they can be prepared for sample re-
ceipt. Immediately after shipment, the laboratory should be contacted and
2-41
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the following information relayed:
• Sample numbers
• Name of carrier
• Airbill numbers
• Number of containers
• Date and time of shipment
• Estimated date and time of delivery
• Any problems relating to shipment
When the samples are received at the laboratory, they should be
recorded in a permanent log book. This log book should include for each
sample, date and time received, source of sample, sample number, how trans-
ported to the laboratory, and the number assigned to the sample by the
laboratory if this number differs from the field number. Although this re-
cording procedure may seem laborious, it is absolutely imperative that pre-
cise records be kept for all samples so that the data generated by the sam-
pling and analysis effort is of unquestionable integrity.
An accurate written record should be maintained which can be
used to trace possession of the sample from the moment of its collection
until it has been analyzed. A chain of custody tag (Figure 2-9) should be
placed on all coolers in which samples are stored and shipped. This should
have appropriate spaces for signatures when the sample is transferred from
one person to another. The date and time at which the custody is trans-
ferred should be indicated on the tag.
2.6 Quality Assurance/Quality Control Procedures
Proper Quality Assurance and Quality Control (QA/QC) measures
are essential for ensuring that the sampling and analytical techniques em-
ployed produce data with known accuracy and precision. Before any sampling
occurs, a detailed QA/QC program should be coordinated between the sampling
team leader and the laboratory QA/QC officer. Although development of a
specific QA/QC program is beyond the scope of this effort, the following
controls should be incorporated:
2-42
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CHAIN OF CUSTODY RECORD
fTA'MM
NUM**
IT* now kOCATto*
•
e*ti
RttinquisJwd bv !«*•**'
Rttinoui*ft«d by: «*— *
Retinqui*ft«d by: «»•..»<
RKWVWJ by: '*»*•*«
Dispatched by: », .. .1
Mmtwd of Shipmwn:
Oltt
run
SAMPLERS -i— «
•MM>
c««*.
On»
*•
tia
MO.
•to. o«
COMTAINIftl
MUlVflt
•«oui*ia
Rtcwved by: r«^—i Oats/Tim*
Received by:»^»- Oa»/T«ie
flecaived by: <*~-~. Date/Tim*
analysis: ap»*.>
/Ttmt
1 Caoi-Surm Ciiardiniinr f*t» f<*t
Figure 2-9. Chain-of-Custody Tag.
2-43
-------
1. Sample blanks - Distilled water brought from the lab is
poured, in the field, into the collecting device, then
into the composite Jug, and then into the respective
sample containers. This is a check on contamination
from containers and lab water.
2. Reagent blanks - Distilled water is poured into sample
containers for cyanide, metals, and phenols, and ap-
propriate preservatives are added. This is a check
on the purity of reagents.
3. Spiked samples - A known quantity of pollutant is added
to one of a pair of duplicate samples. This is a check
on accuracy of analytical results and possible loss of
pollutants during storage.
4. Duplicate samples - Identical samples are taken, as-
signed separate sample numbers, and sent back to the
lab* This is a check on the precision of analytical
results.
The number of samples taken for each of the above controls should be suf-
ficient to identify contamination problems and to quantify accuracy and
precision.
2-44
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2.7 References
American Public Health Association (APHA). 1976. Standard
Methods for the Examination of Water and Wastewater. 14th
Ed. Washington, D.C. 1193 p.
Anderson, C.H., and J.M. Long. 1980. Interim Method for De-
termining Asbestos in Water. U.S. EPA. Office of Research
and Development. Athens, GA. EPA-600/4-80-005.
Bellar, J.A., W.L. Budde, and J.W. Eichelberger. 1979. The
Identification and Measurement of Volatile Organic Compounds.
In: Aqueous Environmental Samples in Monitoring of Toxic
Substances. D. Schuetzle, Ed. ACS Symposium Series No. 94.
American Chemical Society. Washington, D.C.
Cooper, L.H.N. 1958. A System for International Exchange of
Samples for Trace Element Analysis of Ocean Water. Journal
of Marine Research. 17:128-132.
Federal Working Group on Pest Management (FWGPM). 1974.
Guidelines on Sampling and Statistical Methodologies for
Ambient Pesticide Monitoring. National Technical Information
Service. U.S. Department of Commerce. Washington, D.C.
PB-239-798.
Feltr, H.R., and J.K. Culbertson. 1972. Sampling Procedures
and Problems in Determining Pesticide Residues in the
Hydrologic Environment. Pesticide Monitoring Journal.
6(3):171-178.
Feltz, H.R., W.T. Sayers, and H.P. Nicholson. 1971.
National Monitoring Program for the Assessment of Pesticide
Residues in Water. Pesticide Monitoring Journal. 5(1):54-59.
Giam, C.S., H.S. Chan, and G.S. Neff. 1975. Sensitive
Method for Determination of Phthalate Ester Plasticizers in
Open-Ocean Biota Samples. Analytical Chemistry.
47:2225-2229.
Harris, D.J., and W.J. Keffer. 1974. Wastewater Sampling
Methodologies and Flow Measurement Techniques. U.S. En-
vironmental Protection Agency, Region VII. Surveillance and
Analysis Division. Kansas City, MO.
King, D.L. 1971. Sampling in Natural Waters and Waste
Effluents. _In L.L. Ciaccio, (ed.), Water and Water Pollution
Handbook, Volume 2. Marcel Dekker Publishing, Inc., N.Y.
Kittrell, F.W. 1969. A Practical Guide to Water Quality
Studies of Streams. Federal Water Pollution Control
Administration. Cincinatti, Ohio.
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Mackenthun, K.M. 1969. The Practice of Water Pollution Bio-
logy. Federal Water Pollution Control Administration.
Washington, D.C.
Robertson, D.E. 1968. Role of Contamination in Trace Ele-
ment Analysis of Sea Water. Analytical Chemistry. 40(7):
1067-1072.
Sanders, T.G. (ed.). 1979. Design of Water Quality
Monitoring Networks. Colorado State University. Ft. Collins,
Colo.
Segar, D.A., and G.A. Berberlan. 1975. Trace Metal Con-
tamination by Oceanographlc Samplers. Amer. Chem. Soc.,
Advances in Chem. Series. 147:9-21.
Shelley, E. 1977. Sampling of Water and Wastewater. En-
vironmental Research Information Center Office of Research
and Development. U.S. EPA. Cincinnati, Ohio.
Tolg, G. 1972. Extreme Trace Analysis of the Clements - I:
Methods and Problems of Sample Treatment, Separation and En-
richment. Talanta. 19:1489-1521.
Truitt, R.E., and J.H. Weber. 1979. Trace Metal Ion
Filtration Losses at pH 5 and 7. Analytical Chemistry.
51(12):2057-2059.
U.S. Environmental Protection Agency. 1979a. Handbook for
Analytical Quality Control in Water and Wastewater
Laboratories. Environmental Monitoring and Support
Laboratory. Office of Research and Development. Cincinnati,
Ohio. Chapter 10.
U.S. Environmental Protection Agency. 1979b. Methods for
the Chemical Analysis of Water and Wastes. Environmental Mon-
itoring and Support Laboratory. Office of Research and De-
velopment. Cincinnati, Ohio.
U.S. Environmental Protection Agency. 1980. Draft Protocols
for the Analysis of Priority Pollutants. Methods 601-613, 624
and 625. Monitoring Technology Division. Office of Research
and Development. Washington, D.C.
U.S. Environmental Protection Agency. 1981. Handbook for
Sampling and Sample Preservation in Water and Wastewater.
Environmental Monitoring and Support Laboratory. Office of
Research and Development. Cincinnati, Ohio. (EMSL 0220).
U.S. Geological Survey. 1980s. National Handbook of
Recommended Methods for Water Data Acquisition. Office of
Water Data Coordination. Reston, VA.
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U.S. Geological Survey. I980b. USGS Cone Splitter. Quality
of Water Branch Technical Memorandum No. 80. Reston,
Virginia.
U.S. National Bureau of Standards. 1974a. Sampling, Sample
Handling, and Analysis. Symposium on Accuracy in Trace
Analysis. Proceedings of the 7th IMR Symposium. P.D. Lafleur,
Ed. NBS Special Publication 422. Washington, D.C.
U.S. National Bureau of Standards. 1974b. Marine Pollution
Monitoring (Petroleum). Proceedings of a Workshop. NBS
Special Publication 409. Washington, D.C.
U.S. National Bureau of Standards. 1976. A Survey of
Current Literature on Sampling, Sample Handling, and Long
Term Storage for Environmental Materials. U.S. Department of
Commerce. Washington, D.C.
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3.0 SAMPLING BED SEDIMENT
Sediment is an important indicator of pollution because it acts as an
information integrator and repository for many contaminants. Unlike water
»«»oples, which show only instantaneous conditions, bed sediment samples can
show long-term trends in the quality of the overlying water for some of the
priority pollutants. Most of the priority pollutants are partitioned more
strongly in sediment than in water; thus, if these pollutants have been
present recently and are not quickly degraded or desorbed, they are evident
in the sediment analysis. Bed sediment analysis can thus be an important
tool in studying the occurrence and distribution of priority pollutants.
Feltz (1980) summarizes the significance and use of bed sediment (bottom
material):
1. As a historical water quality integrator.
2. As a reconnaissance tool.
3. In planning analytical schedules.
4. In conducting short-lived studies.
5. For deriving short- and long-term trends.
6. For identification of problem areas.
As discussed later in this chapter, the capacity of sediments to
adsorb, coprecipitate, and otherwise bind pollutants varies widely depend-
ing upon such factors as grain size, organic carbon content, and iron and
manganese concentrations. For this reason, it is recommended that study
designs include provisions for analyzing particle size distribution, total
organic carbon, iron concentration, manganese concentration, and any other
parameters which could help In interpreting data from various sites.
This section has been developed as a guide for sampling bed sediment
in streams, rivers, lakes, and estuaries for bulk analysis for the EPA-
deaignited priority pollutants (Appendix A). It is intended to specify the
procedures that should be used in collecting, storing, and preserving
sediment samples, with the understanding that the results will be used to
indicate areas of potential water quality problems rather than as an exact
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historical record of long-cent conditions. As such, the equipment, pro-
cedures, and site selection techniques are geared to a semi-quantitative
approach emphasizing convenience, efficiency, and protection from'con-
tamination. Site selection is discussed in Section 3.1 of this report;
sampling equipment and techniques are described in Section 3*2; and sample
handling, preservation, and transport are discussed in Section 3.3.
3.1 Site Selection
The selection of sampling sites when screening for priority pol-
lutants requires considerable forethought to maximize the returns while
considering relative costs, data accuracy, and physical limitations. The
programs for which this manual has been written will use sediment samples
to indicate whether or not priority pollutants are present or have been
present in the overlying water column. Site selection, therefore, should
be limited to areas where (1) pollutants will have the best chance of being
detected if they are or have been present in the surrounding ambient
waters; (2) access is relatively easy in order to limit extensive field
operations and minimize costs; and (3) the sampling devices recommended in
this manual can be readily used.
The ability of sediments to sorb pollutants is related to
several factors, one of the more important of which is grain size. As
grain size decreases, the surface-to-volume ratio increases, so that there
is more contact with the water column and generally greater sorptive
capacity. The finer sediments such as clays and silts, therefore, will
have a greater probability of exhibiting pollutants than coarse sediments
such as sand and gravel. Thus, site selection should be limited to areas
where fine sediments are present in active deposits. A further advantage
of sampling fine-grained, unconsolldated sediments is that such sediments
tend to be relatively high in organic carbon. Organic carbon content is
another major factor affecting the amount of adsorption of both metals and
organic pollutants. It should be noted that consolidated sediments are
often representative of sediments deposited some time ago; therefore
unconsolidated sediments should be sampled.
Depositing areas are found where the current speed is slow.
Greatest deposition occurs where a stream slows down from fast current to
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slow current. Typical depositing areas include insidea of river bends,
downstream of islands, and downstream of obstructions in the water. Es-
tuarine areas, the region where fresh water mixes with salt water, are
often an area of enhanced deposition. This zone represents an area of con-
stant shoaling and a turbidity maximum usually occurs in the water column
(Holliday, 1978). Sites that are located immediately above or below the
confluence of two streams or rivers should generally be avoided. This is
because the flows from two tributaries do not necessarily immediately mix,
and the sediment may be moving almost as two streams in proportion to the
inflow from two tributaries. Potential sites upstream from the confluence
with another stream may also be unsuitable at times due to possible back-
flow which can disrupt the normal movement of sediment. When lakes, poods,
and reservoirs are to be sampled, consider sampling at the center of water
mass. This is particularly true for reservoirs that are formed by the
impoundment of rivers or streams. Generally, the coarser grained sediments
are deposited near the headwaters of the reservoir, and the bed sediments
near the center of the water mass will be composed of fine-grained mate-
rials (USGS, 1980). The shape, inflow pattern, bathymetry, and circula-
tion must all be considered when sampling sites are selected in lakes,
reservoirs, or estuaries.
Potential sampling sites must also be evaluated for accessibil-
ity. Selecting sites that can easily be reached by vehicle will greatly
reduce the time necessary for field activities. Bridges, piers, and other
projections over the water are readily accessible, but sediment conditions
may not be representative or appropriate for sampling (e.g., dredging is
often done around piers; bridges are often constructed at the narrowest
point in the water body and the constriction increases water velocity).
Frequently water body conditions will require the use of a boat; the site,
therefore, should be located in the vicinity of launching ramps or areas
where a boat is easily carried and launched.
Equipment availability will also help in determining suitable
sampling sites. Because the primary recommended collection tools are
Teflon* or glass tubes, shallow wadeable areas are best suited for sam-
pling. However, areas that are exposed during low flow or low tide periods
3-3
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should not be sampled, because they do not represent continuous exposure
and some pollutants can be more readily oxidized or volatilized when the
sediments are not Inundated. To ensure that periodically exposed sediments
are not collected, it is best to sample during low flow or low tide peri-
ods. It will not always be possible to locate sites in shallow, wadeable
areas. In some Instances (particularly in estuarine areas), it may be
necessary to sample deep waters or coarse substrates to determine sediment
quality in particular locations. In such cases, gravity corers or grabs may
be used to collect samples, and sits selection can be modified according-
ly.
As pointed out by Felt2 (1980), fresh sediment deposits
represent seasonal transport, and repetitive sampling and analysis of
samples from fresh deposits can reveal seasonal or short-term trend data.
In order to maximize information regarding such trends, the following
procedure may be used.
The geological study of stratigraphy is based upon the principal
of a vertical sequence of strata with older layers at the bottom of the
column and more recent layers at the top. Methods have been developed for
dating formations according to their characteristic fossils. While the
sediments of concern in the programs to which this manual pertains are all
of recent origin and do not contain characteristic fossils, a rough method
of identifying recent deposits within a specific area may be utilized. The
essential requirements for this procedure are a coring device capable of
taking three to four foot core samples and a means of identifying a precise
area of the bottom during repeated sampling efforts. The easiest means of
identifying such an area is to use a piling, stage height gage, bridge,
pier, or other permanent fixture.
During the first reconnaissance trip to the site, a core sample
is taken which includes the deeper and more permanent sediment layers. A
representative core from the site Is either saved for future reference or
carefully photographed. The stratigraphy of this core is documented and a
deep (old) layer is noted for future reference. All overlying layers are
recorded in terms of distance (in cm) above this layer. When sampling
personnel return to the site during the next scheduled sampling excursion,
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another core is taken which penetrates to at least the same depth. The
characteristic reference layer is identified, and again distances are
measured from this layer to each of the overlying strata. By this means,
newly deposited strata may be identified and any disturbance of the upper
strata resulting from storm events, etc., should be identifiable. This
method enables the Investigators to identify recent strata and determine
the extent of deposition or scour which has occurred since the last
sampling program. Such data are extremely valuable in interpreting
sediment transport phenomena.
Alternative methods of measuring deposition and scour are pos-
sible. A very simple method involves accurately measuring the distance
from the top of a piling, staff gauge, or other reference point to the top
layer of sediment each time. This procedure will only work, however, if
the water body has not been subjected to ice cover since the last sampling
program. Even moderate ice cover is capable of pulling piles upward as the
water level rises. Yet another possibility, more sophisticated and diffi-
cult, involves some means of taking a fix upon a permanent landmark on the
shore and determining the exact location by triangulation methods. Cores
are then taken in proximity to each other during the repeated sampling
trips.
If a piling is used as the reference point, samples should
always be taken upstream of the piling. This reduces the Influence of
scour which might occur as a result of propwash from motorboats which use
the piling for mooring purposes. The boats will naturally tend to drift
downstream of the piling because of the current, and scour is more likely
to occur in that direction.
If the effects of particular sources of pollutants on water and
sediment quality are to be investigated, sampling sites should be located
both upstream and downstream of the source. In order to facilitate com-
parison of samples, special care should be taken to ensure that sites are
located in areas with similar grain size characteristics. It should be
taken into consideration that most or all of the sediment at the downstream
location could have been upstream as recently as the last storm event and
may not have been exposed to pollutants long enough to accumulate them
significantly.
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Once a site has been selected, It will be the responsibility of
the Investigator to collect a sample which is representative of the de-
posited sediment in that area. In most circumstances, a number of samples
should be collected along a cross section of a river or stream in order to
adequately characterize the bed material. Investigators do not need to
establish a complex sampling transect when screening sediments for priority
pollutants. A common procedure is to sample at quarter points along the
cross section of the site selected. When the sampling technique or equip-
ment requires that the samples be extruded or transferred at the site, they
can be combined into a single composite sample. However, samples of dis-
similar composition which may alter or shed doubt on the representativeness
of the composited sample should not be combined but should be analyzed
separately in the laboratory.
In review, the criteria that should be used in selecting sample
sites include:
1. Depositing area, with slow current, where silt/mud/clay
is settling out. Nothing more coarse than sand (max-
imum grain size 2mm) should be collected, and silt
or clay is preferred.
2. Readily accessible by field personnel. Shallow wade-
able waters where the coring tubes can be driven by
hand are preferred. However, bridges, piers, or boats
may also be used, provided adequate sampling equipment
is available.
3.2 Sampling Equipment and Use
A number of collection techniques have been developed to sample
sediment material representative of different substrates. In the selection
of a sampling device, the investigator oust consider: (1) the nature of
the bed material to be sampled; (2) the structural detail of sediment
layering desired; (3) the amount of sediment material needed for analysis;
(4) the depth of water above the sediment; (5) the degree of sediment dis-
turbance by the sampling device; and (6) the possible interferences or
contamination introduced by the sampling device. In response to these
criteria, sediment sampling equipment can be divided into three basic
categories:
3f
-o
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1. Scoops or drag buckets.
2. Coring devices.
3. Mechanical grabs.
While sampling for priority pollutants, it is important not to
disturb the top layers of sediment and to minimize the loss of low-density
deposits during any sampling process. This is because the main emphasis is
on the sampling of recently deposited material, which may Indicate water
quality conditions in the fairly recent past (Feltz and Culbertson, 1972).
This material is generally unconsolidated and easily disturbed unless the
proper sampling equipment and precautions are used. All sampling equipment
will cause the formation of an hydraulic disturbance, or "shock wave," as a
result of its design and the manner in which it is used. The shock wave
may result in the displacement of the fine overlying sediment material be-
fore the sample is contained, or as it is retrieved to the surface. A
number of investigators (Wigley, 1967; Flannigan, 1970; Hudson, 1970;
Paterson and Fernando, 1971; Bowmiller, 1971) have reported on this
phenomenon. Most of their research has focused on the displacement of ben-
thic macroinvertebrates such as the small crustaceans, worms and larvae.
The principles, however, apply to the sediment fines as well. Of the three
major types of sampling devices, the scoops and drag buckets cause the
greatest degree of disturbance and, therefore, are not recommended for
priority pollutant sediment sampling. Corers and mechanical grabs also
disturb the sediment-water Interface to some extent; however, if pre-
cautions are taken, this disturbance can be minimized. The particular
limitations for each sampling device are discussed In their respective
sections•
The degree of contamination introduced by the sampling device or
technique must also be carefully considered. Plastics are particularly to
be avoided because they can introduce phthalate esters and compounds that
can interfere with pesticide analyses; they are also known to sorb pesti-
cides. Metal devices, which are prone to corrosion, may also introduce
interfering substances and if possible should also be avoided (Cooper,
1958; Robertson, 1968; EPA, 1977a). When conditions dictate the use of a
3--
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metal-containing device, strict attention should be given to the recom-
mendations for minimizing metal interference; otherwise, only devices con-
structed of high quality stainless steel should be used. High quality
•
glass and Teflon*, when properly cleaned, offer the most satisfactory
material for priority pollutant sampling. Since the use of these materials
is limited to the coring devices, they are recommended as the primary tools
for priority pollutant sediment sampling. The various coring and mechani-
cal devices are reviewed in the following sections, with a discussion of
their limitations and of the precautions to be observed in their use.
Table 3-1 at the end of this section summarizes the equipment discussed,
the suitable environment for equipment use, and the equipment's advantages
and disadvantages.
Finally, for reproducibility of sediment sampling results, re-
gardless of the equipment or technique used, operator training is absolute-
ly essential and sampling should not be entrusted to well-intentioned but
untrained staff or volunteers. This is, of course, true for all types of
environmental sampling.
In terms of volume, at least 500 mL (about 1 pint) should be col-
lected per site, and as a rule, it is advisable to collect 1000 mL (about 1
qt), in order to run duplicates or confirmation analyses when desired.
Check with the laboratory doing the analyses for the exact amount required.
Note that if additional analyses (e.g., particle size distribution, total
organic carbon) will be performed, additional volume will be required.
3.2.1 Corers
Core samplers are used to sample vertical columns of sediment.
They are particularly useful when a historical approach to sediment de-
position is desired, for they preserve the sequential layering of the de-
posit. Many types of coring devices have been developed depending on the
depth of water from which the sample is to be obtained, the nature of the
bottom material, and the length of core to be collected. Core samplers
vary from hand pushed tubes to explosive or weight driven devices. Since
priority pollutant sampling is generally concerned with the uppermost
sediment layers, only the devices that can adequately take shallow cores
are discussed below.
3-8
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Coring devices are particularly useful In pollutant monitoring
because (1) the "shock wave" created by descent is minimal, thus the fines
of the sediment-water interface are not disturbed; (2) the sample, is with-
drawn intact permitting the removal of only those layers of interest; (3)
core liners manufactured of glass or Teflon* can be purchased, thus re-
ducing possible sample contamination; and (4) the samples are easily de-
livered to the lab for analysis in the tube in which they are collected.
The disadvantage of coring devices is that a relatively small surface area
and sample size are obtained, necessitating repetitive sampling in order to
obtain the required amount needed for analysis. Because it is felt that
this disadvantage is offset by the Advantages, coring devices are recom-
mended in sampling sediments for priority pollutants. Following is a re-
view of the most commonly used coring devices.
3.2.1.1 Teflon or Glass Tube
In shallow wadeable waters, the direct use of a core liner or
tube manufactured of Teflon* or glass is recommended for the collection of
sediment samples. Their use can also be extended to deep waters when SCUBA
equipment is available. Teflon* is preferred to avoid glass breakage and
possible sample loss. The use of the tube by itself eliminates any pos-
sible metal contamination from core barrels, cutting heads, and retainers
and also cuts down on disturbance of surficial sediments (Flannigan, 1970)*
The tube should be approximately 13 cm (5 in) in length since only re-
cently deposited sediments (8-13 cm (3-5 in)) are to be sampled. Soft or
semi-consolidated sediments such as mud and clays have a greater adherence
to the inside of the tube and thus can be sampled with large diameter
tubes. Because coarse or unconsolidated sediment such as sand and gravel
will tend to fall out of the tube, a small diameter is required for them.
Since silt or clay materials are preferred for analysis of priority pol-
lutants, a tube about 5 cm (2 in) in diameter is usually the best size.
The wall thickness of the tube should be about 3 mm (1/3 in) for either
Teflon* or glass.
Actual sample collection is an easy operation conducted by one
person. Caution should be exercised, when the sample is obtained by wading
in shallow water, not to disturb the area to be sampled. The core tube is
3-9
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pushed into the substrate until only 2.5 cm (1 in) or less of the tube is
above the sediment-water interface. When sampling hard or coarse sub-
strates, a gentle rotation of the tube while it is pushed will facilitate
greater penetration and cut down on core compaction. The tube is then
capped with a Teflon* plug or a sheet of Teflon* held in place by a rubber
stopper or cork. After capping, the tube is slowly extracted, the negative
pressure and adherence of the sediment keeping the sample in the tube.
Before the bottom part of the core is pulled above the water surface, it
too is capped.
To help prevent contamination from direct contact between the
sampler's hands and the upper part of the tube, a collar-type device can be
constructed of wood and should have a circular recess to accept the top of
the tube. The recess should have a hole in it to allow water to pass
through when the tube is pushed in, and should be lined with sheet Teflon*.
Handles should be attached to the sides of the collar. After the tube is
driven in, impart a wide circular motion to help loosen the core for easy
removal; take off the collar device; cap the top of the tube (as described
above); pull it up out of the sediment layer; and cap the bottom of the
tube before removing it from the water.
Another method of obtaining recently deposited sediments in
shallow, wadeable waters with a core tube is to use the tube as a
horizontal scoop. The tube should be placed on Its side on the sediment
surface and carefully inserted into the sediment so that the top inside
surface is just at the sediment water interface. It is important to
disturb the fines as little as possible. After the tube is filled, both
ends should be capped with a Teflon* plug, as described above, before
the tube is removed from the sediment. If this method is used with a tube
having an outer diameter of 5.2 cm and wall thickness of 3 mm, only the top
4.5 cm of sediment will be sampled (allowing a 1 mm clearance between the
sediment surface and top inside of the tube).
Cores should always be kept upright, and should be immediately
transferred to a cooler.
As previously mentioned, a minimum of 500 mL, and preferably 1
liter, of sediment should be collected at each site. This translates into
3-10
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a minimum of 3, and preferably 5, tubes with 10 em-long (4 in) cores,
outer diameter of 5.2 cm (2 in), and wall thickness of 3 mm (1/8 in) (the
volume of eech core would be about 190 mL). For other tube sizes and core
lengths, the number of tubes necessary can, of course, be calculated by
using the formula for the volume of a cylinder.
When sampling water slightly deeper than wading depth or when
sampling shallow water from a boat, the coring tube may be modified by
adapting a handle to the tube. Many devices have been improvised Co
accomplish this goal; two custom-made devices (Maltland, 1969; Bouma, 1969)
are provided as examples (Figures 3-1 and 3-2). It will be necessary,
however, to Incorporate a valve or capping device at the top of the coring
Cube to prevent sample loss when the tube is retrieved.
When the sediment material is difficult to penetrate with a
Teflon* or glass tube, a commercially available hand coring device may be
used (Figure 3-3). These devices are equipped with a metal barrel with a
handle and a core liner. The liner is inserted and then held in place by a
screw-on core cutter, usually manufactured of stainless steel. The core
cutter, along with the handle attached to the core barrel, Increases Che
efficiency of sediment penetration. After the sample has been obtained,
the cutting head is removed and the liner carefully withdrawn and immedi-
ately capped as previously described. When coarse grain deposits such as
sand are sampled, the use of a core retainer will increase the efficiency
of sample retention. Only retainers manufactured of stainless steel should
be used in order to minimize the risk of trace metal contamination. When
several samples are to be obtained, it Is advisable to carry several core
liners to the sample site. This eliminates the need for time consuming
extrusions and permits the use of the core liners as sample containers for
shipment to the lab.
3.2.1.2Gravity Corers
Gravity corers, so named because the energy that causes them to
penetrate the bottom is derived from the momentum of free fall, are design-
ed for use in deeper bodies of water. Many variations of the gravity corer
have been devised; however, all have the same general characteristic of
taking a section of the bottom material that preserves the details of
layering.
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Figure 3-1. Coring Tube Adapted with Handle (from Maitland, 1969).
3-12
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Figure 3-2. Two Types of Coring Tubes with Handles (Van Stratten Tubes),
(A) for use in water up to wading depth
(3) for use from a small boat
(from Bouma, 1969).
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(Figure 3-3. Band Corer (Kahl Scientific Instrument Corp.)
Gravity corers essentially consist of a metal tube, preferably
stainless steel, with a detachable metal cutting nose on the lover end and
a weight and valve on the other. The metal nose facilitates penetration,
and the core catcher allows the sediment to slide up into the corer, but
prevents it from slipping back out. The valve permits the free passage of
water through the sampler as it descends, but aa it is retrieved, the water
pressure on the top of the corer keeps it securely closed. Many gravity
corers have been modified to accommodate a core liner. This eliminates the
need for time-consuming extrusions when repetitive sampling is necessary
and minimizes the introduction of interfering substances from the metallic
core barrel. Only those devices that can accommodate either a Teflon* or
glass liner are recommended when sampling for priority pollutants.
Selection of a gravity coring device depends upon the nature of
the substrate to be sampled. Coarse substrates such as gravel and sand are
generally unconsolidated and easily lost from the core barrel unless it has
a small diameter. Such substrates are most effectively cored by using
Phleger corers (Figure 3-4), which have a small diameter. The main dis-
advantage in the use of Phleger corers is that a relatively small sample is
taken, and repetitive sampling is necessary in order to obtain the required
amount for analysis. Finer sediments, such as mud, silt, and clay have a
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greater cohesive tendency, and thus larger core barrel diameters can be
used.
d
Figure 3-4. Fhleger Corer (from APHA, 1976).
The addition of a stainless steel core catcher to the bottom of
the corer greatly increases the efficiency of sample retention. Since core
catchers are made of metal and the chance of trace metal contamination ex-
ists, they should be used only when absolutely required.
The gravity corer is easily operated by a two-person crew from a
boat or any structure extending over the water surface. The equipment,
fastened to a flexible line of rope or wire, is lowered to within 2 or 3
meters of the bottom. Terminal velocity is generally achieved within this
distance (Bourne, 1969), and better accuracy and corer orientation is ob-
tained than with a free fall from the surface. The corer is retrieved to
the surface, cutting head unscrewed, and liner with sediment removed. Cau-
tion must be exercised at this point not to lose the sample, particularly
if it is coarse in nature. Only those cores that have some water in the
core tubes above the sediment should be retained. This ensures that the
sediment surface is intact and provides a reference point for determining
the sample depth below the sediment-water interface. After the core liner
has been removed from the barrel, the bottom and top of the liner should be
capped (as described in Section 3.2.1.1) and stored upright in an ice
filled cooler for delivery to the lab. The operation is repeated with a
new liner until sufficient sample for analysis is obtained.
3-15
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3.2.1.3 Free Fall or Boomerang Corers
Free fall or boomerang corers are gravity corers chat are oper-
ated without the use of flexible lines such as wire or rope. These devices
are equipped with an expendable weight and casing assembly. After the
bottom has been penetrated, a delay timer releases the weight and casing
and a buoyant device pulls the corer from the substrate and floats it to
the surface. This device is particularly convenient because cable connec-
tions to the surface are eliminated, and several corers can be dropped at
the same time. Simultaneous sampling saves time when sampling very deep
water, and higher impact velocity and accuracy are obtained (Hopkins,
1964). The device, however, should never be used in flowing water bodies
where corers can easily be swept away.
Free fall corers are primarily a deep-sea device, and their ex-
pense, large size, and unwieldiness make them generally unsuitable for
routine priority pollutant sampling.
3.2.1.4 Piston Corers
The piston corers have all the features of the gravity and hand
corers except that they incorporate a stationary piston within the core
barrel. Piston corers can be pushed by hand or operated by a flexible line
of rope or wire. After contact with the sediment, the piston remains sta-
tionary at the sediment surface while the coring tube penetrates the sub-
strate either by its own or by some externally applied weight. Since the
piston remains immobile, a partial vacuum is created over the core sample,
increasing core penetration and retention in the core barrel. The piston
also aids in easy extrusion of the core sample from the barrel or liner.
A common hand operated piston corer used for obtaining samples
from shallow water is the BMH-53 (Figure 3-5) developed by the Federal
Interagency Sedimentation Project (1940). The instrument is 120 cm (46 in)
long and is usually made of corrosion resistant materials. The collecting
end of the sampler is a stainless steel thin-walled cylinder 5 cm (2 in) in
diameter and 20 cm (8 in) long fitted with a rod mounted piston. The
BMH-53 is operated much like other hand pushed corers, being placed in a
vertical position on the stream bed with the piston extended to the open
3-16
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Figure 3-5. BMH-53 Piston Corer (USGS, 1930)
3-17
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end of the barrel* The barrel Is then pushed over the piston Into the
sediment. When coarser materials are being sampled, pulling on the piston
rod while pushing on the cylinder increases penetration and retention of
the sample in the liner. The piston also serves to force the sample from
the cylinder in a manner that results in a sample column with a minimum of
distortions. Since the BMH-53 does not have a liner, repeated extrusions
into cleaned sample jars are required in order to reuse the sampling de-
vice. After each extrusion, the sampler should be thoroughly rinsed with
site water to remove any remaining sediment material. If two fieldmen are
used to sample, this procedure can be quite rapid; however the operation la
quite awkward and time consuming for one man (Guy and Norman, 1969). The
BMH-53 can also be used in deeper water by attaching extension rods.
Because the BHM-53 is constructed primarily of metal parts, it is not
recommended for sampling inorganic priority pollutants.
3.2.1.3 Multiple Tube Corers
Since single coring tubes are limited in obtaining sufficient
material for chemical analysis, several types of multiple coring tube ap-
paratus have been developed. A triple corer (Kemp e£ aJL., 1971) (Figures
3-6A and B) has been developed that is particularly suited for sampling the
mud-water interface. Basically, three stainless steel core barrels are
welded inside a stainless steel ring. Each tube is provided with a valve
*
at the top end to ensure passage of water through the barrels during de-
scent and sealing upon retrieval. The nose end of each barrel contains a
screw mounted stainless steel cutting head. The core cutters are removed
for insertion of core liners. Only glass or Teflon* liners should be used;
plastic liners are unsuitable, because they may Introduce phthalate esters
and sorb organics.
Operation of the triple corer is the same as for wire supported
single tubes. Best results are obtained by lowering the device to within 2
or 3 meters of the bottom and then allowing it to free fall. Kemp et al.
(1971) state that good results have been obtained when sampling the fine
substrates such as mid, silt, and clays; however, the device was unsatis-
factory on coarse substrates such as sand and glacial tills.
One disadvantage introduced by connecting tubes in this manner is
that the increased surface area may cause a "shock wave" as the tubes de-
3-13
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Diagram of section of triple corer.
A-Core cube. B-Core barrel. C-Outer
ring. D-Retaining stops. E-Lifting
rod. F-Trigger mechanism. G-Valve
pin. H-Benthos valve in closed posi-
tion. I-Mud sample with overlying
water. J-Metal core cutter. K-Valve
in open position. L-Inner core barrel,
Sectional diagram of trigger
mechanism. A-Lifting rod.
B-Collar. C-Trigger pin. D-
Safety pin.
Figures 3-6A and B. Multiple Coring Tube (from
3-19
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acend which could disturb the sediment fines before they can be contained.
With caution and proper handling, this can be avoided, and the multiple
corer may eliminate much of the need for repetitive sampling. Never-
theless, in vadeable areas, not even the multiple corer is as expedient (or
contamination-free) as single core tubes for sampling sediment.
3.2.2 Mechanical Grabs
Various types of mechanical grabs (or dredges) have been de-
veloped depending upon the physical nature of the sediment material to be
sampled. All are similar in that they are devices with jaws which are
forced shut by weights, lever arms, springs, or cords. Caution should be
observed in the selection and use of a mechanical device. All, to varying
degrees, create a "shock wave" on descent which may disturb the fine mate-
rial of the sediment-vater interface where many of the priority pollutant
compounds are concentrated. Some degree of sample disturbance is also
possible as the device is retrieved. Mechanical grabs are constructed of
metal and may introduce trace contaminants. The major advantages in the
use of mechanical grabs is that they can sample a relatively large area and
secure a large amount of material, thus reducing the need for repetitive
sampling.
The use of a mechanical grab will require that the sample be
transferred from the sampler to a glass or Teflon* container for delivery
to the lab. Subsampling from the center of the sediment sample obtained,
particularly one with the integral structure of layering preserved by the
sampling device, with Teflon* or glass tubes will minimize the possibility
of metals introduction from the frame of the sampling device and will
provide a container for delivery of the sample to the laboratory (Greig «_t
aJU, 1977; Greig and McGrath, 1977; Prank e£ al._, 1977). Alternatively,
the use of a Teflon* or glass scoop is recommended for removal of the
sample. Contact of the sample with hands or other surfaces that may
introduce Interfering substances should be avoided. Following is a review
of the note commonly used mechanical grab sampling devices. Many of these
may be used, with proper precautions, for priority pollutant sampling. A
comprehensive review of most types of sediment sampling equipment is
provided by Hopkins (1964).
3-20
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3.2.2.1 Ekman Grab or Box Dredge
This device (Figure 3-7) Is widely used to sample mud, silt, and
other soft substrates (Welch, 1948; APHA, 1976). In studies conducted on
the Great Lakes, the Ekman dredge was found to be the most satisfactory de-
vice for sampling soft, unconsolidatad substrates (Howmiller, 1971). It is
not particularly efficient on coarse grain substrates such as sand, gravel,
and rock because of United penetration of the substrate and because small
pebbles or grit may prevent the proper closing of the jaws.
The device is made of 12 to 20 gauge brass or stainless steel.
The box-like sample container has spring operated jaws on the bottom that
are manually cocked. The jaws are triggered with a catch that is released
by a messenger (weight) after the sampler is resting on the bottom. The
top of the sample container is covered by two hinged overlapping lids that
are held partially open during descent by water passing through the sample
compartment. The lids are held shut by water pressure during retrieval.
The Ekman dredge is available in various sizes depending on the needs of
the investigation.
The device can be operated by one person from a boat or any ob-
ject extending over the water surface. It is slowly lowered by wire or
rope to within a few meters of the bottom before it is released for free
fall. This gradual descent will ensure proper orientation of the grab be-
fore impact and will reduce the possibility of a "shock wave." After the
sampler is resting on the bottom, a messenger (weight) is lowered to trig-
ger the jaws, and the grab is slowly retrieved. After retrieval, the sam-
ple may be easily subsampled through the lids of the box with coring tubes,
thus minimizing the possibility of sample contamination from the frame of
the device. The major disadvantage of sampling soft substrates with the
Ekman Dredge is that a significant portion of the finer sediments often
gets "washed out" of the dredge when water flows through it.
3.2.2.2 Peterson Grab
This device (Figure 3-8) is especially designed for sampling hard
or coarse substrates such as sand, gravel, and clay in swift currents and
deep water. It is a clam type grab manufactured of corrosion resistant
steel in various sizes that will sample an area of 0.06 to 0.09 m2. Weight
3-21
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Figure 3-7. Ekman or Box Dredge (from APHA, 1976),
3-22
-------
Figure 3*3. Pecarsen Grab
(from APHA, 1976).
3-23
-------
ranges from 14 kg (30 Ibs) to 32 kg (70 Ibs) depending on its size and on
whether auxiliary weights are bolted to its sides. The primary reasons for
using extra weights are to achieve stabilization in swift currents and
additional cutting power in the jaws.
The device can be operated by one person from a boat or object
extending over the water surface (heavy models nay require the use of a
winch). The jaws are set and the device is lowered slowly to the bottom in
order to minimize the "shock wave" which may disturb the lighter bottom
materials* The rope or cable is eased to release the locking catch. When
the grab is raised, the lever system closes the jaws and secures the
sample.
The construction of the Petersen Grab does not permit direct
access to the secured sample without opening of the closed jaws. This
process may further cause disturbance of the sediment material by destroy-
ing the integral nature or sequential layering of the sediment, particular-
ly if the sediment is unconsolidated as with muck and mud. Subsampllng
with coring tubes or Teflon* scoops may, therefore, not provide a repre-
sentative sample of the bottom sediment. The lack of a screen or door on
the top of the sample compartment which would allow water to pass through
the compartment as it descends, also results in the formation of a
relatively large shock wave. In comparative analysis with the Ponar Grab
(subsequently discussed), the Petersen Grab was observed to cause a much
larger displacement of the sediment fines (Wigley, 1967). Furthermore, the
metallic construction of the Petersen Grab may result in the introduction
of trace contaminants to the secured sample. It is, therefore, not
particularly recommended for priority pollutant sampling. If current or
substrate conditions dictate Its use, particular attention must be given to
its limitations.
3.2.2.3 Ponar and VanVeen Grabs
These devices (Figure 3-9) are modifications of the Petersen
Grab and are similar in size and weight. They have been modified by the
addition of side plates and a screen on the top of the sample compartment.
The screen over the sample.compartment permits water to pass through the
sampler as it descends thus reducing the "shock wave" and permitting direct
3-24
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access Co Che secured sample withouc opening the closed jaws. As previous-
ly reported, this advantage has been demonstrated in comparative analysis
with the Fetersen Grab (Wigley, 1967).
Figure 3-9. Fonar Grab Sampler (from APHA, 1976)
The Fonar and VanVeen Grabs are easily operated by one person in
the same fashion as the Fetersen Grab. The Ponar Grab is regarded as one
of che most effective samplers for general use on all types of substrates
(Flannigan, 1970; Hudson, 1970; Hovmiller, 1971; APHA, 1976). Access to
the secured sample through the covering screens permits subsampling of Che
secured material with coring tubes or Teflon* scoops, thus minimizing the
chance of metal contamination from the frame of the device.
3.2.2.4 Smith-Mclntyre Grab
This device is also similar to the Fetersen Grab. Its weight is
increased by the use of heavy gauge steel, and the jaws are closed by
spring loaded coils rather than the use of levers. While its efficiency on
hard or coarse substrates is better than the Fetersen or Ekman Grabs, its
bulk and weight require that it be operated from a boat equipped with a
winch. The bulk and weight of the grab also creates a relatively large
"shock-wave" on descent, thus disturbing the lighter bottom sediments be-
fore they may be contained. 'Generally, the Smlth-Mclntyre Grab is used for
sampling the benthos of deep lakes, estuaries, and areas of the continental
shelf because its great weight provides stability wnile in descent. It is
not recommended, however, for routine sediment sampling in shallow waters.
3-25
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If this grab is used, operators should use extreme caution as it is a
cumbersome and dangerous instrument.
3.2*2.5 Jawed Grab Samplers
These devices, such as the Orange-Peel Grab (Figure 3-10), are
designed for use on hard substrates in estuaries and deep lakes. The jaws
are operated by a wheel and sprocket mechanism within the upper framework,
which may be operated by a second cable or by a stack release mechanism
activated by a messenger. These devices frequently are impeded by incom-
plete closure of the Jaws caused by grit, gravel and other objects re-
sulting in sample loss. In a comparative study of the Orange-Peel Grab
versus the Ekman and Ponar samplers, the Orange-Peel Grab was found to be
the least efficient in terms of sample retention (Hudson, 1970). For this
reason, as well as the other limitations discussed for grab samples, the
jawed grabs are not recommended for priority pollutant sampling.
3.2.2.6 Pola-Operated Grabs
Many pole operated devices have been developed incorporating the
mechanics of the previously discussed grab samplers* These devices are
particularly useful for sampling shallow water lakes and streams. They
also offer the investigator a much greater degree of control over the sam-
pling operation than can be obtained by cable or hand line. The Ekman or
box-type pole operated grab sampler is recommended over the clam type grabs
because direct access to the secured sample is provided permitting sub-
sampling of the secured material with Teflon* or glass coring tubes. As
mentioned previously, this not only minimizes the risk of trace metal con-
tamination from the frame of the sampling device but also provides a suit-
able container for delivery of the sample to the laboratory.
An example of a custom made device which overcomes many of the
limitations of the other grab sampling devices is a controlled depth, volu-
metric bottom sampler (Figure 3-11) developed to sample to a constant depth
in most types of sediment over the entire area encompassed by the jaws
(Jackson, 1970). The sampler has closed ends and screened relief openings
to prevent blowout during placement or washout when the sampler is closed
and retrieved. Depth guides attached to the sides of the sampler determine
the depth to which the sampler can be thrust, thus preventing the jaws from
3-26
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Figure 3-10. Jawed Grab (from APHA, 1976).
3-27
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(J)
(I)
Figure 3-11.
Controlled Depth Volumetric Bottom Sampler:
Sampler in Open Position (A) Pivot bolt,
(b) Optional pivot bolts, (c) Removable
handle, (d) Pivot plate, (e) Lever arm,
(f) End plate, (g) Depth guide, (h) Guide
plate, (i) Carrier bolt and (j) Axis rod
(Jackson, 1970).
3-28
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penetrating too deeply in soft bottoms and exceeding the capacity of the
grab* This feature is particularly useful in sampling for priority
pollutants since it is the uppermost sediment deposits that may interact
with the overlying water. If used for priority pollutant sampling, this
grab should preferably be constructed of high quality stainless steel in
order to minimize the introduction of trace metal substances to the sample.
3.2.2.7 Shipek Grab Sampler
This grab sampler (Figure 3-12) consists of two concentric half
cylinders* When the grab touches bottom, inertia from a self-contained
weight releases a catch, and helical springs rotate the inner half cylinder
180 degrees. A sample is taken which is 0.4 m2 (8 in by 8 in) in surface
area and approximately 10 cm (4 in) deep at the center. Its bulk and
weight require that it be operated from a boat equipped with a winch. This
sampler is generally used for sampling marine waters and large inland bod-
ies of water. The Shipek retains fine sediments effectively, but there is
some chance of sample contamination due to its metal construction. This
grab is usually effective for deeper waters but may be unwieldy for routine
sediment sampling in shallow waters. Like all other samplers, quality con-
trol samples should be run initially to ensure the absence of contamination
before this grab is used.
3*2*3 Scoops and Buckets
3.2.3.1 Rotating Bucket Sampler - BMH-60
The USBHM-60 developed by the Federal Interagency Project of
1963 is a hand line, spring-driven, rotary bucket bed material sampler.
This device is designed primarily for use in coarse bed streams and rivers
(Guy and Norman, 1969). It is not particularly efficient In mud or other
soft substrates because its weight will cause penetration to the deeper
lying sediments, which is not desired when sampling for priority pollu-
tants* It is also difficult to release secured samples in an undisturbed
manner so that they could be subsampled. Nevertheless, the BMH-60 is
streamlined and is one of the few samplers capable of sampling in moving
water from a fixed platform. The BMH-60 may be used for priority pollutant
sampling if care is taken to collect only subsamples that have not been in
contact with the metal walls of the sampler.
3-29
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Figure 3-12. Shipek Grab Sampler (From APIA, 1976)
3-30
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3.2.3.2 Scoops and Drag Buckets
Scoops and drag buckets are designed for general bottom sampling
or exploratory work only. Many designs have been developed depending on
the type of bottom material to be sampled and the depth of water from which
it is to be obtained. All have similar characteristics in that they obtain
a sample by direct insertion and removal, as with a shovel, or by being •
dragged along the bottom for a select distance by a rope or wire. The most
common device is the simple hand scoop used in waters of wadcable depth.
t
The use of scoops or drag buckets for priority pollutant mon-
itoring is not recommended, unless the scoop is enclosed on five sides and
capable of being capped before removal from the sediment. The fine mate-
rials of the sediment surface are easily lost by the action of the sampling
motion or may be washed away as the sample is brought to the surface. The
various devices do not provide for a consistent technique that will ensure
the securing of a truly representative sample, and it is difficult to
quantify the bottom area sampled.
Table 3-1 summarizes the use and advantages and disadvantages of
the sampling equipment discussed above.
3.3 Sample Handling, Preservation, and Shipment
Sediment samples obtained for priority pollutant analysis re-
quire careful handling to avoid the possibility of introducing interfer-
ences, both positive and negative, in the sampling process, in the
containers used for storage, or during transport to the lab for analysis.
Possible routes of positive interference or contamination include residues
on the sampling equipment (such as rust and corrosion products), leaching
of materials from containers, paint leached from the hulls of ships and
boats, and dust and other micro-particles in the sampling environment.
Negative interferences may arise as a result of the adsorption of chemicals
to surfaces of containers or from the breakdown of samples because of
improper preservation procedures. The risks of contamination, adsorption,
and desorption have been reviewed by a number of investigators (Cooper,
1958; Robertson, 1968; Tolg, 1972; NBS, 1974a, 1974b, and 1976). The
purpose of this section is to summarize the currently accepted procedures
3-31
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Table 3-1
Summary of Bottom Sampling Equipment
Device
Us*
Advantage*
01sadvantages
Teflon* or Glass Tub*
Shallow wadeable waters or
waters If SCUBA available. Soft
or semi-consolidated deposits.
Preserves layering and permit* historical
study of sediment deposition. Rapid -
samples Immediately ready for laboratory
shipment. HlnlMl risk of contamination.
Inexpensive.
Small sample size requires
repetitive sampling.
Hand Corer with rwMvabl* Taf Ion*
or ylas* liners
Sam* as abov* «xcopt aor* consoll-
datad s«llMints can b* obtalnad.
ttea a)tt*ndod to Mtars of 4-6 to«t
by tha usa of axtanslon rods.
Hand las prowIda for greater aas* of sub-
strata penetration.
Requires removal of liners
befor* rao*tltlv« sampling,
Slight risk of netaI con-
tamination from barrel and
cora cutter.
Ekaan or Box-Orado»
I
u>
N>
Soft to seal-soft sadlamits. Can
be used fro* boat, .bridge, or plar
In waters of various depths.
Obtains a larger sample than coring tubas*
Can ba subsampled through box lid.
Possible Incomplete Jaw
closure and sample loss.
Possible shock wave which
any disturb the fines.
Metal construction may
Introduce contaminants.
Possible loss of "fines" on
retrieval.
Gravity corers
I.e., Phleger Carer
Deep lakes and rivers. Semi-
consolidated sediments.
Low risk of sample contamination.
Small sample, requires re-
petitive operation and re-
moval of liners. Tina
consuming.
Ponar Grab Sampler
VanVoen Sampler
Deep lakes, rivers, and estuaries.
Useful on sand, silt, or clay.
Most universal grab sampler.
Adequate on mast substrates.
Large sample obtained Intact,
permitting subsampllng.
Shock wave from descent may
disturb "fines". Possible
Incomplete closure of Jaws
and sample loss. Possible
contamination from metal
frame construction. Sample
exist be further prepared
tor analysis.
-------
Table 5-1 (coot»
Summary of Bottom Soap I Ing Equipment
Device
Us*
Advantages
Disadvantages
UHH-S3 Piston Carer
Maters ol 4-6 (not deep when
used Milk extension rod. Soft
to sa« I-consolidated deposits.
Platan provides for greater
•ample retention.
Cores «ust be extruded on
site to other containers -
•ataI barrel Introduces
risk of natal contamination
UHH-60
Sampling moving waters Iroei a
fixed platform.
Streamlined configuration
allows sampling where other
devices could not achieve
proper orientation.
Possible contamination fro*
•atat construction. Sub-
samplIng difficult. Not
effective tor sampling
fine sedlmaats
Hetorsen Grab Sampler
Deep lakes, rivers, and estuaries.
Useful on mo»t substrates.
Large sample} can penetrate
most substrates.
Heavy, may require winch.
No cover lid to permit sub-
sampling. All other disad-
vantages of Ekman and
Ponar.
St.l(i«k Grab
Used primarily la marine waters
and large Inland lakes and
reservoirs.
Sample bucket may be opened to
permit subsampllng. Retains
fine grained sediments effectively.
Possible contamination from
metal construction. Heavy.
may require winch.
Grab
S«lth-Mclntyre Grab
Deep lakes, rivers, and estuaries.
Useful on most substrates.
Designed for sampling hard substrates.
Loss of fines. Heavy - may
require winch. Possible
matal contamination.
Scoops. Oraij Buckets
Various envlronmants depending
on depth and substrate.
Inexpensive, easy to handle.
Loss of fines on retrieval
through water column.
-------
with regard to container selection and cleaning and sample handling,
preservation, and transport that will maintain the integrity of the sample
until analysis.
In general, because of the risk of contamination, the amount of
sample handling should be kept to an absolute minimum. It is recommended,
therefore, that samples be transported in bulk to the laboratory and not
divided into individual groups for specific analysis, such as metals,
volatiles, and-extractable organics. This will not only reduce handling
but also the packaging or container requirements. Core tubes or liners, as
discussed in Section 3.2.1, can be used for sampling as well as for con-
tainers for laboratory shipment. Coring tubes should be manufactured of
high quality borosilicate glass or Teflon*. Plastics should not be used
because they are known to introduce plasticizers, such as phthalate esters,
and also because they can sorb organics to the container walls. Coring
tubes should be equipped with cork or rubber stoppers that have been cov-
ered either with Teflon* sheet or aluminum foil.
An alternative to the use of coring tubes is the use of one-
quart wide-mouth glass jars- with screw cap lids. The cap should also be
lined with Teflon* sheeting or aluminum foil to prevent contamination of
the sample by the cap material. Some USEPA Regions recommend that samples
for volatile organics analysis be collected separately in the field in a
sealed container which can be fitted to a purge and trap device.
The core liners, glass jars, caps, stoppers, Teflon* sheeting,
and aluminum foil should be thoroughly cleaned before the sampling effort.
Since it is recommended that the containers be used for bulk collection and
storage, they must be cleaned with regard to all the parameters on the
priority pollutant list (see Appendix A). The following cleaning procedure
is summarized from Methods for Chemical Analysis of Water and Wastes (EPA,
1979) and the Office of Research and Development proposed analytical pro-
cedures for priority pollutant analysis (EPA, 1980).
1. Wash all coring tubes, containers, caps, Teflon* sheet-
ing and aluminum foil with a non-phosphate laboratory
grade detergent and tap water.
2. Triple rinse with tap water.
3. Rinse with 1:1 nitric acid (HN03 - Reagent grade).
3-34
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4. Rinse with deioniaed-distilled water.
5. Rinse with 1:1 hydrochloric acid (HCL - Reagent grade).
6. Triple rinse wish deionized-distilled water.
7. Rinse with acetone followed by a final rinsing with
pesticide grade hexane.
8. Dry in a contaminant free area such as a laminar flow
hood.
The nitric acid-hydrochloric acid washes are designed to remove trace
metals, and the acetone-hexane rinses are to remove organic impurities
which may interfere with the subsequent priority pollutant analyses. After
drying, the core tubes and containers should be sealed and stored in a
clean area until ready for use. Additionally, although it is impractical
to clean all sampling equipment, common sense doea apply. Sampling equip*
ment that has been obviously soiled by oils, grease, or household and
laboratory solvents should not be used. All equipment should be carefully
stored away from chemical solvents and household items such as paints,
cleansers, and disinfectants.
Core samples are prepared for packaging by capping the ends of
the coring tubes with an appropriately sized Teflon* or aluminum foil cov-
ered stopper as described in Section 3*2.1. The overlying water should be
retained in the core tubes, and they should be sealed with no air space in
order to prevent loss of volatiles during shipment and storage. If samples
are collected by dredging, they should be carefully transferred to a one
quart wide-mouth glass Jar by means of a Teflon* spatula or, as described
in Section 3.2.1, subsampled with coring tubes to minimize the risk of
trace metal contamination from the frame of the sampling device. These
tubes should be capped as described above. When wide-mouth glass Jars are
used they should be filled as nearly to the top as possible and topped off
with sample water and sealed with the Teflon«-llned screw cap. Maximum ef-
fort must be made to seal the sample with a minimum of gaseous headspace to
prevent loss of volatiles. The sample must remain sealed until aliquot3
for volatile organlcs are taken for analysis (EPA, 1977t>).
Each sample container (e.g., core or Jar) should be labeled with
a unique number by which it can be readily identified in the laboratory.
3-35
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This identification number should have as few digits as possible to dis-
courage abbreviation. The label should be waterproof, and all information
should be written with a ballpoint pen in waterproof ink. The labels
should include, in addition to the identification number, the date and the
initials of the sampling personnel.
Other pertinent information such as the time the sample was
taken, location, approximate depth, stratigraphy, and water quality (e.g.,
temperature, DO, pfl) should be recorded in a field notebook. If coring
devices are used, a note should be made of the compaction of the core if
this information is available. Any coring will result in compaction of the
sediments within the coring tube. Usually mud will adhere to the outside
of the corer at some distance above the level of the sediment within the
core. Compaction is frequently two or more to one. An estimate of the
compaction ratio should be noted if observed. All data in this notebook
must, of course, be cross-referenced to the actual sample by using the
identification number as previously discussed.
After it is labeled, the sample should be placed in a cooler or
freezer chest. The cores should be stored in an upright position. If
samples are to be analyzed within 7 days of collection, they should be
maintained at 4*C until analysis (ZPA, 1980). The use of pre-frozen sealed
glycol-based coolant (e.g., "blue ice") is preferred over crushed ice. The
cores and glass jars can be stored in a cooler by alternating rows of
samples and synthetic ice containers. This method of storing sediment
samples for shipment is efficient and results in the least possibility of
contamination. For long-term storage, it is recommended that the samples
be immediately frozen. Preservation with dry ice (frozen C02) is recom-
mended as a means of ensuring that the sample is frozen rapidly and that it
remains frozen. This is very important to prevent decomposition and loss
of volatile materials (FWGPM, 1974; Bruce et. al., 1974; Straughan, 1974).
If freezing is necessary, try to avoid the use of glass containers because
of the possibility of breakage.
Dry ice requires special packaging precautions before shipping
to comply with DOT regulations. The Federal Code of Regulations classifies
dry ice as ORM-A (Other Regulated Material). These regulations specify the
3-36
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amount of dry ice which may be shipped by air transport and the type of
packaging required.
For any amount of dry ice co be shipped by air, advance arrange-
ments must be made with the carrier. Not more than 440 pounds of dry ice
may be shipped by air freight unless special arrangements have been made
previously between the shipper and the aircraft operator. Quantities of
dry ice needed for sample preservation are usually considerably less than
440 pounds.
The regulations further specify that the packaging must be de-
signed and constructed in a manner to permit the release of carbon dioxide
gas which, if restricted, could cause rupture of the package. If samples
are being transported in a cooler, several vent-holes should be drilled to
allow for escape of the sublimated gas. The vents should be near the top
of the vertical sides of the cooler, rather than in the cover, to prevent
debris from falling into the cooler. Furthermore, wire screen or cheese-
cloth should be installed to help keep foreign materials from entering the
vents. When packaging the samples, care should be taken to keep these
vents open to prevent the buildup of pressure.
Dry ice is exempted from shipping paper and certification re-
quirements if the amount is less than 440 pounds and the package meets
design requirements* The package must be marked "Carbon Dioxide, Solid" or
"Dry Ice" and also marked with an identification that the material being
refrigerated is to be used for diagnostic or treatment purposes (e.g.,
frozen samples).
Upon receipt at the laboratory, the sediment samples should be
placed in a refrigerator and maintained at 4*C or, if frozen, placed in a
freezer and maintained at a temperature less than -20*C until samples are
prepared for analysis. All sediment samples should be kept in their
original containers until they are ready to be prepared for analysis.
When the samples are received at the laboratory, they should be
recorded in a permanent log book. This log book should include for each
sample date and time received, source of sample, sample number, mode of
transportation to the laboratory, and the number assigned to the sample by
the laboratory if this number differs from the field number. Although this
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recording procedure may seem laborious, it is absolutely imperative that
precise records be kept for all samples so that the data generated by the
sampling and analysis effort is of unquestionable integrity.K
An accurate written record should be maintained which can be
used to trace possession of the sample from the moment of its collection
until it has been analyzed. A chain of custody tag should be placed on all
coolers in which samples are stored and shipped. This should have appro-
priate spaces for signatures when the sample is transferred from one person
to another. The date and time at which the custody is transferred should
be Indicated on the tag.
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3.4 References
American Public Health Association (APHA). 1976. Standard Methods
for the Examination of Water and Wastewater. 14th Ed. Washington,
D.C. 1193 p.
Bruce, H.E., and S.P. Gran. 1974. Sampling Marine Organisms and
Sediments for High Precision Gas Chromatographic Analysis of Aromatic
Hydrocarbons, jn Marine Pollution Monitoring. Proceedings of a
Workshop held at National Bureau of Standards. Gaithersburg, MD.
(NBS Special Publication 409: 181-182).
Bouma, A.H. 1969. Methods for the Study of Sedimentary Structures.
John Wiley & Sons, N.7. pp. 301-380.
Cooper, L.H.N. 1958. A System for International Exchange of Samples
for Trace Element Analysis of Ocean Water. Journal of Marine Re-
search. 17: 128-32.
Federal Interagency Sedimentation Project. 1940. Equipment Used for
Sampling Bedload and Bed Material. U.S. Interagency Report No. 2.
University of Iowa Hydraulics Laboratory. Iowa City, Iowa.
Federal Working Group on Pest Management (FWGPM). 1974. Guidelines
on Sampling and Statistical Methodologies for Ambient Pesticide Mon-
itoring. National Technical Information Service. U.S* Department of
Commerce. Washington, D.C. PB-239-798.
Feltz, H.R. 1980. Significance of Bottom Material Data in Evalua-
ting Water Quality. In Contaminants and Sediments, Vol. I. R.A.
Baker (ed.). Ann Arbor Science Publications. Ann Arbor, Mich. pp.
271-287.
Feltz, H.R., and J.K. Culbertson. 1972. Sampling Procedures and
Problems in Determining Pesticide Residues in the Hydrologic Environ-
ment. Pesticide Monitoring Journal. 6(3):171-178.
Flannigan, J.F. 1970. The Efficiencies of Various Grabs and Corers
in Sampling Freshwater Benthos. Journal of the Fisheries Research
Board of Canada. 27(10):1691-1700.
Frank, R., M. Holdrinet, H.E. Braun, R.L. Thomas, A.L.W. Kemp, and
J.M. Jaquet. 1977. Organochlorlne Insecticides and PCBs in
Sediments of Lake St. Clair (1970 and 1974) and Lake Erie (1971). The
Science of the Total Environment. 8:205-227.
Greig, R.A., R.N. Reid, and D.R. Wenzloff. 1977. Trace Metal Con-
centrations in Sediments from Long Island Sound. Marine Pollution
Bulletin. 8(8):183-188.
Greig, R.A., and R.A. McGrath. 1977. Trace Metals in Sediments of
Rarltan Bay. Marine Pollution Bulletin. 8(8):188-190.
3-39
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Guy, H.P., and V.W. Norman. 1969* Field Methods for the Measurement
of Fluvial Sediment. In Techniques of Water Resources Investiga-
tions, Book 3, Chapter C2. U.S. Geological Survey. Res ton, 7A.
Holliday, B.W. 1978. Processes Affecting the Fate of Dredged Mater-
ial. U.S. Army Engineer Waterways Experiment Station. Technical Re-
port DS-78-2. Vicksburg, Miss.
Hopkins, T.L. 1964. A Survey of Marine Bottom Samplers, jn M.
Sears, (ed.), Progress in Oceanography, Volume II. Pergamon Press,
N.7. pp. 215-253.
Hough, J.L. 1939. Bottom Sampling Apparatus, ^n P.O. Trask, (ed.),
Recent Marine Sediments. Dover Publications, Inc., N.Y. pp.
632-664.
Howmiller, R.P. 1971. A Comparison of the Effectiveness of Ekman
and Ponar Grabs. Transactions of the American Fisheries Society.
100(3): 560-564.
Hudson, P.L. 1970. Quantitative Sampling with Three Benthic
Dredges. Transactions of the American Fisheries Society. 99(3):
603-607.
Jackson, H.W. 1970. A Controlled Depth, Volumetric Bottom Sampler.
Progressive Fish Culturist. 32 (2): 113-115.
Kemp, A.L.W., H.A. Savile, C.B. Gray, and A. Mudrochova. 1971. A
Simple Corer and Method for Sampling the Mud-Water Interface.
Limnology and Oceanography. 16(4): 689-694.
Larimore, R.W. 1970. Two Shallow-Water Bottom Samplers. Progres-
sive Fish Culturist. 32(2): 116-119.
Llnd, O.T. 1974. Handbook of Common Methods in Limnology. The C.V.
Mosby Co. St. Louis, Mo.
Maitland, P.S. 1969. A Simple Corer for Sampling Sand and Finer
Sediments in Shallow Water. Limnology and Oceanography.
Paterson, C.G., and C.H.. Fernando. 1971. A Comparison of a Simple
Corer and An Ekman Grab for Sampling Shallow-Water Benthos. Journal
of the Fisheries Research Board of Canada. 28(3):365-368.
Robertson, D.E. 1968. Role of Contamination in Trace Element
Analysis of Sea Water. Analytical Chemistry. 40(7): 1067-1072.
Schwoerbel, J. 1974. Methods of Hydrobiology (Freshwater Biology).
Pergamon Press. Oxford, England.
3-40
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Straughan, D. 1974. Field Sampling Methods and Techniques for
Marine Organisms and Sediments, Marine Pollution Monitoring. In
Proceedings of a Workshop held at National Bureau of Standards.
Gaithersburg, MD. (NBS Special Publications 409).
Tolg, G. 1972. Extreme Trace Analysis of the Elements - I: Methods
and Problems of Sample Treatment, Separation and Enrichment.
Talanta. 19:1489-1521.
U.S. Environmental Protection Agency. 1977a. Analysis of Pesticide
Residues in Human and Environmental Samples. Health Effects Research
Laboratory. Office of Research and Development. Research Triangle
Park. Horth Carolina.
U.S. Environmental Protection Agency. 1977b. (Revised October
1980). Interim Methods for the Sampling and Analysis of Priority
Pollutants in Sediments and Fish Tissue. Environmental Monitoring
and Support Laboratory. Office of Research and Development.
Cincinnati, Ohio.
U.S. Environmental Protection Agency. 1979. Methods for the
Chemical Analysis of Water and Wastes. Environmental Monitoring and
Support Laboratory. Office of Research and Development. Cincinnati,
Ohio.
U.S* Environmental Protection Agency. 1980. Draft Protocols for the
Analysis of Priority Pollutants. Methods 601-613, 624 and 625.
Monitoring Technology Division. Office of Research and Development.
Washington, D.C.
U.S. Geological Survey. 1980. National Handbook of Recommended
Methods for Water-Data Acquisition. Office of Water Data Coordi-
nation* Reston, VA.
U.S. National Bureau of Standards. 1974a. Sampling, Sample
Handling, and Analysis. Symposium on Accuracy in Trace Analysis.
Proceedings of the 7th IMR Symposium. P.D. Lafleur, Ed. UBS Special
Publication 422. Washington, D.C.
U.S. National Bureau of Standards. 1974b. Marine Pollution
Monitoring (Petroleum). Proceedings of a Workshop. NBS Special
Publication 409. Washington, D.C.
U.S. National Bureau of Standards. 1976. A Survey of Current
Literature on Sampling, Sample Handling, and Long Term Storage for
Environmental Materials. U.S. Department of Commerce. Washington,
D.C.
Weber, C.I., editor. 1973. Biological Field and Laboratory Methods
for Measuring the Quality of Surface Waters and Effluents. U.S.
Environmental Protection Agency. Office of Research and Development*
Cincinnati, Ohio. EPA 670/4-73-001.
3-41
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Welch, P.S. 1948. Llmnological Methods. The Blakiaton Company.
Fniladeiphia, ?A. pp. 175-186.
Wigley, R.L. 1967. Comparative Efficiencies of VanVeen and
Smith-Mclntyre Grab Samplers as Revealed by Mocloa Pictures.
Ecology. 48(1): 168-169.
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4.0 SHELLFISH SAMPLING
Shellfish monitoring can provide Important information on distribu-
tion and occurrence of pollutants. Shellfish live on or in the substrate
and nay accumulate pollutants from either water or sediment. They are also
an important food item for some "higher" aquatic organisms. Therefore,
they constitute an interface among the sediment, water column, and biotic
components of aquatic and estuarine systems. la terms of equipment and
study design, shellfish sampling shares some characteristics with both
sediment and fish sampling. Some of the equipment used in sediment sam-
pling can be used to collect bivalves; techniques for site selection and
sample preservation and handling are similar to those used in fish sam-
pling. These similarities are evident on comparing this chapter with Che
respective sections of Chapters 3 and 5.
There are two major assumptions governing the techniques recommended
in this chapter: (1) shellfish will be sampled in estuaries only, and (2)
target species will be limited to bivalve molluscs (pelecypods). The rea-
son for limiting shellfish sampling to estuaries is that in fresh water,
fish are preferable because of their wider distribution, potential for
consumption by humans, and more convenient size (many of the fresh water
shellfish are small and would take intensive collecting and identifying
efforts to provide the minimum 500 gram tissue sample size). On the other
hand, in estuaries, many of the fish are anadromous or catadromous and
tissue concentrations may not reflect environmental conditions at the site
sampled. Some of the common estuarine bivalves offer an excellent alterna-
tive to fish because they are sessile, are fairly easy to identify and
collect, and are consumed by humans.
The reason for limiting shellfish sampling to bivalves is primarily
that they are sessile (with the exception of some scallops) and therefore
cannot move out of polluted areas as can gastropods and the crustacean
shellfish. The filter feeding employed by bivalves also exposes them to
large quantities of water thereby increasing the opportunity for uptake of
pollutants. These two factors result in bivalves being rapid indicators of
pollution and highly representative of the areas from which they are
sampled.
4-1
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There are, however, several problems encountered when using bivalves
as indicators of pollution. Aa with any organisms used to indicate pol-
lution, uptake and depuration varies widely among species. This is the
principal reason for restricting the number of target species. The large
volume of water pumped by bivalves in their filter feeding has disadvan-
tages as well as advantages. One major disadvantage is that sporadic
episodes, of pollution may not show up aa readily in these species as they
would with finfish. The reason for this is that depuration tends to occur
more rapidly in these molluscs because of the large amount of water they
process. Excretion is often nearly as rapid as uptake.
Another related problem is the tendency of many bivalves to accumu-
late appreciable quantities of silt in their digestive system. This may
result in high concentrations of metals and other pollutants if the whole
organism (not including the shell) is analyzed. If the intent of the study
is to determine concentrations available for human exposure, this is per-
haps a more representative case since the whole organism is often consumed,
including the stomach and contents. On the other hand, if the intent is to
determine "tissue" levels of pollutants, this factor can bias the results
giving values higher than the actual tissue concentrations.
Aa discussed in the following chapter on fish, methods can be adopted
in developing a sampling program to reduce the difficulty in interpreting
data. These include establishing target species for collection (thereby
eliminating interspecific variations), sampling shellfish of similar age
and size, and limiting sampling at different sites to as short a period as
possible to reduce seasonally-related differences. Analytical techniques
should also be used to improve the utility of the data. These include
analysis of lipid content, analysis of water content, quality control (such
as analysis of duplicate samples), and compositing tissues from five or
more individual bivalves. Analytical techniques are currently being devel-
oped by EPA1s Environmental Support Laboratory in Cincinnati, Ohio, and are
provided in a separate document.
4.1 Site Selection
Maximum benefits can be obtained from a sampling program by
giving careful attention to the process of site selection. Factors which
4-2
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play an Important role la site selection include:
1. Purpose of sampling program.
2. Presence of shellfish at the site.
3. Proximity of sites for sampling water, bed sediment,
and fish for priority pollutants.
4. Previous shellfish sampling for priority pollutant
analysis.
5* Type of equipment available.
6. Accessibility of site.
Each of these factors is discussed in greater detail in the following
paragraphs•
Since bivalve shellfish sampling is recommended only for es-
tuarlne areas, there are basically two types of areas related to the pur*
pose of the sampling program. If the program's goal is to provide an over-
all evaluation of pollutant levels, the sites should be located in open
water, characteristic of the overall estuary. On the other hand, if the
goal is to identify sources of pollutant input, the sites should be select-
ed near river mouths or suspected sources of pollutant discharge.
The presence of bivalves at the site is an obvious requirement.
Finding a suitable site is often less obvious. In many cases, the State
Department of Natural Resources (or other related agencies) maintains maps
showing the location of major concentrations of commercially important
shellfish. Consulting maps of this sort or commercial fishermen as to the
location of large beds of shellfish can save a lot of time and eliminate
the hit-or-miss sampling technique. Beware of the fact that some shellfish
beds are leased from state governments. Before sampling such beds, secure
any necessary permits and the consent of the lessee.
There are advantages to locating the shellfish sampling sites
near sites selected for priority pollutant sampling of water, bed sediment*
or finfish. The principal benefit of this sampling design is the possibil-
ity of developing at least a simple model of the dynamic distribution of
pollutants in that area. Correlations among pollutant levels in the dif-
ferent compartments (finfish, shellfish, sediment, and water) may be es-
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tablished. Selecting sites close to each other also allows a more
efficient use of time by providing the opportunity to combine sampling
trips.
The availability of historical data on pollutant body burdens
should be checked before making any final decisions regarding sample sites.
In most cases these data are nonexistent, but in some areas such sampling
has been conducted previously. In those fev cases where this information
is available, consideration should be given to choosing sampling sites near
the areas sampled previously in order to construct a historical record of
pollution in the area. Closed shellfish areas often have historical data.
In addition, they are presumably close to pollution sources and contain
many large, old bivalves which may have had the greatest opportunity to
accumulate high concentrations of pollutants in their tissues.
The availability of equipment and personnel experienced in its
use is also an important consideration when selecting sample sites. If a
site which meets the other criteria cannot be sampled with available
equipment, there is often the option of coordinating efforts with other
biologists or commercial fishermen who have the appropriate equipment and
experience.
Another factor closely related to the equipment availability is
the depth and accessibility of the site. If hand tongs, a rake, or similar
equipment are to be used, the water depth at the site should be 6 meters
(20 feet) or less. In any case, choosing a representative site is more
important than choosing a site which is highly accessible.
The best approach to developing a sound sampling program is to
consult with local biologists regarding the best candidate species and
areas in which those species can be located. Before sampling, a reconnais-
sance should be performed to locate sampling sites. Once again, it is
vital to arrange for any necessary collection permits before implementing a
sampling program.
4.2 Target Species
The principle of selecting "target" species is designed to limit
the number of variables and facilitate comparison of results from different
4-4
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monitoring studies* Because of the great diversity of estuarine habitats
in the United States, it is impossible to select a single target species
for all monitoring studies. An attempt should be made, however, to
concentrate on a few widely distributed species.
Characteristics of a good target species include:
1. Wide range (e.g., broad distribution).
2. Easy identification.
3* Pollution tolerance.
4. Commercial importance.
Wide range is important since it permits sampling the same
species in various areas, thereby facilitating direct comparison between
studies. Easy identification is a practical factor in sampling since
identification of some pelecypod species requires careful examination of
minute details. Pollution tolerance is an important consideration since
many of the sites of interest may be polluted to some extent. Commercially
important species are preferred because of their larger size and the fact
that they represent an important route of human exposure to pollutants.
It is recommended that at least five shellfish of the same
species be collected per site. Each mollusc should be wrapped individually
and then placed in a plastic bag containing all shellfish of the same
species from the same site, as described in Section 4.4. The minimum
sample mass (per species) is 500 grams of tissue. It is important to note
that the shell of bivalve molluscs represents a considerable percentage of
the total organism weight. Therefore, several molluscs should be removed
from their shells and tissue weights taken to determine the size needed to
obtain the required 500 grama. If five molluscs do not weigh an aggregate
500 grams, more should be collected until this minimum is reached.
If it is not possible to collect one of the recommended species,
congeneric species should be collected if available. ' It Is always a good
policy to collect the same species throughout a sampling program; however,
this is not always possible. The most important principle is to be
consistent.
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The shellfish listed In Table 4-1 satisfy most of the criteria
listed previously for selecting target species* This list is not intended
to be exclusive, but should serve as guidance for selecting bivalves to
sample.
Table 4-1
Target Species For Bivalve Shellfish
Eastern Oyster (Craaaostrea virginica)
Native Pacific Oyster (Ostrea lurida)
Common Blue Mussel (Mytilus edulis)
Northern Quahog or Hard-shell Clam
(Mereenaria mereenaria)
Soft-shell Clan (Mya arenaria)
4.3 Sampling Equipmentand Use
Bivalves, such as clans, scallops, mussels and oysters, are
typically found burled in benthic substrates or attached to submerged ob-
jects such as rocks and piers. Various types of sampling equipment or
tools have been developed for the purpose of collecting these bivalves,
depending on the type of environment or physical conditions in which the
particular target organism is found* Some bivalves such as the eastern
oyster (Crassostrea virginiea) and the hard clam (Mereenaria mereenaria)
enjoy wide popularity because of their edible and abundant nature. Commer-
cial fisheries have developed sophisticated sampling equipment for each of
these particular species in order to maximize the catch efficiency.
Sophisticated methods and equipment have also been developed for the
purpose of quantifying the biomass of particular bivalves.
When collecting bivalves for the purpose of screening
pollutants, the sampling approach is generally qualitative and the equip-
ment need not be extremely sophisticated since only a small sample size is
required. The equipment, however, should be specifically adapted to the
4-6
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target species being sampled and the type of environment in which they are
to be found. The specific considerations that are important in the selec-
tion of sampling equipment include (1) the size of the target species to be
sampled; (2) the number of organisms required in the sample; (3) t'he type
of substrate in which the target species is found; and (4) the depth of.
water in which the target species is located. Some of the equipment useful
for sampling bivalves was discussed In the sediment sampling section of
this manual. A brief review is offered in this section, however, for the
purpose of discussing equipment use and limitations when sampling for bi-
valves. This section will also introduce various other types of equipment
that are applicable for sampling particular target species.
Regardless of the equipment used, a minimum of 500 grams (1.1
Ibs) of body tissue (not including shell) should be collected for priority
pollutant analysis. It is recommended that the average tissue weight of
the selected size of specimens to be retained for analysis be established
prior to actual sample collection. This will preclude the removal and
sectioning of tissue material in the field which would increase the risk of
sample contamination. Prior Icnowledge of average tissue weight per speci-
men size will facilitate determination of the actual number of individual
samples required for subsequent priority pollutant analysis.
4.3.1 Mechanical Grabs
Section 3.2.2 of the sediment sampling portion of this manual
discusses the various types of mechanical grab (or dredge) samplers that
have been developed to sample different sediment types. All are similar la
that they are devices with jaws which are forced shut by the action of
weights, lever arms, springs, or cord. Many of the devices discussed in
that section are applicable to sampling the various types of bivalves. A
brief discussion of the advantages and disadvantages in collecting pele-
cypods is offered according to the major categories within which the de-
vices can be grouped. These categories include (1) the pole operated grab
buckets and tongs and (2) the line operated grab buckets.
4.3.1*1 Pole Operated Grab Buckets and Tongs
Pole operated grab buckets and tongs are probably the most ef-
ficient means for sampling bivalves such as clams and oysters that are
4-7
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located on or slightly buried in bottom sediments. The various models are
all designed to be used in shallow water areas of less than 6 meters (20
feet) either from a boat or structure such as a pier projecting over the
*
water surface. At depths greater than 6 meters (20 feet), the pole oper-
ated devices generally become too difficult to manually operate.
Pole operated grab device efficiency is attributable to the com-
plete control the operator has over the placement of the jaws on the
substrata. It also allows the direct application of the needed energy for
substrate penetration if the target species is buried. Single-pole operated
grab devices, such as the Eloaan or Orange-Feel grabs, are not as efficient
as the double-pole or scissor-like grab devices. This is because the
single-pole grab whose jaws are generally operated by some type of spring
loaded mechanism must be repeatedly retrieved and emptied. These devices
are also generally small and not suitable for collecting large specimens
such as oysters and hard clams.
The double handled grab samplers or "tongs," as they are fre-
quently called, are a much more efficient tool for sampling bivalves in
shallow water, lakes, rivers, and estuaries. Essentially the "tong" is
composed of a pair of long poles fastened together near their distal ends.
The end of each pole has claws or baskets, and the two sets are brought
together by closing the handles together like a pair of scissors. Since
the collection of surrounding or overlying sediments is not required when
sampling for bivalves, the jaws are generally open baskets. This not only
reduces the weight of the device but also allows the washing of collected
specimens to remove the mud, clay, or other sediments in which they are
found. Experienced "totigers" can also feel, via the poles, whether samples
have been collected avoiding the necessity of repeated jaw retrieval for
inspection. Tongs are frequently used in the commercial industry for
gathering oysters, hard clams and scallops.
• 4.3.1.2 Line or Cable Operated Grab Buckets
Included in this category are the various types of mechanical
grabs discussed under sediment sampling such as the Ekman, Petersen, Ponar
and Orange-Peel dredges. These devices are generally used for sampling
benthlc substrates in deep lakes, rivers, and estuaries as discussed in the
4-8
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sediment sampling section of this manual (Section 3.2.2). Besides sediment
sampling, these devices are frequently used for quantitative sampling of
small benthic macroinvertebrates. These devices, however, are not partic-
ularly effective for collecting the large pelacypods such as oysters and
clams. The limitations include the small size of the jaws, incomplete
closure of the jaws caused by the jamming action of shells and other
debris, lack of penetration into substrates to the depths at which many
bivalves may be found, and the relative inefficiency of repeated raising
and lowering of the device to inspect the sample containers for success of
capture. The only notable exception to the various line operated grab
samplers are the patent tongs commonly used in the Chesapeake Bay area for
collecting oysters. This device is a variation of the previously discussed
hand tongs; for this version, their depth effectiveness has been increased
by the use of pulleys. The jaws or baskets are attached to short handles
that are lowered by a rope which runs through a block on the upper end of
one handle, then across and through a block on the upper end of the oppo-
site handle, where it runs down to the base. Pulling on the rope closes
the tongs or jaws together. This device can be operated by hand-line; how-
ever, more frequently it is done from a boat supplied with a winch. Since
it is unlikely that most priority pollutant investigators would have access
to this equipment, it is generally not a practical equipment alternative.
4.3.2 Biological Dredge
Biological dredges are heavy duty nets or bags which are dragged
along the bottom of a deep water body to collect stones, bottom debris, or
large stationary macroinvertebrates such as clams, scallops, and oysters.
Unlike the line operated mechanical grabs which sample a unit area and can
be used for quantitative sampling, the tow dredges are generally an unso-
phisticated tool for simple qualitative sampling of bottom substrates. As
such, biological dredges are a valuable tool for securing the samples
required for priority pollutant studies.
Many types of biological dredges or drag nets are available de-
pending on the physical nature of the substrate and/or the organism to be
sampled. These devices are made of heavy metal frames, frequently with
rake-like teeth on the lower edge to which is attached a heavy duty bag of
4-9
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canvas or nat possibly made of chain or wire. The dredge is pulled along
the bottom by means of a tow line. Often the natal frame is designed to
dig into the bottom sediments, dislodging buried bivalves as it is pulled
along* The length of tow depends upon the site of the dredge and prior
knowledge of the density of organisms to be found in the area to be sam-
pled. Since only a small number of organisms are needed, the length of tow
should only be long enough to obtain the required number of bivalves neces-
sary for analysis. It will be difficult to pinpoint the exact area of sam-
ple collection unless the length of tow is relatively short. Because of
the scouring operation of biological dredges which may damage the shells of
bivalve specimens, It is important that all specimens selected for priority
pollutant analysis be inspected. Damaged specimens should be discarded,
thus preventing the chance of contamination of the tissues.
4.3.3 Coring Devices
The various types of coring devices discussed in the sediment
sampling section of this manual are not suitable for collecting bivalve
specimens unless the target species are very small and are known to Inhabit
a specific area in a relatively high density. The limited diameter of
coring devices makes them unsuitable as a tool for the collection of
organisms such as clams, mussels, and oysters.
4.3.A Miscellaneous Devices
In addition to the various types of equipment previously de-
scribed there are many other kinds of devices for collecting bivalve speci-
mens that do not readily conform to the established categories. These
various types of tools are discussed in the following subsections.
4.3.4.1 Scoops or Shovels
Scoops or shovels can frequently be used for collecting bivalves
buried in shallow water sediments accessible by wading or SCDBA equipment.
The hard clam or quahog (Mereenarla mercanaria) and the soft-shell clam
(Mya arenaria) as well as many other shallow water bivalves can be easily
collected using some form of scoop or shovel. The scoop is simply used to
scrape or dig away the overlying sediment or to dislodge the buried organ-
isms. Care must be exercised so as not to damage the shell of desired
4-10
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specimens and ehus introduce possible contaminants to the tissue of the
organisms. Shovels and scrapers can also be used to dislodge bivalves,
such as mussels or oysters, that are attached to the surface of submerged
objects.
4.3.4.2 Rakes
Various types of rakes have been designed to collect bivalves
that lie on the bottom substrate or are buried in shallow water sediments.
These rakes are usually long-handled and can be operated while the sampler
is wading or can be used in shallow waters from a boat. Many of the types
of "clam" rakes are frequently designed or constructed with a catch basket
made of wire or heavier gage metal to maximize the catch efficiency. The
mesh size of the basket is determined according to the size of the target
organism desired. The rakes are simply operated by "raking" the benthic
substrate or, if the target species is buried, by a digging and raking
action to force the dislodged organisms into the catch basket. Bakes can
also be used to dislodge organisms such as oysters and mussels that are
attached to submerged objects such as rocks and the pilings of piers.
A variation of a rake is the scraper with a handle to which is
attached a net or canvas bag. The scraper is used to dislodge specimens,
and the net is used to capture them before they sink or are carried away by
any current that might be present.
4.3.4.3 Dip Wets and Other Assorted Devices
Dip nets made of wire or heavy twine are also acceptable sam-
pling tools in shallow water when the target organisms can be visually
identified. Grappling hooks, pocket knives, pry bars, etc., have all been
used as effective methods for collecting bivalves in different areas and
under different conditions. Regardless of the collection technique chosen,
the investigator or sampling personnel must exercise care in the sampling
process so as not to damage the specimens, which could cause contamination'
4.3.5 Purchasing Specimens/Coordinated Sampling
The most cost-effective and efficient means for obtaining bi-
valve specimens for priority pollutant analysis when sampling equipment
4-11
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and time are limited is to coordinate sampling trips with other types of
sampling investigations or to obtain the needed specimens from a commercial
fisherman* Local federal agencies such as the Fish and Wildlife Service
and the Bureau of Commercial Fisheries, and state agencies such as the De-
partments of Fish and Game and Natural Resources, conduct routine sampling
programs* The priority pollutant investigator need only accompany these
fishery biologists or sampling personnel to acquire the needed specimens
for subsequent1 priority pollutant analysis. This coordination will greatly
reduce the cost for equipment and provide for a sharing of Information that
may be beneficial to all Investigators involved.
In areas where large commercial operations are ongoing, the pri-
ority pollutant investigator may consult with local commercial fishermen to
assist them in obtaining the specimens necessary for analysis. If this ap-
proach is considered, the investigator must accompany the commercial fish-
ermen and should remove the sample from the collection device* This will
not only ensure the proper handling of specimens for subsequent priority
pollutant analysis, but will also ensure accurate recording of exact time
and place the sample was obtained.
4.3.6 Summary
In summary, the particular equipment or methods selected to ob-
tain bivalve samples for priority pollutant tissue analysis depend on a
number of factors. These factors include the type of bivalve, depth of
water in which it is to be found, and the nature of the substrate from
which it will be removed. Because pelecypods are found in a wide variety
of habitats and because they are sessile, the specific collection device or
sampling equipment should be based on substrate, water depth, and size of
the target organisms. As a general rule, however, pelecypods can be sam-
pled in shallow waters from a boat or with the use of pole-operated mechan-
ical grabs or tongs. Where wading is possible the use of clam rakes as a
collection tool may be desirable. When investigators are sampling deep
water estuaries, the use of biological tow dredges is preferred over line
operated mechanical grabs. The most efficient sampling effort is the
coordination of effort with other investigative personnel in order to
reduce costs, equipment needs, and time.
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4.4 Sample Handling, Preservation, and Shipment
Bivalve samples obtained for priority pollutant analysis re-
quire careful handling to avoid the possibility of contamination either in
•
the sampling process or in transport to the lab for analysis. Initiallyt
all samples should be thoroughly rinsed with site water to remove any
surrounding sediment deposits. After rinsing, all samples should be care-
fully inspected for damage in the sampling process before packaging for
laboratory transport. Shells or body tissue that have been damaged by the
sampling equipment should be discarded. Additionally, although it is un-
likely that the bivalves will be contaminated by the sampling equipment it-
self, common sense does apply. Sampling equipment that has been obviously
soiled by oils, grease, or solvents should not be used.
Specimens should not be removed from their shells in the field,
except to weigh a few to ensure that minimum sample size has been attained.
After the samples are frozen (as described below) they can be excised from
their shells at the lab. This reduces the chances of contamination in the
field.
After the selected samples have been rinsed and inspected, they
should be placed in either clean Teflon* bags, such as the air sample bags
manufactured by Pollution Measurement Corporation, or wrapped in clean
heavy duty aluminum foil. Teflon* bags are the ideal packaging material;
however, they have the disadvantage of high cost. Polyethylene or polypro-
pylene packaging materials, as well as all other plastic containing mate-
rials, should not be used because they can introduce contaminants (e.g.,
phthalate esters) to the sample. If aluminum foil is selected as a pack-
aging material, it should be cleaned beforehand by rinsing with acetone,
rinsed again with pesticide grade hexane, and allowed to dry in a contami-
nant free area* Samples should be completely wrapped in aluminum foil and
then sealed in polypropylene bags to retain moisture.
Each sample container (e.g., bag or foil) should be labeled with
a unique number by which it may be readily Identified in the laboratory.
This identification number should have as few digits as possible to dis-
courage abbreviation. The label should be waterproof, and all information
should be written with a ballpoint pen in waterproof ink* The labels
4-13
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should Include, In addition to the identification number, the date and the
initials of the sampling personnel.
Other pertinent information such as the time the sample was
taken, location, approximate depth, species, substrate, and water quality
(e.g., temperature, DO, pH) should be recorded in a field notebook. The
data in this notebook must, of course, be cross-referenced to the actual
sample by using the identification number as previously discussed.
After labeling, the sample should immediately be placed in a
freezer chest with dry ice. Preservation with dry ice (frozen C02) is
recommended as a means of ensuring that the sample is frozen rapidly and
that it remains frozen. This is very important to prevent decomposition
and loss of volatile materials. Minimum deterioration occurs if biological
samples are frozen immediately after death and properly packaged to prevent
loss of moisture and the entrance of oxygen into the tissues (Royce, 1972;
EPA, 1980). Dry ice is the most effective means of rapidly freezing tissue
during field sampling.
Dry ice requires special packaging precautions before shipping
to comply with DOT regulations. The Code of Federal Regulations classifies
dry ice as ORM-A (Other Regulated Material). These regulations specify the
amount of dry ice which may be shipped by air transport and the type of
packaging required.
For any amount of dry ice to be shipped by air, advance arrange-
ments must be made with the carrier. Not more than 440 pounds of dry ice
may be shipped by air freight unless special arrangements have been made
previously between the shipper and the aircraft operator. Quantities of
dry ice needed for tissue preservation are usually considerably less than
440 pounds.
The regulations further specify that the packaging must be de-
signed and constructed in a manner to permit the release of carbon dioxide
gas which, if restricted, could cause rupture of the package. If samples
are being transported in a cooler, several vent holes should be drilled to
allow sublimated gas to escape. The vents should be near the top of the
vertical sides 'of the cooler, rather than in the cover, to prevent debris
4-14
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from falling into the cooler. Furthermore, wire screen or cheesecloth
should be installed to help keep foreign materials from entering the vents*
When the samples are packaged, care should be taken to keep these vents
open to prevent the buildup of pressure.
Dry ice is exempted from shipping paper and certification re-
quirements if the amount is less than 440 pounds and the package meets de-
sign requirements. The package must be marked "Carbon Dioxide, Solid" or
"Dry Ice" and also marked with a statement indicating that the material
being refrigerated is to be used for diagnostic or treatment purposes
(e.g., frozen tissue).
An alternative to dry ice is to enclose the bivalves in a
waterproof bag and ship them in a cooler with wet ice (^0). Bivalves
can survive several days at these temperatures and remain closed and
inactive.
Upon receipt at the laboratory, the tissue samples should be
placed in a freezer and maintained at a temperature less than -20"C (Royce,
1972; EPA, 1980) until the samples are prepared for analysis. All tissue
samples should be kept in their original packaging until they are ready to
be prepared.
When the samples are received at the laboratory, they should be
recorded in a permanent log book. This log book should include for each
sample date and time received, source of sample, sample number, mode of
transportation to the laboratory, and the number assigned to the sample by
the laboratory if this number differs from the field number. Although this
recording procedure may seem laborious, it is absolutely imperative that
precise records be kept for all samples so that the data generated by the
sampling and analysis effort is of unquestionable integrity.
An accurate written record should be maintained which can be
used to trace possession of the sample from the moment of its collection
until it has been analyzed. A chain of custody tag should be placed on all
coolers in which samples are stored and shipped. This should have appro-
priate spaces for signatures when the sample is transferred from one person
to another. The date and time at which the custody is transferred should
be indicated on the tag.
4-15
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4.5 References
American Public Health Association (APHA). 1976. Standard
Methods for the Examination of Water and Wastewater. 14th
Ed. Washington, D.C. 1193 p.
Feltz, H.R. and J.K. Colbertson. 1972. Sampling Procedures
and Problems in Determining Pesticide Residues in the
Rydrologic Environment. Pesticide Monitoring Journal.
6<3):171-178.
Flannigan, J.F. 1970. The Efficiencies of Various Grabs
and Corers in Sampling Freshwater Benthos. Journal of the
Fisheries Research Board of Canada. 27(10):1691-1700.
Hopkins, T.L. 1964. A Survey of Marine Bottom Samplers.
In M. Sears, (ed.), Progress In Oceanography, Volume II.
Pergamon Press, N.Y. pp. 215-253.
Hough, J.L. 1939. Bottom Sampling Apparatus. Jta P.D.
Trask, (ed.), Recent Marine Sediments. Dover Publications,
Inc., N.Y. pp. 632-664.
Howmiller, R.P* 1971. A Comparison of the Effectiveness of
Ekman and Ponar Grabs. Transactions of the American
Fisheries Society. 100(3):560-564
Hudson, P.L. 1970. Quantitative Sampling with Three Benthlc
Dredges. Transactions of the American Fisheries Society.
99(3):603-607.
Larlmore, R.W. 1970. Two Shallow-Water Bottom Samplers.
Progressive Fish Culturist. 32(2):116-119.
Lind, O.T. 1974. Handbook of Common Methods in Limnology.
The C.V. Mosby Co. St. Louis, Mo.
Rounsefell, G.A., and V*H. Everhart. 1953. Fishery
Science: Its Methods and Applications. John Wiley & Sons,
Inc., New York.
Royce, W.F. 1972. Introduction to the Fishery Sciences.
Academic Press Inc., New York. pp. 214-214, 284-295.
Schwoerbel, J* 1974. Methods of Hydrobiology (Freshwater
Biology). Pergamon Press, Oxford, England.
U.S. Environmental Protection Agency. 1977a. Analysis of
Pesticide Residues In Human and Environmental Samples.
Health Effects Research Laboratory. Office of Research and
Development. Research Triangle Park, North Carolina.
4-16
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U.S. Environmental Protection Agency. 1977b. (Revised
October 1980). Interim Methods for the Sampling and
Analysis of Priority Pollutants in Sediments and Pish
Tissue. Environmental Monitoring and Support Laboratory.
Office of Research and Development. Cincinnati, Ohio.
U.S. Environmental Protection Agency. 1980. Draft Proto-
cols for the Analysis of Priority Pollutants. Methods 601 -
613, 624 and 625. Monitoring Technology Division. Office
of Research and Development. Washington, D.C.
Weber, C.I., editor. 1973. Biological Field and Laboratory
Methods for Measuring the Quality of Surface Waters and
Effluents. U.S. Environmental Protection Agency. Office of
Research and Development. Cincinnati, Ohio. 670/4-73-001.
Welch, P.S. 1948. Limnological Methods. The Blaklston
Company, Philadelphia, PA. pp. 175-186.
4-17
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5.0 SAMPLING FISH
Pish can be used to monitor toxic pollutants because of the tendency
of fish to bioconcentrate many kinds of chemical substances. The.general
public regards fish as the most important type of aquatic animal and is
more concerned with the effects of water pollution on fish than on other
aquatic organisms. Germane to the monitoring of toxic pollutants by fish
tissue analysis is the fact that fish are relatively easy to sample, gener-
ally occupy high trophic levels, accumulate many pollutants to much higher
levels than those concentrations found in ambient water, have a relatively
long life span and thus represent long-term conditions, and represent a
direct route for human uptake of these pollutants via ingestion. The pre-
sence and concentration of toxic pollutants in fish tissue thus indicate
the occurrence of these pollutants in the environment, the pathways through
which they travel in aquatic ecosystems, and the hazards to humans through
dietary exposure.
There are several problems in using the concentration of pollutants
in fish tissue to indicate the extent of the toxic pollutant problem. Dif-
ferences in fish size, age, lipid content, migratory patterns, position in
the food web, and uptake and clearing rates make it difficult to analyze
pollutant exposure. Even within a species, temporal factors (season/lipld
content, age/feeding habits, etc.) can make interpretation of data a spec-
ulative matter. Some methods can be adopted in developing a sampling pro-
gram to reduce the difficulty in interpreting data, such as establishing
target species for collection (thereby eliminating interspecific varia-
tions), sampling fish of similar age and size, and limiting sampling at
different sites to as short a period as possible to reduce seasonally-
related differences. Analytical techniques should also be used to Improve
the utility of the data. These include analysis of lipid content, quality
control (such as analysis of duplicate samples), and compositing tissues
from five or more individual fish. The recommended method of compositing
fish tissue is to grind each fish Individually and composite the homo-
genates so that analysis of individual concentrations can be performed, if
required (if you homogenate several fish together, rather than compositing
the homogenates, it is impossible to analyze individual fish when unusual
results warrant further investigation). Also, individual homogenates may
5-1
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b« desirable to establish the variance of tissue concentrations for various
pollutants. Analytical techniques are currently being developed by BPA's
Environmental Support Laboratory in Cincinnati, Ohio, and are provided in a
separate document.
Guidance on fish sampling is provided in the following sections and
is intended for use in rivers, streams, lakes, and estuaries throughout the
U.S. There is considerable overlap in the fish fauna and collection meth-
ods used in freshwater and estuarlne environments as well as in lentic
(standing water) and lotic (running water) environments; these systems are
/
not discreet, but rather different positions in a spectrum of aquatic en-
vironments. Because of the variety of conditions that will be encountered
specific policies on site selection, equipment, target species, number of
fish per sample, and collection schedule should be made as part of the ini-
tial planning of each sampling program. Regardless of how much planning is
done, the sampling crew will need to make judgements in the field. The in-
formation in this manual should be helpful in making these decisions, but
is no substitute for experience. We recommend that each sampling crew
include at least one experienced member for this reason.
One aspect of sampling fish that differs from sampling ambient water
and bed sediment is that a collection permit is often required. Before
performing any sampling, check with the appropriate fishery agency(iea) to
arrange for any necessary permits. If possible, coordinate joint sampling
efforts with other agencies to improve cost efficiency and data interpreta-
tion. The U.S. Fish and Wildlife Service (TVS) conducts periodic surveys
in some areas, and also collects some fish samples for tissue analysis, so
PWS is usually a good starting point for coordinating efforts.
5.1 Site Selection
The process of site selection should be given careful consid-
eration to ensure maximum benefits for the sample program. Many factors
play an important role in site selection and should be considered in an at-
tempt to maximize the success of the effort. Important factors that should
be considered include:
1. Purpose of sampling program.
5-2
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2. Proximity of sites for sampling water and bed sediment for
priority pollutants.
3* Previous priority pollutant sampling/analysis for fish
tissue at the site.
4. Availability of data on fish community structure*
5. Bottom conditions.
6. Type of equipment available.
7. Accessibility of site.
Each of these factors is discussed in greater detail in the following
paragraphs.
The specific purpose of the sampling program has an important
role in the selection of suitable sampling areas. In rivers and streams,
for example, a program designed to identify sources of pollution may re-
quire the selection of sites immediately up- and downstream of suspected
sources. In lakes and estuaries, sites may be selected in open water areas
if the program's goal is to provide an overall evaluation of pollutant
levels, or near river mouths or outfalls if the goal is to identify the
sources of pollutant input. The type of problem being studied will dictate
the site, the conditions at the site will affect target species selection,
and the site and species will affect equipment selection.
There are several advantages to locating the fish sampling sites
near sites selected for priority pollutant sampling of water and sediments*
The most important benefit of such a sampling design is the possibility of
developing at least a simple model of the dynamic distribution of pollu-
tants in that area. This consideration has greater weight if the target
species has a limited territory and spends most of its time in the immedi-
ate area. In cases such as this, correlations between pollutant levels ia
the different compartments (fish, sediment, and water) may be established.
Selecting sites in proximity to each other also allows a more efficient use
of time by providing the opportunity to combine sampling trips.
The availability of historical data on pollutant body burdens
should be cheeked before making any final decisions regarding sample sites*
5-3
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In most cases these data are nonexistent, but in some areas such sampling
has been conducted previously; in those few cases where this information is
available, consideration should be given to choosing sampling sites near
the areas sampled previously in order to construct a historical record of
pollution in the area.
Data on the ecology of fish communities should play an Important
role in the final selection of sample sites. Information on food chain re-
lationships, preferred feeding areas, spawning areas, and movement patterns
of target species is a valuable asset in site selection. Knowledge of this
sort is useful in locating populations of the target species. This type of
Information is available from fishery biologists familiar with the area.
Even though such an expert may not have previously sampled in the water
body in question, the. information provided regarding habitat preferences
may significantly reduce the time required to locate a suitable population
of the species. In areas having commercial fishing operations, experienced
commercial fisherman can also often provide valuable information.
Bottom condition is another factor closely related to the ecol-
ogy of the target population. Once the habitat preferences of a species
are known, the next step is to locate those preferred areas in the water
body being sampled. This factor includes such considerations as the pre-
sence of deeper areas (holes) preferred by some species, vegetation beds
which are often utilized as feeding areas, and dead trees and other shel-
ters which provide protective cover. Obviously, the bottom condition has
Important considerations related to equipment use as well. Obstructions
such as snags or oyster beds should be avoided when trawls or seines are
used. Depth contours and the presence of larger obstacles are readily
determined in coastal areas and larger navigable rivers by consulting
navigation charts. Other sources of this information include fisheries
biologists and commercial fishermen familiar with the candidate area, or
where possible use of depth finders.
The availability of equipment and personnel experienced in its
use is also an important consideration when selecting sample sites. If a
site which meets other criteria cannot be sampled with available equipment,
there is often the option of coordinating efforts with other fishery biol-
5-4
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ogiats or commercial fishermen who have the appropriate equipment and ex-
perience •
Another factor closely related to equipment availability is the
accessibility of the site. This is sometimes a problem with smaller
streams where it is impractical to use a boat. In such instances it is
desirable to locate sampling sites where there is good land access. The
same is true of land-locked lakes, particularly in mountainous areas. When
seining or shocking in areas which must be reached by land, factors to con-
sider are: necessary permission to cross private property, the presence of
brush and other obstacles which could make it difficult to carry a large
seine or electroshocklng equipment, and the depth and bottom gradient of
the sample site. If access is by water, consideration should be given to
the location of nearby boat ramps and marinas and the depth of water re-
quired to operate the boat safely.
The factors described above are among the important considera-
tions which should be evaluated in the selection of sampling sites. Other
elements may be important in some cases. For example, commercial fishing
and popular sport fishing areas may be good sampling sites since fish from
these sites represent vehicles for human exposure to water-borne pollu-
tants.
Obviously, the taxa of fish present will vary with the kinds of
sites selected. A list of "target species" has been compiled (Section 5.2)
representing common aquatic habitats throughout the country. If conditions
at several sites are to be compared, it is recommended that the same spe-
cies and age classes of fish be sampled from each site, since species and
age classes differ considerably in their propensity to accumulate and re-
tain pollutants. This, of course, requires that the sites selected be
similar as far as providing the habitat necessary for the desired species.
Like water and sediment, fish can be sampled upstream and down-
stream of point sources of pollutants to indicate the effects of these
sources on pollutant loading and aquatic health. Because of the mobility
of fish, however, such data should be interpreted carefully. The well
documented ability of fish to avoid noxious conditions can result in
situations where populations have avoided exposure to maximum concentra-
5-5
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tions of a pollutant they are able to detect* For the purpose of com-
paring conditions at nearby sampling stations, it is best to sample fish
with relatively narrow territorial confines. Several fish on the target
species list given in the next section exhibit well-defined territorial-
ly.
The best approach to developing a sound sampling program is to
consult with local fisheries biologists regarding the best candidate
species and areas in which those species can be located. Before sampling,
a reconnaissance should be performed to locate sampling sites and access
points. Once again, it is vital to arrange for any necessary collection
permits before implementing a sampling program.
5.2 Target Species
Because of the differences in habitat, niche, and pollutant up-
take among fish species, it is very difficult to compare the results from
different monitoring studies unless the same species are used. It is ob-
viously impossible to sample for the same species in every study; never-
theless, the number of "target" species should be fairly small to limit the
number of variables involved when interpreting data from different sites or
studies.
Several characteristics are important for selecting a target
species:
1. Wide ranging (e.g., broad distribution).
2. Iton-migratory.
3. Easy to identify.
4. Easy to capture.
5. Pollution tolerant.
6. Foodfish.
7. Abundance.
Wide range is vital since it allows sampling the same species in various
studies, thus enabling direct comparison among different sites. Non-mig-
ratory fish, especially those with a small individual territory, are also
5-6
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preferable, since they more accurately reflect ambient water quality at the
site where they were sampled. Species which are easy to capture and easy-
Co-identify are practical choices for sampling. Tolerance to pollution is
an important attribute because tolerant species can withstand pollutants
and accumulate them in their tissues; intolerant species will be absent in
such environments. Foodfish are preferred because they are large enough
and have a long enough life span to accumulate detectable concentrations of
pollutants, and they also represent a route of human exposure to pollu-
tants .
It is recommended that both predators and bottom feeders be
sampled. Predators and bottom feeders are not mutually exclusive designa-
tions but conveniently describe two ecological groupings of fish that are
apt to be exposed to priority pollutants in large concentrations. Predators
can indicate the presence of pollutants that are biomagnified (accumulated
in the food web). Bottom feeders (most of which prey on benthic Inverte-
brates) may come in contact with heavy pollutant concentrations because
many of the organics and metals partition strongly from water to sediment
and can then be accumulated by benthic and eplbenthic organisms.
In lacustrine and estuarine systems, planktlvores should also be
sampled, if possible. Plankton are essential to the aquatic food web in
these systems and play an important role in pollutant uptake.
It is recommended that five fish of the same species be collect-
ed per site for each of the trophic groupings that are to be sampled (i.e.,
predator, bottom feeder, planktivore). For instance, at a warm water
stream site, a sampling crev might collect five largemouth bass (pre-
dator) and five white suckers (bottom feeder). . Mixing species at the same
site is not recommended (i.e., three largemouth bass, two blueglll). Each
fish should be wrapped individually and then placed in a plastic bag con-
taining all fish of the same species from the same site, as described in
Section 5.6. The minimum sample mass (per species) is 300 grams; if five
fish do not weigh an aggregate 300 grams, more should be collected until
this minimum is reached.
Target species for freshwater and estuarine systems are dis-
cussed in Sections 5.2.1 and 5.2.2, respectively. Synopses of the range
5-7
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and habitat preferences of the target species are provided in Appendix B.
Often, it will not be possible to collect one of the recommended species.
In this case, collect congeneric species, if available. It is always ,a
good policy to collect the same species throughout a sampling project;
however, this is not always possible. The most important principle is to
be consistent.
5.2.1 Freshwater Target Species
Recommended target species for warm water, cold water, and the
Great Lakes and other lentic systems are listed in Table 5-1. Detailed
information on range and habitat preference for the target species and
similar species can be found in common field guides, such as McClane (1978a
and 1978b). The U.S Fish and Wildlife Service and state fisheries agencies
can usually provide information on types of fish present as well.
The fish that are listed in Table 5-1 satisfy most of the crite-
ria listed previously for selecting target species. This list is not in-
tended to be exclusive, but should serve as guidance for selecting fish to
sample.
Some minnows are widely distributed and easy to capture, but
identification of these fish is fairly difficult, especially in the field.
Because most minnows have a relatively short life-span and are fairly low
in the food chain, their capacity for accumulating pollutants may not be as
great as most of the species listed in Table 5-1. Other than the carp,
squawfish, and stoneroller, which are long-lived, large cyprlnids, minnows
are absent from the target species list. They can, however, be useful for
screening purposes to determine the presence of pollutants in ambient
water, and, in some headwater streams, minnows may be the only alternative
to stocked trout.
Although trout are of a suitable size and may be the only large
species in many cold-water streams, caution should be exercised in using
trout as a representative species. Many trout are stocked on a "put and
take" basis, so they may not represent long-term exposure conditions for
the pollutants in the stream in which they are found. The agency respons-
ible for stocking trout should be able to provide information as to when
5-8
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Table 5-1. Target Species for Warm Water, Cold Water, and the
Great Lakes and other Cold Water Lentlc Systems
WARM WATER
Predators:
**Largenouth bass (Mieropterus salaoides)
**Smallmouth bass (M. dolomieul)
**Yellow perch (Perea flavescenk)
*Bluegill (Lepomls aacrochlrua)
^Channel catfish (Ictalurus punctatus)
*Chaln pickerel (Esox nlger)
COLO WATER
Predators:
Bottom Feeders:
**White sucker (Catostomus
commersoni)
**Carp (Cyprlnus earpio)
*Stoneroller (Campostoma
anomalum)
*Spotted sucker (Mlnytrema
melanopa)
*Sllver redhorse (Moxostoma
anlaurum)
^Freshwater drum (Aplodinotug.
grunnlens)
Bottom Feeders:
**Rainbow trout (Salno galrdnerl)
**Brown trout (S. trutta)
**Brook trout (Salvellnus fontinalls)
*SquawfIsh (Ptyehocheilus oregonensls)
*Round whlteflsh (Prosoplun cylindraceun)
*Moutain whltefish (P. villiamsoni)
**White sucker (Catostomus
eommeraoni)
**Largescale sucker (C.
aacrocheilua)
*Carp (Cyprinus carplo)
*Sllver radhorse (Moxostoma
anisunaa)
*Northera(shorthead)redhorse
(M. taacrolepldotum)
•Freshwater drum (Aplodinotua
grunnlena)
Predators:
GREAT LAKES & OTHER COLD WATER LZNTIC SYSTEMS
Bottom Feeders:
**Lake trout (Salvellnus namayeush)
**Yellow perch (Perca flavescens)
*Walleye (Stlzostedian vitreum)
*Coho salmon (Oncoryhnchus kisuteh)
*Sauger (S_. eanadense)
^Northern pike (Esox luclus)
*Round whlteflsh (Prosoplum eyllndraceum)
*Lake whltefish (Coregonus elupeaforatis)
*Ralnbov smelt (Osmerus mordax)
**Whlte sucker (Catostomus
commersoni)
**Carp (Cyprlnus earpio)
*Sllver redhorse (Moxostoma
anlsurum)
*Northern(shorthead)redhorse
(M_. macrolepldo turn)
*Fres¥water drum (Aplodinotus
grunnlens)
**Preferred target species
*Good target species
5-9
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a particular stream was last stocked. The best policy is to avoid using
stocked fish at all, whether they are trout or other species.
As previously mentioned, planktivores should- be included when
lacustrine systems are sampled if the budget will allow it. Clupeids
(Herring family) are generally good target planktivores since they have a
high lipid content and thus accumulate llpophilic compounds.
A taxonomic key and field guide should be a part of the field
equipment. Identification must be done in the field, as it is much easier
to do with live or recently dead specimens than those that are frozen.
5.2.2 Estuarine Species Selection
Ideally, estuarine fish samples should comprise predators, bot-
tom feeders, and planktivores as suggested for lacustrine areas. Plankti-
vores play a very important role as forage fish in the highly productive
waters of most estuaries, and priority pollutant data for planktivores may
furnish valuable insights regarding the pathways of pollutants in these
waters* Predators and bottom feeders also tend to accumulate high concen-
trations of pollutants, as previously discussed.
Some special problems are encountered in the selection of target
species for estuarine areas* One major problem is the lack of species
which are represented on both the Atlantic and Pacific coasts. Another
difficulty encountered in estuarine areas is the tendency of many species
to move seasonally. This results in uncertainty as to the source of any
pollutants detected in many estuarine species. A related factor is the
role of estuaries as spawning and nursery areas for many species. In many
cases, sexually mature adults only enter the estuary to spawn. As mention-
ed in Section 5.4, spawning populations should not normally be sampled. If
only juvenile stages of a species occur in the estuary, another species
should be selected since juveniles may not have had sufficient time to ac-
cumulate pollutants. In some cases, these seasonal movements may vary from
estuary to estuary for a given species. The best approach to selecting
estuarine species is to consult with an authority familiar with the estuary
to be sampled.
5-10
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AM described in Section 4, an alternative approach for detecting
pollutants in estuarine biota is to sample shellfish rather than finfish.
Shellfish, such as clams and oysters, are sessile and may represent local
pollutant levels more accurately than the highly mobile finfish.
Table 5*2 lists some recommended estuarine fish for the Atlantic
and Pacific coasts. Many of the Atlantic species are found along the Gulf
Coast also. When possible, this list recommends closely related species
for the Atlantic and Pacific coasts. Other local species may be more suit-
able in some cases. In any event, careful consideration should be given to
choosing species which will represent local conditions as much as possi-
ble.
Table 5-2. Target Species In Estuaries
West Coast
Starry flounder (Platichthys stellatus)
Striped mullet (Mugil cephalua)
Rainbow smelt (Osmerus mordaae)
Whitebait smelt (Allosmerus elongatus)
Pacific staghorn aculpin (Laptocottus armadus)
Atlantic and Gulf Coast
Winter flounder (Pseudopleuroneetes americanua)
Striped mullet (Mugil cephalus)
Spot (Leioetomus zanthurus)
Croaker (Mierepogon undulatus)
Rainbow smelt (Osmerus mordaz)
Yellow perch (Perea flaveseens)
White perch (Morone americana)
5.3 Sampling Equipment and Use
A number of collection techniques have been developed to sample
fish representative of different habitats, sizes, and behaviors. In re-
sponse to the variations in physical conditions and target organisms of
interest, fisheries biologists have had to devise methods which are, for
the most part, intrinsically selective for certain species and sizes of
fish. Although this selectivity can be a hindrance in an investigation of
community structure, it is not a problem where fish tissue analysis is con-
cerned. Results from different samples can only be compared if uncontrol-
led factors such as differences in taza and size are minimized.
5-11
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Collection methods can be divided into two categories, active and
passive. Individual methods are discussed under these categories in
Sections 5.3.1 and 5.3.2.
5.3.1 Active Collection
Active collection methods utilize electroshock units, seines,
trawls, angling equipment, and chemical poisons. Although active collec-
tion requires a greater amount of fishing effort, it is usually more ef-
ficient than passive collection for covering a large number of sites and
catching the relatively small quantities needed from each site for tissue
analysis. Active collection methods are particularly useful in shallow
waters such as streams, along lake shorelines, and along the shallow coast-
al areas of estuaries. When sampling must be conducted in deep water, how-
ever, active collection methods have distinct disadvantages because, as
previously mentioned, they are more Intensive requiring large numbers of
personnel and expensive equipment. This problem, however, may be overcome
when sampling efforts to collect fish samples for priority pollutant an-
alysis are coordinated with other scientific or commercial collecting ef-
forts. In such cases, a subsample can be taken from the entire catch to
obtain a sample for priority pollutant analysis.
The following discussions describe each of the most common active
collection methods and equipment, the procedures for proper use, and the
advantages and disadvantages of each as tools for the collection of fish
samples for priority pollutant analysis.
5.3.1.1 Zleetroflahlng
Electrofishing is the most efficient and least selective sampling
method available with the exception of radical methods such as poisoning or
draining. An AC, DC, or pulsed DC electrical current is applied to the
water, which stuns the fish. Pulsed DC is commonly used because it is
apparently most efficient in terms of power and because it often attracts
fish to the anode. The attraction is caused because the swimming muscles
of the fish are stimulated, causing orientation in the direction of the
electrode. Although fish are usually just stunned, higher voltages can be
lethal. Most stunned fish usually float to the top, where they can easily
be collected, but some sink when stunned (e.g., catfish) and thus are not
readily sampled in this manner.
5-12
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Electrofishing is not a viable alternative in brackish, salt, or
extremely soft (hardness less than 25 mg/1, as C&CQ$) waters (Bennett,
1970; Battalia, 1975). In soft waters, alternating current or pulsed DC
may be more effective (Weber, 1973). In most other fresh waters, DC or
pulsed DC is usually employed. Areas where these considerations may be
very important are the Southeast where the conductivity is frequently too
low and in the West where it is often too high*
Two types of electroshocking units are in general use: the boat
mount shocker and the backpack shocker. The boat mount shocker uses a gen-
erator run by an air cooled gasoline engine. The approximate weight of the
motor/generator assembly is 70 Ibs, and it has a utility of about 500 watts
and 110 volts. The unit is usually mounted in the middle of the boat with
the probes extending from bars on the front of the boat. The depth of the
electric field can be lowered or raised by adjustment of these probes.
Three investigators are required to operate this unit; one operator is
responsible for the generator and boat while the other two are responsible
for keeping the electrode probes in position and collecting the stunned
fish. Boat mount shockers can be used in shallow rivers, ponds, lakes, and
impoundments, but they are usually not effective in water deeper than 5
meters.
Smaller backpack shockers are available for small wadeable water-
ways. At least two, and preferably three or four, investigators are
required to carry the backpack, deploy the probes, collect the fish, and
carry the sample containers and cooler.
Caution should always be employed when electrofishing because the
currents used can not only stun fish, but also kill humans* For this rea-
son, safety precautions such as rubber linesman's gloves, waders, and a
readily available cut-off switch should always be included as mandatory
equipment when backpack shocking; a "deadman switch" should always be
installed in all shockers. Metal handled collection nets should not be
used. It is important to emphasize that the sampler responsible for the
generator must be capable of shutting off the electricity instantly in case
of mishap.
5-13
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All personnel using eleetrofishlog techniques should receive
basic training from experienced operators and have electrofishing safety
manuals. For sampling with an eleetroshock unit, it is a good safety mea-
sure to have at least one experienced operator per sampling crew. All mem-
bers, of the crew should know basic C?R (cardio-pulmonary resuscitation)
techniques. The need for safety precautions when working with eleetroshock
units cannot be too strongly emphasized.
Although there is some added element of danger Inherent in elec-
trofishing, with caution, problems can be avoided. Electrofishing is, for
many applications, probably the best method available because:
1. There is minimal damage to the fish, which concomitantly
reduces the danger of contamination.
2. It is very efficient in terms of catch per unit time.
3. It is one of the least selective techniques, so that it
provides flexibility in "target" species.
4. It can be adapted for use under a variety of conditions.
5.3.1.2 Seines
A seine consists of a wall-like collecting net held upright in
the water by floatlines at the top and lead weights on the bottom. Poles
are attached at each end of the net to provide a grip and help keep the net
stretched vertically. The nets vary in mesh size as well as length and
usually are operated by two or more individuals. Select a mesh size that
is small enough to retain the smallest fish to be sampled; very small mesh
sizes are disadvantageous because of their drag.
Seines can be used in relatively shallow waters in which
organisms can be captured by surrounding an area and pulling the net ashore
so as to enclose the specimens. It is extremely important to keep the lead
line on the bottom because most fish will attempt to go under the net.
Often larger specimens of fish such as bass and trout will attempt to jump
the net. Seines are not effective in lakes or streams with irregular
bottoms and snags.
5-14
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Good technique is required for efficient use of a seine. One
sampler temains close to shore and plays out the seine so that it doesn't
get tangled. The other pulls the seine away from the shore, keeping an eye
•
out for snags, and then turns parallel to the shore. When the seine is
completely deployed, the shoreline sampler starts moving in the same direc-
tion as the leading sampler. The seine should then be in a long "J" shape.
The leading sampler turns quickly into shore, and both ends of the seine
are brought close together. The poles are dropped, and the seine is pulled
in, one hand on the float line and one hand on the lead line, taking care
that these stay on the surface and bottom, respectively. If the seine is
snagged, give it slack, and while one sampler holds the ends, the other
wades out and frees the snag. In flowing waters, the seine should always
be let out as the samplers move upstream.
Seines are somewhat less selective than other net collection
forms in that they will take any specimen which cannot pass through the
mesh. Since they are relatively cheap and easy to operate, they are widely
used.
5.3.1.3 Trawls
Trawls are specialized seines most commonly used in larger open
bodies of water. They vary in types and sizes but all are pulled with
boats at speeds sufficient to overtake the fish. The two types most com-
monly used in fresh and estuarine waters are the otter trawl and the beam
trawl, both of which are used in collecting fish near or on the bottom.
The beam trawl incorporates a log like beam responsible for holding the net
open and scaring the fish up from the bottom into the net (See Figure 5-1).
This travl has a very rigid opening and is quite difficult to operate from
small to medium sized boats. In recent years beam trawls have been more or
less replaced in fish collection methods by the more efficient otter trawl.
The otter trawl utilizes otter boards which are sections of white oak bound
with heavy iron runners to protect the wood. These boards are placed at
either side of the mouth of the net and serve to keep the net open (see
Figure 5-2).
Despite this modification in design, crawls cannot be used in
water bodies with irregular bottoms. Operation of a crawl often requires
5-15
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Figure 5-1. Beam Trawl (from Weber, 1973).
Figure 5-2. Otter Trawl (from Weber, 1973)
5-L6
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the use of a boat equipped with heavy winches and motors as well as
experienced personnel. Furthermore, since the number of fish collected by
trawling far exceeds the number required for fish tissue analysis at any
given site, this method of sampling is probably too intensive (and expen-
sive) for most priority pollutant investigations unless small sampling
trawls are readily available*
5.3.1.4 Angling
Angling is one of the most selective forms of fish collection.
This fora makes use of the hook and line method of fishing. Many varia-
tions of angling exist, the moat productive being set lines or long lines
used primarily for larger, non-schooling species of fish. These are essen-
tially one long mainline anchored or stretched between floats with smaller
drop lines with baited hooks attached at intervals. (Trot lines, which are
single baited lines attached to branches or other fixed points, may be more
useful in rivers.) These lines are normally checked daily by two Investi-
gators working from a boat. This method may be especially productive for
particular species, such as catfish.
Another fora of angling commonly utilized is the use of rod and
reel, ranging from large tuna poles to the sport model rods and reels. Al-
though angling Is not a dependable means of collection and is generally not
as efficient as other methods, in some cases it may be easier to catch fish
with a hook and line than to use some of the more elaborate techniques.
Therefore, angling should be considered as an acceptable procedure, par-
ticularly for deeper bodies of water. Bank fishing, drift fishing, and
trolling are all effective for catching fish with rod and reel. Deep
trolling is a good way to catch fish such as lake trout which inhabit deep
water.
5.3.1.5 Poisoning
Various poisons have been used to sample entire fish communities
for purposes of determining community structure and population dynamics.
Although the use of these poisons is justifiable for such studies, they
should be avoided when sampling fish for analysis of toxic pollutants
because they may induce physiological changes which could alter the concen-
5-L7
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tration of pollutants in the tissue. It is interesting to note that sev-
eral of the chemicals historically used for fish sampling are now on the
EPA priority pollutant list (e.g., cyanide, copper, toxaphene).
5.3.2 Passive Collection
Passive collection methods include gill nets, Fyke nets, trammel
nets, hoop nets, pound nets, D-traps, and purchasing fish from commercial
fishermen. These forms of fish collection generally require less fishing
effort than the active forms but are usually less desirable for shallow
water collecting because of the ability of many species to evade entangle-
ment and entrapment devices. These methods normally yield a much greater
catch than necessary from a particular site and are time-consuming; in deep
waters, however, passive collection techniques are generally more efficient
than active methods.
5.3.2.1 Gill Nets
The gill net consists of a loosely hung single wall of diamond
shaped mesh with a float line on top and a lead line on the bottom. Fish
are captured when they swim part way through the mesh and are entangled by
the net behind the gills (see Figure 5-3). The degree of selectivity in
Figure 5.3 Gill Net (from Battelle, 1975)
5-18
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this net is controlled by the mesh size chosen (Weber, 1973). The recom-
mended range of use for the gill net includes lakes, reservoirs, estuaries,
or rivers where fish movement can be expected. They are used extensively
in deeper water where they can be set at any desired depth by adjusting the
length of the anchor and buoy lines. These nets are set from a boat and
require at least two individuals for proper setting. Gill nets have proved
effective in collecting pelagic fish, but are also characterized by severe
tangling problems when used for species with large or barbed spines such as
the channel catfish.
The gill net is commonly set for a period of 24 hours, after
which the sampling crew returns to remove specimens that have become en-
tangled during the sampling period. One Important consideration is that
the gill net will eventually kill entangled fish. Specimens that have been
killed and allowed to remain entangled for extended periods may undergo de-
gradation or other physiological changes that could possibly alter the con-
centration or character of toxic pollutants contained in the fish tissue.
Therefore, only those fish that are still living or have recently died
should be sampled.
5*3.2.2 Trammel Nets
Trammel nets consist of a light, small-mesh net hung between two
walls of large mesh webbing. The fish are captured when they hit the light
gill net, pulling a pocket of this netting through the mesh of the larger
net (see Figure 5-4). These nets are equipped with a float and weight
system identical to the gill net and are commonly fished across waterways
or by surrounding visible schools of fish. Trammel nets are most commonly
used for commercial fish which can be scared into the nets, such as carp
and catfish (Bennett, 1970). Two investigators are required to set these
nets from a boat, but additional personnel are recommended to aid in
removing severely tangled fish from the netting. As with gill nets, only
living or recently dead specimens should be sampled.
5.3.2.3 Hoop, Fyke, and Pound Nets
The hoop net is essentially a shallow water gear owing -to the
difficulty of setting it effectively in deep water. The hoop net is used
5-19
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extensively in river sampling and is best adapted for use in a fair current
or where fish move in predictable directions.
Figure 5-4. Trammel Net (from Weber, 1973).
The typical hoop net (Figure 5-5) is a long cone shaped bag
mounted on one or more hoops* The hoops serve a double purpose; they keep
the net from collapsing, and they form the attachment for the series of
internal funnels (throats) which prevent the fish from escaping readily.
The hoops are most frequently constructed of metal or wood, the size of
which depends on the depth to be sampled and the particular species to be
captured. Most commonly, the trap consists of 5 hoops of decreasing size,
the largest hoop being at the front of the net. The hoops are equally
spaced and overall length may vary from 3 to 15 meters. Two funnel shaped
throats lead inside the net. The first throat is attached peripherally to
the front hoop and posteriorly to the third hoop. The second throat, which
leads to the cod or pot end, is similarly attached to the third and fifth
5-20
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hoops. The mesh at Che pot or cod end of the net is equipped with a
drawstriag that may be released for easy removal of the catch. Nets of
this type are also available with square frames, which reduces the tendency
to roll and twist.
Figure 5-5. Hoop Nets (from Battalia, 1975).
Construction of the hoop net facilitates handling and transport.
The net will collapse upon itself, and thus can easily be stored or carried
on the deck of a small boat. The net is set parallel to, and usually fac-
ing, the current. This arrangement reduces current resistance and also
places the throat end in the path of downstream moving fish. The net can
be secured and kept taut by anchors or posts secured in the substrate. The
net may or may not be baited depending upon the needs or selectivity of the
investigation.
The Fyke net (Figure 5-6) is essentially a hoop net with one or
more wings attached to the first frame in order to increase its efficiency*
5-21
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Fylce nets are also used in shallow waters, particularly where little or no
current exists and where fish movement is more random such as in lakes,
impoundments, or estuaries.
-•t-\.#Y 4k*1
•?sHnri 5S*!
Figure 5-6. Fyke Net (from Weber, 1973).
The wings and leader of a Fyke net consist of net meshing of the
same size as the hoop. The wings are set obliquely on either side of the
mouth of the bag and are generally 1.5 times as long as the length of the
bag. The leader extends directly out perpendicular to the mouth of the
bag, and is generally 5 times the total length. The wings and leader of a
Fyke net will have a depth equal to the diameter of the first hoop. The
top of the net is supported by floats and the bottom by a lead line. The
entire structure is kept taut by posts and anchors secured in the sub-
strate.
5-22
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By nature, the Fyke net la a rather semi-permanent structure.
The net can easily be set up by two people and thereafter serviced by one
person. The nature of the wings and leader Increase the capture efficiency
of the hoop net. As fish move and strike the wings or leader, they are de-
flected toward the mouth of the net.
Found nets are a particularly effective method for large-scale
capture of migratory species that tend to follow a shoreline. There are
scores of designs and variations, but the operating principles are the same
for all. Fish moving along the shoreline encounter a lead of wire, brush,
or net. They follow this lead in order to attempt to pass around it, but
are led into one or more enclosures from which escape is extremely dif-
ficult.
The typical pound net used on the Great Lakes or Atlantic sea-
board uses wire mesh or net that is hung from poles driven In the bottom.
These poles are arranged in a straight line leading away from the shoreline
towards the Impoundment area in deeper water* There, piles are driven into
the bottom in depths of up to 25 meters or more from which nets are also
hung. This area, often called the heart, is opened from the lead, and fish
entering are further funneled to the trap area. The designs of the trap
area are also extremely variable, but in essence, they all facilitate the
hauling of the catch to the surface for removal to the servicing boats.
Pound nets are typically used for commercial purposes and not
routine sampling programs. Many persons are Involved in the operation of a
pound net, and it is therefore not a feasible operation for priority pollu-
tant sampling programs. It is, however, quite possible that members of the
sampling team could accompany commercial fishermen who are operating pound
nets and buy fish for sampling purposes. If this alternative is used, the
members of the sampling team should take responsibility for removing the
desired fish, weighing, and measuring them.
5«3«2 «4__ D-Traps
The D-trap is a particularly effective method of capturing slow
moving fish (e.g., sunfish, perch, catfish) and crustaceans (e.g., crabs,
lobsters) that move about on, or just above, the river, lake, or estuary
5-23
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bottom. The D-trap is usually small enough so that several can be piled on
the deck of a small boat and light enough to be operated by one person.
The D-trap is so named for its construction. Wire, wood slats
and cotton or synthetic mesh are supported by three attached D-shaped
frames. The trap consists of two compartments, the "chamber" and the "par-
lor." Fish enter the chamber through a suspended funnel shaped throat and
then pass through a second funnel to the parlor. The fish are removed from
the parlor through a draw-string pocket or a door if the trap is con-
structed of wire or wood. Mesh and trap size depend on the needs of the
investigation.
The trap can be set from a boat or the shoreline of a river or
lake in either shallow or deep water. The trap is weighted with a few
bricks, steel rods, or stones, and a buoy line is attached to the lower
corner of the chamber end. The trap may or may not be baited. A typical
set is 24 hours; however, this also varies with the study.
Many variations of the D-trap have evolved with time (Figure
5-7). The most commonly used type are those traps or pots used in the
lobster fishery. Frequently, rectangular traps are used. These are par-
ticularly popular because they are easily made and stored. All pots or
D-traps are fished in the same manner, and they differ in their efficiency
and selectivity. Efforts should be made to find which shape or size best
fits the needs of the particular investigation about to be undertaken.
D-trapa can be useful in sampling fish for priority pollutant
analysis, especially when sampling fast, deep waters where other methods
are difficult to use. They are usually fairly effective for sampling
bottom feeders and less effective for the more visually oriented predators.
They are less effective for all species when water is clear than when it is
turbid. Although it is generally true that the passive collection methods
work best during periods of extensive fish movement, this seems par-
ticularly true for D-traps. Because of the highly variable catch ef-
ficiency of D-traps, they are not a good choice for a primary sampling
technique, but are valuable as a back-up for other methods.
5-24
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HALF-ROUND LOBSTER POT
CRA8 POT USED IN
CHESAPEAKE BAY
HALF-SOUND EEL POT
Figure 5-7. Modifications of the D-Trap (from Battelle, 1975)
5-25
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5.3.2.5 Purchasing Specimens
Purchasing fish from commercial fishermen is a common and ac-
cepted method of obtaining specimens. When collecting for priority pol-
lutant analysis, it is imperative that samples be preserved immediately and
with the least amount of handling. To ensure that proper preparation tech-
niques have been observed, a member (or members) of the sampling team
should accompany the fishermen during the operation. In this way, trained
personnel can remove the fish from the nets then weigh, measure, and pack-
age them with minimal chance of contamination. This is a good method of
obtaining specimens of commercially important species in areas such as the
Great Lakes and coastal estuarine areas.
Another good possibility in some cases is to work in cooperation
with the state fishery agency if they are planning a sampling program.
Members of the sampling team can accompany the fishery biologists and
remove the desired specimens from the nets or traps.
5.3.3 Summary
In summary, sampling methodologies may be divided into active
methods, in which the sampling team can generally catch small-quantities of
fish in a short period of time, and passive methods, which are usually
more selective means of capturing large numbers of a particular species.
Active methods of collection are usually less selective and allow the sam-
pling crew to exercise some control over the sample size. Electroshocklng
is probably the best of the active means of acquiring samples for tissue
analysis, but the other active collection methods described may be used,
with the exception of poisoning which, until proven otherwise, must be
assumed to induce physiological changes that may alter the concentrations
of pollutants in the tissues. In deep water or strong currents, passive
collection methods may be more efficient in collecting samples. In all
cases, the method used depends primarily on the water conditions of the
sampling area, the judgement of the sampling team, and the equipment avail-
able. Table 5-3 summarizes the use and advantages and disadvantages of each
type of sampling equipment or method previously discussed.
Regardless of the sampling technique employed, five or more fish
weighing a total of at least 300 grams should be collected from each sam-
5-26
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TobU 5-J
ry of Fish SIMP I Ing Equipment
Device
Use
Advantages
Disadvantages
Active Methods
Electroflshlng
Shallow rivers, lakes, and stream.
Most efficient non-selective Method.
Nlnlswl damage to fish. Adaptable
to a number of sailing conditions
(e.g., boat, wading, shorelines, etc.l
Particularly useful at sites where other
active Methods cannot be used (e.g.,
around snags and Irregular bottom
contours!.
Nan-selectivity - stuns or Mils
Most fish. Cannot be used In
brackish, salt, or extremely soft
water. Requires extensive operator
training. CANCEROUS when not used
properly.
Seines
Shallow rivers, lakes, and stres
Shoreline areas of estuaries.
Relatively Inexpensive and easily
operated. Hesh size selection
available for target species.
Cannot be used In deep water or
over substrates with an Irregular
contour. Not completely efficient
•s fish can get over, around, and
under during seining operation.
Trawls
Used from boats In deep open bodies
of water.
Effective In deep waters not accessible
by other Methods. Allows collection of
number of samples.
Requires boat and personnel with
operator training.
Angling
Generally species selective Involving
use of hook and line.
Host selective netbod. Does not
require use of large number of per-
sonnel or expensive equipment.
Inefficient and not dependable.
Passive Methods
GUI Nats
•Lakes, rivers, and estuaries. Where
fish movement can be expected or
anticipated.
Effective for collecting pelagic fish
specie*. Nat particularly difficult
to operate. Requires less fishing
effort then active Methods. Select-
ivity can be controlled by varying
Mesh site.
Not effective for bottom-dwelling
fish or populations that do not
exhibit Movement patterns. Nets
prone to tangling or damage by
large and sharp splned fish. GUI
nets will kill captured speclMens,
which, when left for extended
periods. May experience physio-
logical changes In the tissues.
-------
TabU 9-3 (Cont.l
Summary of Fish Sampling Equipment
Do vie*
Use
Advantages
Disadvantages
Passive HttthodS
Nats
u»
I
i^>
en
Same as gill nets. Frequently
used where fish My be scared
Into the net.
Slightly mora efficient than • straight
gill net.
Same as for gill nets. Tangling
problttu aay ba anr« savar*.
Hathod of scaring fish lato aat
raqulras aora aarsonnal or possibly
boats ja daap jatar araas.
Hoop, Fyka
Pound Met*
Shallow rlvars, lakas, and astuarlas
Mhara curraats ara prasant or Mhaa
•ovaaants of fish ara pradlctabla.
frequently usad la coaaarclal opar-
atlon.
U»at»andad oparatlon. Vary afflclant
In ragard to long tar* ratura and
axpandad affort. Particularly useful
la araas Mhara active Methods ara
Impractical.
laafflclaat for short-tans.
Difficult to sat up and anlntaln.
O-Traps
Used for long-tans capture of clow
aovlng fish, particularly bottoa
speclas. Can ba used In all
environments.
Easy to operate and sat. Unattended
operation. Particularly useful for
capturing bottom dwelling organisms
In daap waters or other types of In-
accessible areas. Relatively Inexpen-
slve — of tan can ba hand aade.
Efficiency Is highly variable. Nat
eftactive for pelagic fish or fish
that are visually oriented.
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pling site as previously mentioned. Where several small fish are combined
to make up the sample, these must all be of the same species.
5.4 When to Collect
A seasonal monitoring program provides the most complete infor-
mation about levels of pollutants in fish tissue and has the advantage of
building a good data base upon which future monitoring programs can rely.
Financial constraints usually limit the amount of sampling which can be
performed, and therefore consideration must be given to choosing the best
time of year to sample. The state fishery agency (or comparable state
agency) may be able to provide suggestions as to when and where to collect*
The necessary collecting permits will have to be obtained from these agen-
cies in any event.
The spawning season should normally be avoided since fish are
usually stressed during spawning and samples may not be representative of
the normal population. Changes in feeding habits, fat content, respiration
rate, etc., occur during this period which may influence pollutant uptake
and clearing. Information on the spawning season of a species may be found
in standard references such as Carlander (1969) and should be checked
before a sampling program is started. The additional stress of collecting
fish during their spawning season may also have adverse consequences on
some species such as trout or bass, by inhibiting their spawning behavior
or damaging the spawning grounds* On the other hand, populations of some
species, such as bluegllls, may actually benefit if their population size
is reduced during spawning. For most species, spawning occurs during the
spring and this season should be avoided unless the target species is known
to spawn during some other season.
Fall sampling is probably the best choice for avoiding the
v
spawning season. Water levels are typically lower In many areas during the
fall which may make active collection easier. Fat content represents an
Important reservoir for many pollutants in fish and is often higher in many
species during the fall. Another advantage of collecting in the fall is
that a new year-class has usually reached maturity, resulting in greater
abundance of the species. Again caution should be exercised to ensure that
5-29
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sampling does not occur during the spawning season since some salmonids and
other fish spawn during the fall.
Collection hours should be those which maximize sampling.success
and reduce injury to extra fish collected and released. The best time per-
iod to collect is during the night, but this may be inconvenient for a num-
ber of reasons. The work schedule of the sampling team must be adjusted,
and an additional person is usually required to operate some type of light
while the rest of the team samples. If sampling is to be done at night,
state and local regulations regarding the use of a light to fish must be
checked and any necessary permission should be secured from the appropri-
ate authority.
Although the catch is not usually as great as at night, the most
practical time to sample is usually early morning or evening. Fish are
frequently easy to catch at these times since they often come into the
shallows to feed. In addition, the stress to the population resulting from
sampling activities is less than at mid-day when water temperature is
highest. Every effort should be exercised by the sampling team to ensure
that fish are not needlessly injured or killed. Injured fish are much more
susceptible to fungal infection which is brought on by damage to the slime
layer.
When samplers collect in estuarine areas, tidal stage may in-
fluence the availability of many species. Local fishery experts can pro-
vide helpful advice as to which tidal stage is most appropriate for the
target species. Another consideration in this case is that different areas
will be inundated by the tides and those shorelines having too many snags
to seine at high tide may be relatively clear at low tide.
5.5 Container Selection and Cleaning
The selection of containers or packaging material for the trans-
portation of fish samples to the laboratory for priority pollutant analysis
is dependent upon a number of factors including the risk of sample contam-
ination by the container and the mode of sample transport. The most Im-
portant consideration is to prevent contamination of the sample. Because
polyethylene or polypropylene packaging materials can introduce contami-
nants (such as phthalate esters) to the sample, they, as well as all other
5-30
-------
plastic containing materials, should not be used for packaging fish samples
for priority pollutant analysis. Clean, sterile Teflon® bags, such as the
air sample hags manufactured by Pollution Measurement Corporation (Chicago,
111.) are the ideal packaging material. The principal disadvantage of
Teflon* bags is the high cost. A 4 x 9 inch Teflon* bag typically retails
for about $13.00.
Alternatively, aluminum foil or glass containers are the most
practical materials. Aluminum foil is preferred over glass, however,
because of the risk of container breakage in transport when glass con-
tainers are used. Whichever packaging material is used, it must be care'
fully cleaned to ensure that no contaminants are introduced to the packaged
sample. Aluminum foil should be previously cleaned by rinsing with acetone
and again with pesticide grade hexane and allowed to dry in a contaminant
free area. After the samples have been wrapped with foil, they should be
sealed in polypropylene bags to retain moisture. When glass containers are
used, they should be washed with a non-phosphate laboratory detergent,
i
rinsed with tap water and distilled water, and finally rinsed with acetone
and pesticide grade hexane and .allowed to air dry in a contaminant free
area. Also, when glass containers are used, the cap should be lined with
Teflon* sheeting to further prevent sample contamination by the cap
material (EPA, 1981).
Finally, although it is impractical to clean all sampling equip-
ment such as nets and traps, common sense does apply. Sampling equipment
that has been obviously soiled by oils, grease or household solvents should
not be used. Nets, seines, and other gear should not be treated with pre-
servatives (for this reason, nylon seines are preferred over cotton seines
because the nylon seines are more decay-resistant). All equipment should
be carefully stored away from chemical solvents and household items such **
paints, cleansers, and disinfectants. Finally, all utensils or equipment
that will be directly used in handling fish or tissue samples, such as
forceps and scalpels, should be rinsed with acetone and pesticide grade
hexane and stored in similarly cleaned aluminum foil.
5-31
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5.6 Sample Handling, Preservation, and Shipment
Fish selected for laboratory analysis should be carefully han-
dled to prevent contamination by the sampler's hands or by field equipment.
•
The use of cleaned forceps to handle selected specimens will greatly reduce
this risk. Upon removal from the collection device or water, larger fish
should be properly stunned by a sharp blow to the base of the slcull with a
stick or metal rod. This rod should be used solely for the purpose of
stunning fish, and care should be taken to keep it reasonably clean to
prevent contamination of the samples. Normally, whole fish will be ana-
lyzed, tf previous studies have indicated the presence of a pollutant and
the reason for sampling is to determine potential human exposure, edible
portions should be taken. Filleting, scaling, etc. should be done in the
field.
Weight and total length should be determined for all fish col-
lected for laboratory analysis (use metric units). It is recommended that
weight and length be determined In the field.
Bach sample container (e.g., bag or foil wrapped sample) should
be labeled with a unique number by which it may be readily identified in
the laboratory. This identification number should have as few digits as
possible to discourage abbreviation. The label should be waterproof, and
all information should be written in waterproof ink with a ballpoint pen.
The labels should include, in addition to the identification number, the
date and the initials of the sampling personnel.
It is highly recommended that a few scales be stored separately
and cross referenced by the identification number assigned to the tissue
specimen. For catfish and other scaleless fish, the pectoral fin spines
should be clipped and saved. These scales or spines may be stored by seal-
ing in small envelopes or plastic bags. This technique provides a means by
which the fish may be aged by a fisheries biologist if the need should
arise. Aging provides a good indication of the length of exposure to pol-
lutants.
Other pertinent information such as the time the sample was
taken, location, approximate depth, species, substrate, and water quality
(e.g., temperature, DO, pH) should be recorded in a field notebook. The
5-32
-------
data la this notebook must, of course, be cross referenced to the actual
sample by using the identification number, as previously discussed.
Preservation with dry ice (frozen C02) is recommended as a
means of ensuring that the sample is frozen rapidly and that it remains
frozen. This is very important to prevent decomposition and loss of vol-
atile materials. Minimum deterioration occurs if fish are frozen immedi-
ately after death and properly packaged to prevent loss of moisture and the
entrance of oxygen into the tissues (Royce, 1972; EPA, 1980). Dry ice is
the most effective means of rapidly freezing tissue during field sam-
pling.
Dry ice requires special packaging precautions before shipping
to comply with DOT regulations. The Federal Code of Regulations classifies
dry ice as ORM-A (Other Regulated Material). These regulations specify the
amount of dry ice which may be shipped by air transport and the type of
packaging required*
For any amount of dry ice. to be shipped by air, advance arrange-
ments must be made with the carrier. Not more than 440 pounds of dry ice
may be shipped by air freight unless special arrangements have been made
previously between the shipper and the aircraft operator. Quantities of
dry ice needed for tissue preservation are usually considerably less than
440 pounds.
The regulations further specify that the packaging oust be de-
signed and constructed in a manner to permit the release of carbon dioxide
gas which, if restricted, could cause rupture of the package. If samples
are being transported in a cooler, several vent-holes should be drilled to
allow for escape of the sublimated gas. The vents should be near the top
of the vertical sides of the cooler, rather than in the cover, to prevent
debris from falling into the cooler. Furthermore, wire screen or cheese-
cloth should be installed to help keep foreign materials from entering the
vents. When the samples are being packaged, care should be taken to keep
these vents open to prevent the buildup of pressure.
Dry ice is exempted from shipping paper and certification re-
quirements if the amount is less than 440 pounds and the package meets de-
sign requirements. The package must be marked "Carbon Dioxide, Solid" or
5-33
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"Dry Ice" and also marked with an identification that the material being
refrigerated is to be used for diagnostic or treatment purposes (e.g.,
frozen tissue).
Upon receipt at the laboratory, the tissue samples should be
placed in a freezer and maintained at a temperature less than -20"C (Royce,
1972; EPA, 1980) until the samples are prepared for analysis. All tissue
samples should be kept in their original packaging until they are ready to
be prepared.
When the samples are received at the laboratory, they should be
recorded in a permanent log book. This log book should include for each
sample date and time received, source of sample, sample number, how trans-
ported to the laboratory, and the number assigned to the sample by the lab-
oratory if this number differs from the field number. Although this re-
cording procedure may seem laborious, it Is absolutely Imperative that
precise records be kept for all samples so that the.data generated by the
sampling and analysis effort is of unquestionable Integrity.
An accurate written record should be maintained which can be used
to trace possession of the sample from the moment of its collection until
it has been analyzed. A chain of custody tag should be placed on all
coolers in which samples are stored and shipped. This should have
appropriate spaces for signatures when the sample is transferred from one
person to another. The date and time at which the custody is transferred
should be indicated on the tag.
5-34
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5.7 References
Battelle. 1975. Environmental Impact Monitoring of Nuclear
Power Plants: Source Book of Monitoring Methods, Vol. 2.
Atomic Industrial Forum, Inc. Washington, D.C.
Bennett, G.W. 1970. Management of Lakes and Ponds. Van
Nostrand Reinhold Company. New York. pp. 182-208.
Carlander, K.D. 1969. Handbook of Freshwater Fishes of the
U.S. and Canada, Exclusive of the Perciforaes* 3rd Edition.
Iowa State University Press. Ames, Iowa.
Lagler, K.F. 1956. Freshwater Fishery Biology. Wm. C.
Brown Co. Publications, Dubuque, Iowa.
McClane, A.J. 1978a. Field Guide to Freshwater Fishes of
North America. Holt, Rinehart, Winston, New York.
McClane, A.J. 1978b. Field Guide to Saltwater Fishes of
North America. Holt, Rinehart, Winston, New York.
Royce, W.F. 1972. Introduction to the Fishery Sciences.
Academic Press Inc., New York. pp. 214-214, 284-295.
U.S. Environmental Protection Agency. 1977* (Revised
October 1980). Interim Methods for the Sampling and
Analysis of Priority Pollutants in Sediments and Fish
Tissue. Environmental Monitoring and Support Laboratory.
Office of Research and Development. Cincinnati, Ohio.
U.S. Environmental Protection Agency. 1980. Draft Proto-
cols for the Analysis of Priority Pollutants. Methods 601 -
613, 624, and 625. Monitoring Technology Division. Office
of Research and Development. Washington. D.C.
Weber, C.I., editor. 1973. Biological Field and Laboratory
Methods for Measuring the Quality of Surface Waters and
Effluents. U.S. Environmental Protection Agency. Office of
Research and Development. Cincinnati, Ohio. 670/4-73-001.
5-35
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APPENDIX A
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PRIORITY POLLUTANTS
I. PESTICIDES
1. Acrolein
2. Aldrin
3. a-BHC (Alpha)
4. S-BHC (Beta)
5. Y-BHC (Lindane) (gamma)
6. 5-BHC (Delta)
7. Chlordane
3. DDD
9* DDE
10. DDT
11. Meldrin
12 a-Endosulfan (Alpha)
13. S-Endosulfan sulfate (Beta)
14. Endosulfan sulfate
15. Eadrin
16. Endrin aldehyde
17. Heptachlor
18. Heptachlor epoxide
19. Isophorone
20. TCDD (2,3,7,8-tetrachlorodibenzo-p-dioxin)
21. Toxaphene
II. METALS AND INORGANICS
22. Antimony
23. Arsenic
24. Asbestos
25. Beryllium
26. Cadmi urn
27. Chromium
28. Copper
29. Cyanides
30. Lead
31. Mercury
32. Nickel
33. Selenium
34. Silver
35. Thallium
36. Zinc
III. PCBs AND RELATED COMPOUNDS
37. PCB-1016 (Aroclor 1016)
38. PCB-1221 (Aroclor 1221)
39. PCB-1232 (Aroclor 1232)
40. PCB-1242 (Aroclor 1242)
-------
III. PCBa AND RELATED COMPOUNDS (Continued)
41. PCB-1248 (Aroclor 1248)
42. PCB-1254 (Aroclor 1254)
43. PCB-1260 (Aroclor 1260)
44. 2-Chloronaphthalene
IV,
HALOGENATED ALIPHATICS
V.
45
46
47
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
63.
64.
65.
66.
67.
68.
69.
70.
Methane,
Me thane,
Methane,
Ethane,
Methane, bromo- (methyl bromide)
Methane, chloro- (methyl chloride)
Methane, dichloro- (methylene chloride)
Methane, chlorodibromo-
Methane, dichlorobromo-
Methane, tribromo— (bromoform)
Methane, trichloro- (chloroform)
tetrachloro- (carbon tetrachloride)
trichlorofluoro-
dichlorodifluoro-
chloro-
Ethane, 1,1-dichloro-
Ethane, 1,2-dichloro-
Ethane, 1,1,1-trichloro-
Ethane, 1,1,2-trichloro-
Ethane, 1,1,2,2-tetrachloro-
Ethane, hexachloro-
Ethene, chloro- (vinyl chloride)
Ethene, 1,1-dichloro-
Ethene, trans-dichloro-
Ethene, trichloro-
Ethene, tetrachloro-
Propane, 1,2-dichloro-
Propene, 1,3-dichloro-
Butadiene, hexachloro-
Cyclopentadiene, hexachloro-
ETEERS
71. Ether, bis(choromethyl)
72. Ether, bls(2-chloroethyl)
73. Ether, bis(2-chloroiaopropyl)
74. Ether, 2-chloroethyl vinyl
75. Ether, 4-bromophenyl phenyl
76. Ether, 4-chlorophenyl phenyl
77. Bis(2-chloroethoxy) methane
VI. MONOCYCLIC AROMATICS (EXCLUDING PHENOLS, CRESOLS, PHTHALATES)
78. Benzene
79. Benzene, chloro-
80. Benzene, 1,2-dichloro-
-------
VI. MONOCTCLIC AROMATICS (EXCLUDING PHENOLS, CRESOLS, PHTHALATES)
(Continued)
81* Benzene, 1,3-dichloro-
82. Benzene, 1,4-dichloro-
83. Benzene, 1,2,4-trichloro-
84. Benzene, hexachloro—
85. Benzene, ethyl-
86. Benzene, nitro-
87. Toluene
88. Toluene, 2,4-dinitro-
89. Toluene, 2,6-dlnitro-
711. PHENOLS AND CRESOLS
90. Phenol
91. Phenol, 2-ehloro-
92. Phenol, 2,4-dichloro-
93. Phenol, 2,4,6-trlchloro-
94. Phenol, pencachloro-
95. Phenol, 2-nlcro-
96. Phenol, 4-nitro-
97. Phenol, 2,4-dinitro-
98. Phenol, 2,4-dlaethyl-
99. m-Cresol, p-chloro-
100. o-Cresol, 4,6-dinltro-
7III. PHTHALATE ESTEHS
101. Phthalate, dimethyl-
102. Phthalate, diethyl-
103. Phthalate, di-n-butyl-
104. Phthalate, dl-n-octyl-
105. Phthalate, bis(2-ethylhexyl)<
106. Phthalate, butyl benzyl-
II. POLYCTCLIC AROMATIC HYDROCARBONS
107. Acenaphthene
108. Acenaphthylene
109. Anthracene
110. Benzo(a)anthraeene
111. Benzo(b)fluoranthene
112. Benzo(1c)fluoranthene
113. Benzo(ghi)perylene
114. Benzo(a)pyrene
115. Chrysene
116. Dibenzo(a,H)anthracene
117. ?luoranthene
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IX. POLYCYCLIC AROMATIC HYDROCARBONS (Continued)
118. Fluorene
119. Indeno(l,2,3-cd)pyreae
120. Naphthalene
121. Phenanthrene
122. Pyrene
X. N1TROSAMINES AND OTHER NITROGEN-CONTAINING COMPOONDS
123. Nitroaanine, dimethyl- (DMN)
124. Nitrosaoine, diphenyl-
125. Nitroaamine, di-n-propyl-
126. Benzidine
127. Benzidine, 3,3f-dichloro-
128. Rydrazine, 1,2-diphenyl-
129. Acrylonitrile
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APPENDIX B
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RANGE AND HABITAT PREFERENCE OF TARGET SPECIES
Synopses of the range and habitat preference of target species are
provided below. This information Is summarized from McClane (1978a and
1978b). The target species are listed alphabetically by their common name.
Bluegill (Lepomis macroehirus)
Ranges from coast to coast. Widely distributed in farm ponds in
moat states* Prefers quiet, weedy waters with the larger fish remaining ia
deeper waters in the daytime and moving into shallow areas in morning and
evening to feed.
Brook Trout (Salvelinua fontinalis)
Native to northeastern North America from Georgia to the Arctic
Circle. Widely introduced. Found in rivers, streams, and lakes with a
preferred temperature range of 57 to 61"F.
Brown Trout (Salmo trutta)
European species introduced to the U.S. beginning in 1883* Pre-
fers streams with a temperature range from 55 to 64*F. Very active at
night. Found in cool streams and large lakes.
Carp (Cypriaus earpio)
Introduced into this country in 1876. Has become widely dis-
tributed from coast to coast. Prefers warm streams, lakes, and shallows*
Pollution tolerant.
Chain Pickerel (Esox niger)
Ranges from Maine to east Texas and north to the Great Lakes.
Found in lakes, ponds, and brackish creeks.
Channel Catfish (Ictalurua punctatua)
Great Lakes and Saskatchewan River south to Gulf of Mexico.
Widely introduced east and west of native range. Found in lakes and large
rivers with clean bottoms of sand, gravel, or boulders. Feeds primarily &e
night.
-------
Croaker (Micropcgon undulatus)
Distributed from Rhode Island to Cape Kennedy on the Atlantic
coast and from Tampa Bay to the southern Gulf of Campeche in the Gulf of
Mexico. In the winter, adults move out to deeper, warmer waters.
Freshwater Drum (Aplodinotua grunniens)
Ranges from Hudson Bay drainage and the Great Lakes east to Lake
Champlain and south to the Gulf. Abundant in some of the large, silty
lakes and rivers although the species prefers clean water.
Lake Trout (Salvelinua namaycuah)
•
This species is distributed in cold waters of the U.S. in New
England, the Finger Lakes, the Great Lakes, and scattered western lakes
where it has been introduced. Prefers deep, clear lakes.
Lake Whitefish (Coregonus clupeaformia)
Distributed in cold northern lakes of the U.S., particularly the
Great Lakes. Kay also eater rivers*
Largemouth Bass (Microptarus salmoides)
Ranges from Southeastern Canada through the Great Lakes, and
south in the Mississippi Valley to Mexico and Florida, and up the Atlantic
Coast through New England. Widely introduced in the east and west. Found
in shallow, weedy lakes or river backwaters and usually in water less than
20 feet deep.
Mountain Uhitefish (Prosopium williamsoni)
Distributed in lakes and streams on the western slope of the
Rocky Mountains from northern California to southern British Columbia.
Northern Pike (Baox lucius)
Ranges from New York through the Great Lakes to Nebraska.
Widely introduced in the south and west. Feeding is done during the
daylight hours.
-------
Pacific Staghorn Sculpin (Leptocottus armadus)
Ranges from northern Baja California to northwest Alaska. Found
in bays and inshore, also in and near freshwater at the mouths of streams'
Rainbow Smelt (Oamerua mordax)
Occurs on the Atlantic and Pacific coasts and in the Great Lakes
and other lakes in the northeast U.S. In coastal areas the species is sel-
dom found at depths greater than 20 feet.
Rainbow Trout (Salmo gairdneri)
Ranges from Mexican border to the Aleutian Islands. Found in
clear lakes and streams. Prefers temperatures below 70*7.
Round Whitefish (Prosopiurn eylindraeeum)
Distributed from New Brunswick northward to Ungava Bay and west-
ward through the Great Lakes to Alaska.
Sauger (Stizoatedion canadenae)
Distributed in the Great Lakes, large northern lakes, and in the
Mississippi, Missouri, Ohio, and Tennessee rivers. Often found in
tailwaters immediately below dams.
Silver Redhorse (Moxostoma anisurum)
Ranges from Manitoba to the St. Lawrence drainage and south to
northern Alabama and Missouri. Inhabits large streams, preferring long,
deep pools with slow currents. Tolerant of turbidity.
Smallmouth Bass (Mlcropterus dolomieui)
Ranges from Minnesota to Quebec and south to northern Alabama,
then west to eastern Kansas and Oklahoma. Widely introduced. Prefers
clear, rocky lakes with a minimum depth of 25 to 30 feet and temperatures
between 60 and 80°F in the summer. In streams, prefers a good percentage
of riffles flowing over gravel, boulders, or bedrock.
-------
Spot (Leiostomus xanthurus)
Distributed from Cape Cod to the Gulf of Mexico. Occur over mud
and sand bottoms and oyster beds. The species is more common in deepwatar
during fall and winter. Can tolerate a range of salinities from freshwater
to nearly twice that of seawater.
Spotted Sucker (Minytrema melanops)
Ranges from southern Minnesota to Pennsylvania and south to
Texas and Florida. Inhabits larger streams and lakes having sand, gravel,
or hard clay bottoms. Intolerant of turbid waters or heavy pollution.
Squawfish (Ptyehoeheilus oregonensAs)
Columbia River drainage and coastal streams of Washington and
Oregon.
Starry Flounder (Platlchthys stellatus)
Ranges from central California to Alaska. Prefers shallow water
and dandy bottoms. May enter brackish water and the mouths of rivers.
»
Stoneroller (Campostoma anomalum)
Ranges from Minnesota and Texas eastward. Found in streams and
rivers with a strong preference for riffles.
Striped Mullet (Mugll cephalus)
Found on both Atlantic and Pacific coasts. Atlantic coast range
is from the Gulf of Mexico to Cape Cod. Most abundant in southeastern U.S.
and Gulf of Mexico.
Walleye (Stizostedlan vitreum)
Original range was northern U.S. and Canada. Widespread stock-
ing has extended the range throughout the east and much of the south and
far west. Most common in large bodies of water. Prefers summer tem-
peratures below 85°F and clear water.
-------
Whitebait Smelt (Allosmerus elongatus)
Ranges from the Strait of Juan de Fuca to San Francisco.
White Perch (Morone amerieana)
Occurs in fresh, brackish, and saltwater from Nova Scotia to
North Carolina and inland as far as the Great Lakes. Congregates in the
deeper parts of bays and creeks during the winter.
White Sucker (Catostomus eommersoni)
Range extends fom Canada south to Florida and west to Montana.
Prefers large streams and the deeper water of impoundments. Often found
near dense weed beds. Pollution tolerant.
Winter Flounder (Pseudopleuroneetes americanus)
Occurs from Labrador to Georgia. Prefers muddy sand bottoms.
There is a movement into shallow water during the fall and an offshore
movement in the spring.
Yellow Perch (Perea flaveseens)
Ranges from Canadian border south to Kansas, Missouri, Illinois,
Indiana, and Ohio. Present in Atlantic drainage from Nova Scotia to South
Carolina. Widely stocked in the west. Prefers cool, clear water with sand
or rocky bottoms. Primarily found in lakes, although may be found in
rivers and brackish estuarine areas. Often stays in deeper waters during
the day and moves to shallows in the evening.
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