Unit«d Stitw
Environmental Prottction
Agtncy
Offici of Mann*
and Estuarin* Protection
Washington DC 20460
March 1987
EPA 430.9-86-004
W«t«r
Quality Assurance/Quality Control
(QA/QC) for 301 (h) Monitoring
Programs: Guidance on Field
and Laboratory Methods
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EPA Contract No. 68-01-6939
TC 3953-04
Final Report
QUALITY ASSURANCE/QUALITY CONTROL (QA/QC)
FOR 301(h) MONITORING PROGRAMS:
GUIDANCE ON FIELD AND LABORATORY METHODS
for
Marine Operations Division
Office of Marine and Estuarine Protection
U.S. Environmental Protection Agency
Washington, DC 20460
March, 1987
by
Tetra Tech, Inc.
11820 Northup Way, Suite 100
Bellevue, Washington 98005
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PREFACE
This manual has been prepared by EPA's Marine Operations Division,
Office of Marine and Estuarine Protection in response to requests from EPA
Regional Offices and coastal municipalities with sewage treatment plants
discharging into estuarine and marine coastal waters. The members of the
301(h) Task Force of EPA, which includes representatives for the EPA Regions
I, II, III, IV, IX and X, the Office of Research and Development, and the
Office of Water, are to be commended for their vif role in the development
of this guidance by the technical support contractor, Tetra Tech, Inc.
Under regulations implementing Section 301(h) of the Clean Water Act, munici-
palities are required to conduct monitoring programs to evaluate the impact
of their discharge on marine biota, to demonstrate compliance with applicable
water quality standards, and to measure toxic substances in the discharge.
The collection and analysis of high quality data require that specific,
established quality assurance and quality control (QA/QC) protocols be
adhered to in each of these major monitoring programs.
QA/QC procedures are included in this document for environmental variables
that may be measured in effluent, receiving water, sediment, and organism
tissues sampled during 301(h) monitoring programs. Quality assurance and
quality control procedures are provided for sample collection, field sample
handling, and laboratory processing to implement specific monitoring program
requirements provided in the 301(h) modified NPDES permit.
This manual is the result of several years of experience and effort in
designing, implementing, and reporting results of field and laboratory
monitoring programs with the avowed purpose of encouraging both national and
regional consistency in estuarine and marine monitoring programs. In this
regard, special appreciation is due to the members of the 301(h) Task Force
who contributed their valuable time and expertise in the development of this
manual.
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The information provided herein will be useful to U.S. EPA monitoring
program reviewers, permit writers, permittees, and other organizations
involved in performing nearshore monitoring studies. As the monitoring
variables included in t*vs document are commonly used in many marine and
estuarine monitoring programs, the guidance provided herein has broad applica-
bility beyond the 301(h) program.
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CONTENTS
Page
LIST OF FIGURES y
LIST OF TABLES vi
ACKNOWLEDGMENTS vii
INTRODUCTION ^
PURPOSE j
SCOPE !
FORMAT 2
EFFLUENT MONITORING 7
GENERAL METHODS 7
Sampling Preparation 7
Sampling Procedures 8
Sample Handling g
Field Procedures g
Sample Shipment 15
Laboratory Procedures 19
EFFLUENT ANALYSES 22
Flow 77
PH g
Temperature 28
Turbidity 30
Total Suspended Solids 34
Settleable Solids 37
Floating Particulates 39
Dissolved Oxygen (Winkler Method) 41
Dissolved Oxygen (Probe Method) 44
Biochemical Oxygen Demand (BOD) 47
Total Chlorine Residual 50
Oil and Grease 52
Nitrogen (Ammonia) 55
Nitrogen (Total Kjeldahl) 57
Nitrogen (Nitrate-Nitrite) 59
Phosphorus (Total) 61
Priority Pollutant Metals 63
Priority Pollutant Organic Compounds 68
Total and Fecal Coliform Bacteria 74
Enterococcus Bacteria 77
11
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MONITORING THE RECEIVING ENVIRONMENT 80
GENERAL METHODS 80
Sampling Preparation 80
Sampling Procedures 83
Station Location 83
Water Sampling 84
Grab Sampling 90
Trawl Sampling 97
Sample Handling 103
Field Procedures 103
Sample Shipment 113
Laboratory Procedures 114
Shipboard Laboratory Analyses 117
RECEIVING WATER ANALYSES 119
pH 120
Salinity 124
Temperature 127
Color 129
Transparency 131
Turbidity 132
Total Suspended Solids 135
Settleable So .lids 138
Floating Particulates 140
Dissolved Oxygen (Winkler Method) 142
Dissolved Oxygen (Probe Method) 145
Biochemical Oxygen Demand (BOD) 148
Oil and Grease 151
Nitrogen (Ammonia) 155
Nitrogen (Total Kjeldahl) 157
Nitrogen (Nitrate-Nitrite) 159
Phosphorus (Total) 161
Total and Fecal Coliform Bacteria 163
Enterococcus Bacteria 166
Chlorophyll a 169
Phytoplankton 172
SEDIMENT/INFAUNA ANALYSES 177
Grain Size 178
Total Solids/Water Content 181
Total Volatile Solids (TVS) 183
Total Organic Carbon (TOC) 185
Biochemical Oxygen Demand (BOD) 187
Chemical Oxygen Demand (COD) 190
Oil and Grease 192
111
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Sulfides (Total and Water Soluble) 196
Priority Pollutant Metals 199
Priority Pollutant Organic Compounds 205
Infauna 212
BIOACCUMULATION/TRAWL ANALYSES 234
Priority Pollutant Metals 235
Priority Pollutant Organic Compounds 243
Demersal Fishes and Megainvertebrates 251
REFERENCES 255
GLOSSARY 260
APPENDIX A RECOMMENDED METHODS FOR METALS IN EFFLUENT A-l
APPENDIX B RECOMMENDED METHODS FOR ORGANIC COMPOUNDS IN EFFLUENT B-l
1V
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FIGURES
Number Page
1 An example of chain-of-custody record 16
2 Examples of a sample analysis request form and a shipping seal 17
3 Deployment of a grab sampler 93
4 Examples of acceptable and unacceptable grab samples 95
5 Transect length as a function of urrent speed 101
6 An example of a chain-of-custody record 111
7 Examples of a sample analysis request form and a shipping seal 112
8 Construction of a sieve box 213
9 Example of a sieving stand 218
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TABLES
Number Page
1 Environmental variables included in this document 3
2 Recommended sample sizes, containers, preservation, and
holding times for effluent samples ' 10
3 Recommended methods for measuring effluent variables 12
4 Examples of problems frequently encountered during offshore
surveys and possible solutions to each problem 82
5 Recommended sample si':es, containers, preservation, and
holding times for offshore samples 104
6 Recommended methods for measuring offshore variables 106
7 Some common external abnormalities observed in fishes from
polluted areas . 254
A-l List of approved inorganic test procedures [note: this is a
reproduction of Table IB of U.S. EPA (1984)] A-l
B-l List of approved test procedures for non-pesticide organic
compounds [note: this is a reproduction of Table 1C of
U.S. EPA (1984)] B-l
8-2 List of approved test procedures for pesticides [note:
this is a reproduction of Table ID of U.S. EPA (1984)] B-3
vl
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ACKNOWLEDGMENTS
This document has been reviewed by the 301(h) Task Force of the U.S. EPA,
which Includes representatives from the Water Management Divisions of U.S. EPA
Regions I, II, III, IV, IX, and X; the Office of Research and Development
- Environmental Research Laboratory - Narragansett (located in Narragansett,
RI and Newport, OR), and the Marine Operations Division in the Office of
Marine, and Estuarine Protection, Office of Water. Among the reviewers,
the assistance provided by Dr. Steve Ferraro under the direction of Dr. Donald
J. Baumgartner, Dr. Dona". Phelps under the direction of Dr. Allan Beck,
and Dr. Brian Melzian under the direction of Patricia Eklund in coordinating
the numerous technical comments from the Office of Research and Development
and Region IX is gratefully acknowledged.
This technical guidance document was prepared by Tetra Tech Inc. for
the U.S. EPA Operations Division, Office of Marine and Estuarine Protection,
Office of Water) under the 301(h) post-decision technical support contract
No. 68-01-6938, Allison J. Duryee, Project Officer. This document was
prepared under the direction of Dr. Thomas Ginn (Program Director) and
Dr. Scott Becker (Work Assignment Manager) of Tetra Tech, Inc. The following
Tetra Tech staff members contributed to specific sections: Ms. Ann Bailey
(effluent variables, priority pollutant metals), Mr. Robert Barrick (priority
pollutant organic compounds), Dr. Scott Becker (effluent variables, water
column variables, demersal fishes and megainvertebrates, conventional sediment
variables), Dr. Gordon Bilyard (infauna, conventional sediment variables),
and Ms. Julia Wilcox (priority pollutant organic compounds, priority pollutant
metals). The following outside consultants also contributed to specific
sections: Mr. Jack Word (Evans Hamilton, Inc. - infauna), Dr. Victor Cabelli
(CABs Associates - microbiological variables), Ms. Kathy Krogslund and
Mr. Ron Citterman (University of Washington - water column variables).
vll
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INTRODUCTION
PURPOSE
This document was produced in response to specific regional requests
for assistance on technical issues raised during issuance of 301(h)-modified
NPDES permits. This technical g-idance to regional program offices and
permittees provides the framework for making informed decisions with respect
to the field and laboratory methods used when moo toring the effects of
sewage discharge on marine and estuarine ecosystems. Tht principal objectives
of this document are to ensure that:
• Samples are collected, processed, stored, shipped, and analyzed
using acceptable and standardized procedures
• Quality of generated data is documented adequately
• Results are reported completely and correctly
• Security of samples and data is maintained at all times.
SCOPE
The information presented in this document is designed to complement
the sampling and analysis specifications contained in the permits for individual
monitoring programs. Those specifications identify the sampling strategy
of each survey {e.g., station locations, field replication, time of sampling),
as well as which environmental variables will be measured. It is likely
that different monitoring programs will require different combinations
of variables to be measured, depending upon site-specific and discharge-
specific considerations. In this document, collection and analysis procedures
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are presented for most environmental variables that may be measured in
effluent, receiving water, sediment, and tissue during a 301(h) monitoring
program (Table I).
Information presented in this document can be used to address most
of the quality assurance/qual ity control (QA/QC) plan elements described
in "Guidance for Preparation of Combined Work/Quality Assurance Project
Plans for Environmental Monitoring" (U.S. EPA 1984). In the present document,
guidance is given for the following activities:
t Preparation for sampling
• Sample collection
• Sample processing
• Sample size
• Sample containers
• Sample preservation
• Sample holding times
• Sample shipping
• Logkeeping
• Labeling
• Custody procedures
• Analytical methods
• Calibration and preventive maintenance
• Quality control checks
• Corrective action
• Data reporting requirements.
FORMAT
This document is divided into two major sections: Effluent Monitoring
and Monitoring of the Receiving Environment. This division was used because
these two kinds of monitoring typically are not conducted by the same organ-
ization. For example, effluent sampling frequently is conducted by treatment
plant personnel, whereas sampling/of the receiving environment generally
Is conducted by outside contractors. Because of this possible separation,
each major section of this report is designed to stand alone. Thus, effluent
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TABLE 1. ENVIRONMENTAL VARIABLES INCLUDED IN THIS DOCUMENT
Matrix
Variable
Flow
PH
SaMnity
Temperature
Color
Turbidity
Transanssivity
Transparency
T' :al suspended solids
Se .tleable solids
Floating participates
Dissolved oxygen
Biochemical oxygen demand
Chemical oxygen demand
Total chlorine residual
Oil and grease
Nitrogen (ammonia)
Nitrogen (total Kjeldahl)
Nitrogen (nitrate-nitrite)
Phosphorus (total)
Sul fides
Priority pollutant metals
Priority pollutant organic compounds
Total and fecal col i form bacteria
Enterococcus bacteria
Chlorophyll £
Phy to plankton"
Grain size
Total solids
Total volatile solids
Total organic carbon
Infauna
Demersal fishes and
epibenthic macrolnvertebrates
Effluent
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
Receding
Water
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
Seabed/
Sediment Tissue
X
X
X
»
X
X X
X X
X
X
X
X '
X
X
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sa.-nplers will not have to si^t through information relevant only to those
who sample the receiving environment and vice versa. Although this independence
of sections leads to some redundancy, its overall efficiency outweighs
this drawback.
Within the effluent and receiving-environment monitoring sections,
collection and analysis procedures are presented first for general sampling
methods and then for specific environmental variables. General sampling
method sections include procedures *or 1) preparing for a survey, 2) collecting
t^e samples from which subsamples will be taken for specific variables
(e.g., effluent sampling, water sampling, grab sampling, trawl sampling),
and 3) handling samples in the field (labeling, chain-of-custi ly, storage,
shipping) and in the laboratory (reception, tracking, log keeping). Each
section on sample handling also contains tables that summarize the recommended
collection specifications and analytical methods for each environmental
variable.
The collection and analysis procedures for specific environmental
variables are organized to comply with most of the "elements of a QA project
plan" described by U.S. EPA (1984). The following elements are addressed
for each variable:
1. Field collection
2. Field processing
3. Analytical procedures
4. Calibration and preventive maintenance
5. Quality control checks
6. Corrective action
7. Data quality and reporting.
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1.) The section on field collection describes the recommended container
type, special cleaning procedures, and special collection techniques for
each variable.
2.) In the field processjng section, recommendations are made for
cne preservation technique and recommended maximum holding time before
analysis.
3.) The section on analytical procedures identifies the method(s)
recormended for laboratory analysis for each variable. The methods recommended
in this document take precedence over those identified in Tetra Tech (1985c).
To conform with U.c. EPA's equivalency policy, multiple methods were recommended
for effluent and receiving water variables if they have been shown to produce
equivalent results (U.S. EPA 1984). Multiple methods were confined to
tnose approved by U.S. EPA (1979b, 1984) and the American Public Health
Association (APHA 1985). Although other methods may provide equivalent
results (e.g., American Society for Testing and Materials, U.S. Geological
Survey) they generally were not recommended. However, if no U.S. EPA-
or APHA-approved methods were available for a particular effluent or receiving-
'*ater variable, an alternate source of methods was used. Because no U.S. EPA-
or APHA-approved methods are currently available for sediment and tissue
variables, alternate sources of methods were used for all of these variables.
In all cases, priority was given to methods that were developed under the
guidance of U.S. EPA (e.g., Plumb 1981; Tetra Tech 1986a,b). Methods for
collection and analysis of benthic infauna were described in considerably
greater detail than those for other variables because of the importance
of these organisms as indicators of sewage-related biological impacts and
because of the lack of a published reference having adequate depth and
scope. In addition to specifying the recommended analytical method(s),
the section on analytical procedures describes the major sources of interference
or error that may be encountered when using the methods. This kind of
information was included because it often is incorporated into the method
description without being highlighted and because some methods do not treat
it as comprehensively as others.
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4.) The section on cal Ibration and preventive maintenance describes
the kind and frequency of calibration and equipment maintenance that are
essential to providing quality data for each variable.
5.) In the section on quality control checks, the kind and frequency
of quality control checks (e.g., duplicates, blanks, spiked samples, reference
material analyses) that should be conducted during laboratory analyses
are described for each variable. A duplication level of 10 percent was
considered appropriate for most variaoles. Blanks and spiked samples
generally were recarmended at a frequency of one per batch. It is recommended
that duplicate analyses be distributed relatively evenly throughout the
full sequence of samples analyzed, rather tr in being concentrated at the
beginning or end of the sequence.
6.) The section on corrective action describes major sources of analytical
errors that may be encountered when analyzing for each variable. Possible
solutions also are recommended.
7.) The section on data qua! ity and reporting describes documented
levels of accuracy and precision (if available) achieved using the analytical
method, the kinds of information that should be reported to U.S. EPA, and
the units and significant figures that should be used when reporting results.
Specifications in this section are compatible with the data requirements
of the 301(h) program. Further guidance on the data reporting requirements
is available in the "GOES Data Submissions Manual" (Tetra Tech and American
Management Systems 1985).
Because this document considers a wide range of disciplines, the term-
inology used in certain sections may be unfamiliar to various readers.
Therefore, a glossary is provided at the end of the document to define
specialized words or phrases.
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EFFLUENT MONITORING
Recommended methods for measuring effluent variables during 301(h)
monitoring programs are described in this section. The initial major section
includes general procedures for pre-sampling activities, sampling, and
sample handling. The second major section presents detailed procedures
for measuring 20 effluent variables. The general section on sample handling
contains tables that summarize the recommended collection specifications
and analytical methods for the spe ific effluent variables. Tables of
recommended methods for metals and organic compounds in effluent are also
presented in the appendix.
GENERAL METHODS
Sampling Preparation
The QA/QC Project Plan should be thoroughly reviewed in advance by
sampling personnel to identify the following:
t Responsibilities for each individual associated with the
sampling and analysis
• Statement and prioritization of study objectives
• Background information, sampling locations, sampling frequency,
and sampling procedures
• Variables to be measured and corresponding required sample
sizes, containers preservatives, and holding times
• Sample splits or performance samples to be submitted with
the samples
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• Laboratories to -which samples will be shipped
• Shipping requirements.
The study objectives and their prioritization should be understood
by all personnel involved in the sampling effort. This will ensure that
if modifications of the plan become necessary, their impact on the overall
goals of the sampling effort can be adequately evaluated.
i'o ensure that all required sampling equipment and supplies are available
at the time of sampling an equipment checklist should be constructed.
Spare parts and backup supplies (e.g., extra glass jars, spare pH probe)
should be included in the inventory.
Sampling Procedures
Sample collection equipment, sampling frequency, and sample locations
depend upon the size and nature of the discharge and generally are specified
in the discharge permit. Decisions regarding equipment to be used during
the monitoring program should be made after reviewing related literature
(e.g., Tetra Tech 1982; U.S. EPA 1982). If automatic samplers are used
for collection of flow-proportioned 24-h composite samples, the sampler
should be selected on the basis of the analyses to be performed. The automatic
sampler must be able to collect adequate sample sizes without introducing
contaminants. For example, plastic tubing and bottles cannot be used to
collect samples for priority pollutant organic compounds.
Grab samples, rather than composite samples, must be taken for certain
variables (e.g., dissolved gases, volatile compounds, microbiological variables)
because of changes that are likely to occur during storage. Effluent variables
that must be collected as grab samples are pH, temperature, total and fecal
coliform bacteria, dissolved oxygen, oil and grease, and volatile organic
compounds. For further sample collection requirements, the section on
field collection should be consulted for each variable. Note that certain
variables (e.g., volatile organic compounds) require filling sample containers
completely, leaving no air space, whereas other variables (e.g., total
8
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and fecal coliform, oil and grease) require leaving air space (i.e., headspace)
to allow for adequate mixing or addition of acid.
Sample Handling
Proper sample handling ensures that changes in the constituents of
interest are minimized and guards against errors when collecting, shipping,
and analyzing samples. Recommended sample size, container, preservation
and storage requirements for each effluent variable are summarized in Table 2.
Recommended laboratory methods for each effluent variable are listed in
Table 3. These requirements are also noted in subsequent sections. These
requirements and methods should be reviewed in advance j, laboratory personnel
to ensure that the sample size, containers, preservatives, and all other
specifications are consistent with the needs and capabilities of the laboratory.
Field Procedures-
It is important throughout any sampling and analysis program to maintain
the integrity of the sample from the time of collection to the point of
data reporting. Proper chain-of-custody procedures allow the possession
and handling of samples to be traced from collection to final disposition.
It is recommended that chain-of-custody procedures be used for all sampling
conducted during 301(h) monitoring programs. The documents needed to maintain
proper chain-of-custody include:
t Field logbook — All pertinent information on field activities
and sampling efforts should be recorded in a bound logbook.
The field supervisor is responsible for ensuring that sufficient
detail is recorded in the .logbook. The logbook should enable
someone else to completely reconstruct the field activitiy
without relying on the memory of the field crew. All entries
should be made in indelible ink, with each page signed and
dated by the author, and a line drawn through the remainder
of any page. All corrections should consist of permanent
line-out deletions that are initialed by the field supervisor.
At a minimum, entries in a logbook should include:
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TABLE 2. RECOMMENDED SAMPLE SIZES, CONTAINERS, PRESERVATION,
AND HOLDING TIMES FOR EFFLUENT SAMPLES*
Minimum
Sample
Slzeb
Measurement («l)
pH 25
Temperature 1 .000
Turbidity 100
Total suspended solids 1,000
Settleable solids 1.000
f\? ing pa.-ticulates 5.000
Dissolved oxygen
Probe 300
Kinkier 300
Biochemical oxygen
demand 1 .000
Total chlorine
residual 200
Oil and grease I. 000
Nitrogen
Ammonia-N 400
Total Kjeldahl-N 500
Nitrate * Nitrite-N 100
Phosphorus (total) 50
Priority pollutant metals
Mercury 100
Metals, except mercury 100
Priority pollutant
organic compounds
Ex tractable compounds 4,000
(includes phthalates,
nitrosamines, organo
rhlnrlnp ne«Meide«.
ContalnerC
P.6
P.6
P.S.
P.G
P.G
P.G
G bottle & top
6 bottle ft top
P.G
P.G
6 only
P,G
P.G
P.G
P. 6
P.G
P.G
6 only.
TFE- lined
cap
Preservative^
None
None
Cool. 40c
Cool, 40c
Cool. 4oc
None
None
Fix on site; store
in dark
Cool. 40c
None
Cool, 40c
H2S04 to pH<2
Cool , 40c HjSOa to
pH<2
Cool . 4»c H2S04 to
pH<2
Cool . 4% H2S04 to
PH<2
Cool. 40c H2SO« to
pH<2
HN03 to pH<2
HN03 tc pH<2
Cool . 40 c
0.0081 NajSzOjg
Store in dark
Maiimum
Holding
Time
Analyze immediately^
Measure immediately^
48 h
7 days
48 h
Analyze Immediately6*
Analyze immediately6
8 h
48 h
• Analyze immediately0
28 days
28 days
28 days
28 days
28 days
28 days
6 mo
7 days until
extraction
40 days after
extraction
PCBs, nitroaromatics,
isophorone, polynuclear
aromatic hydrocarbons,
haloether, chlorinated
hydrocarbons, phenols,
and TCOD)
Purgeable compounds
40
G only, TfE- Cool, 4»C
lined septum 0.0081
7 days"
10
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TABLE 2. (Continued)
Total and fecal
conform bacteria 250-500
Enterococcus bacteria 250-500
P. 6 Cool. 40 c
0.0081 NajS^Q
P. 6 Cool, 40 C
0.0081 Ma2S203g
6 h
6 h
a Reference: Adapted from U.S. EPA (1979b. 1984).
0 Recommended field sample sizes for one laboratory analysis. If additional laboratory analyses
are required (e.g.. replicates), the field sample size should be adjusted accordingly.
c Polyethylene (P) or Glass (6)
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TABLE 3. RECOMMENDED METHODS FOR MEASURING EFFLUENT VARIABLES
Method Reference
Variable
pH
Temperature
Turbidity
Total suspended solids
Settleable solids
Floating particulates
Dissolved oxygen
Probe
Uinkler
Biochemical oxygen demand
Total chlorine residual
Oil and grease
Nitrogen
Ammonia-N
Total Kjeldahl-N
Nitrate-t-Nitrite-N
U.S. EPAa
150.1
170.1
180.1
IjQ.Z
160.5
-
360.1
360.2
405.1
330.1
330.2
330.3
330.4
330.5
413.1
350.1
350.2
350.3
351.1
351.2
351.3
351.4
353.1
353.2
353.3
APHAb
423
212
21 4A
214B
209C
209E
206Ad
421F
421B
507
408A
408B
408C
4080
408E
503A
41 7A
417B
4170
417G
417B
417D
417E
420A
4208
418C
41 8F
Otherc
-
-
-
-
-
-
-
-
-
-
12
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TABLE 3. (Continued)
Phosphorus (total)
Priority pollutant metals
Priority pollutant
Organic compounds
Total coliform bacte -ia
Fecal coliform bacteria
Enterococcus bacteria
365.1
365.2
365.3
365.4
Table 1B&
U.S. EPA
(1934)
Table ice
U.S. EPA
(1984)
-
-
-
42 4C
42 4F
42 4G
Table IBS
U.S. EPA
(1984)
Table ice
U.S. EPA
(1984)
908Af
909A9
908Cf
909C9
-
™
p. 114f,h
p. lOSg.h
p. 132f,h
p. 1249,h
U.S. EPA1
a Methods recommended in U.S. EPA (1979b).
D Methods recommended in APHA (1985).
c Methods recommended in sources other than U.S. EPA (1979) or APHA (1985)
when no methods were recommended in the latter two sources.
d This method is tentatively recommended by APHA.
e The list of U.S. EPA and APHA methods for individual components of this
group are listed in the table specified and are too extensive to include
here.
f This method can be used whether or not chlorine is present.
9 This method can be used only when chlorine is absent.
h Page reference of this method in Bordner et al. (1978).
i U.S. EPA is currently finalizing a recommended method for this variable.
13
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Date and time of starting work
Names of field supervisor and team members
Purpose of proposed sampling effort
Description of sampling site, including information
on any photographs that may be taken
Location of sampling site
Details of actual sampling effort, particularly deviations
from standard operating procedures
Field observations
Field measurements made (e.g., pH, temperature, flow)
Field laboratory analytical results
Sample Identification
Type and number of samples collected
Sample handling, packaging, labeling, and shipping
information (including destination).
Chain-of-custody procedures should be maintained with the
field logbook. While being used in the field, the logbook
should remain with the field team at all times. Upon completion
of the sampling effort, the logbook should be kept in a
secure area. All logged information should be summarized
and submitted to U.S. EPA after sampling is completed.
14
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t Sample labels -- Sample labels must be waterproof and must
be securely fastened to the outside of each sample container
to prevent misidentlfication of samples. Labels should
contain at least the sample number, preservation technique,
date and time of collection, location of collection, and
signature of the collector. Labels should be marked with
Indelible Ink and placed on the body of the jar. Abbreviated
labels may also be placed on the cap of each jar to facilitate
sample Identification.
t Chain-of-custody records -- A chain-of-custody record (Figure 1)
must accompany every group f samples. Each person who
has custody of the sample onst sign the form and ensure
that the samples are not left unattended unless secured
properly.
• Custody seals -- Custody seals (Figure 2) are used to detect
unauthorized tampering with the samples. Sampling personnel
should attach seals to all shipping containers sent to the
laboratory by common carrier. Gummed paper seals or custody
tape should.be used so that the seal must be broken when
the container is opened.
For further information regarding proper chain-of-custody procedures, consult
the policies and procedures manual for the National Enforcement Investigations
Center (NEIC; U.S. EPA 1978).
Sample Shipment-
All preserved samples should be shipped immediately after completion
of sampling. This minimizes the number of people handling samples, and
protects sample quality and security. Guidance for shipping hazardous
materials can be found in U.S. Department of Transportation (1984). As
samples are prepared for shipping, the following should be kept in mind:
15
-------
Uniud SutH
Environment* Protection
Agency
RtponlO
1200 SlJrth AOTMM
$••«)• WA 98101
CHAIN Of CUSTODY RECORD
PROJECT
LAB *
STATION
DATE
TIME
RELINQUISHED BY: O««M»
RCUNOUI8HED BY: «*•«•»
REUNQUI8HED BY: /S*«M
RELINQUISHED BY: '!»—*
DISPATCHED BY: «w— *
METHOD OF SHIPMENT:
DATA
SAMPLERS: <&?•*•
SAMPLE TYPE
I
m
m
a
1
5
9
s
a
n
5
REMARKS
MCIIVtD BY: ft+~*
RECEIVED BY: av^um
RECEIVED BY: ts*~~
RECVO BY MOBILE LAB FOR WELD
ANAL.: is****
fTIMI
RECEIVED FOR LAB BY: r*v«—
DATEH1MI
DATE/TIME
DATE/TIME
DATimMB
DATE/TIME
Ontnbuoon
On* Coo»-Surv«v Coaidniiar ftta f»m
Figure 1. An example of * chain-of-custody record. An equiva-
lent form should be used for 301(h) monitoring.
16
-------
ENVIRONMENTAL PROTECTION AGENCY
CLP Sample Management OH ice
P.O. Boa SIS • Alexandria. Virginia 22313
Phone: 703/J37-2MO - FTS/3J7-2»90
SAS Number
SPECIAL ANALYTICAL SERVICE
PACKING LIST
Sampling Officei
Sampling Contacts
(name)
(phone)
Sample
Number*
Sampling Datefsh
Date Shipped:
Site Name/Code!
Ship Tm
AttK
For Lab UM Only
Date Sample* Rec*di
Received By:
Sample Description Sample Condition on
LA, Anaiyiis, UatrU, Concentration Receipt at Ub
1. _
2. .
J. .
a. _
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4. .
7. .
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For Ublte Only
EPA CUSTODY SEAL
KNYiROMUMTAL PtOTICTMN ACtNCY
Meri MM Afl» 71 TUB «•<
*
si
Figure 2. Examples of a sample analysis request form (above)
and a custody seal (below). Equivalent material
should be used for 301(h) monitoring.
17
-------
• Shipping containers should be in good shape and capable
of withstanding rough treatment during shipping.
• Samples should be packed tightly:
Dividers must separate all glass containers
Empty space within shipping boxes should be filled so that
jars are held securely.
• All containers must be leak-proof. If a container is not
leak-proof by design, the interior should be lined with
two heavy-duty plastic bags and the tops of bags should
be tied once samples are inside. Adequate absorbent material
should be placed in the container in a quantity sufficient
to absorb all of the liquid shipped.
•
t All samples should be accompanied by a sample analysis request.
Variables to be analyzed by the laboratory, and total number
and kind of samples shipped for analysis should be listed
on the request sheet. An example sample analysis request
form is illustrated in Figure 2. The laboratory should
acknowledge receipt of shipment by signing and dating the
form, and returning a copy to the designated QA coordinator.
• A chain-of-custody record for each shipping container should
be filled out completely and signed.
• The original chain-of-custody record and analysis request
should be protected from damage and placed inside the shipping
box. A copy of each should be retained by the shipping
party.
• The custody seal should be attached so that the shipping
box cannot be opened without breaking the seal.
18
-------
• For shipping containers carrying liquid samples:
A "This End Up" label should be attached to each side
to ensure that jars are transported in upright position
A "Fragile-Glass" label should be attached to the top
of the box to minimize agitation of samples.
• Shipping containers should be sent by a carrier that will
provide a delivery receiot. This will confirm that the
contract laboratory received the samples and serve as a
backup to the chain-of-custody record.
• All shipping charges should be prepaid by the sender to
avoid confusion and possible rejection of package by the
contract laboratory.
Laboratory Procedures--
At the laboratory, one person should be designated custodian of all
incoming samples. An alternate should also be designated to serve In the
custodian's absence. The custodian should oversee the following activities:
• Reception of samples
• Maintenance of chain-of-custody records
• Maintenance of sample tracking logs
• Distribution of samples for laboratory analyses
t Sending samples to outside laboratories
19
-------
t Supervision of labeling, log keeping, data reduction, and
data transcription
• Storage and security of all samples, data, and documents.
Upon reception of samples, a designated laboratory person should fill
out the chain-of-custody record, indicating time and date of reception,
number of samples, and condition of samples. All irregularities indicating
that sample security or quality may hav«» been jeopardized (e.g., evidence
of tampering, loose lids, cracked jars) should be noted on the sample analysis
request form and returned to the client-designated QA :oorrfinator. In
addition, a designated person should initiate and maintain a sample-tracking
log that will follow each sample through all stages of laboratory processing
and analysis. Minimum information required in a sample-tracking log includes:
• Sample identification number
t Location and condition of storage
• Date and time of each removal from and return to storage
t Signature of person removing and returning the sample
• Reasons for removal from storage
• Final disposition of sample.
All logbooks, labels, data sheets, tracking logs, and custody records
should have proper identification numbers and be filled out accurately.
All information should be written in ink. Corrections should be made by
drawing a line through the error and entering the correct information.
Corrections should be signed and dated. Accuracy of all data reductions
and transcriptions should be verified at least twice. All samples and
documents should be properly stored within the laboratory until the client
authorizes their removal. Security and confidentiality of all stored material
should be maintained at all times. Before releasing analytical results,
20
-------
a',1 information on sample tags, data sheets, tracking logs, and custody
records should be cross-checked to ensure that data pertaining to each
sample are consistent throughout the record.
Originals of the following documents should be sent to the client:
t Chain-of-custody records
t Sample-tracking logs
• Data report sheets
• Quality contiol records.
Copies of all forms should be retained by the laboratory in case originals
are lost in transit.
When replicate analyses are conducted as QA/QC checks, it is recoimended
that they be distributed relatively evenly throughout the full sequence
of samples analyzed, rather than being concentrated at some point (e.g.,
beginning, end) of the sequence. This precaution will enhance the probability
that if problems develop within part of a sequence, they will be detected.
21
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EFFLUENT ANALYSES
In this section, QA/QC procedures are presented for the following
20 effluent variables:
• Flow
• pH
• Temperature
• Turbidity
• Total suspended solids
• Settleable solids
• Floating particulates
• Dissolved oxyg:r. (Winkler metiiod)
t Dissolved oxygen (probe method)
• Biochemical oxygen demand (BOD)
• Total chlorine residual
• Oil and grease
• Nitrogen (ammonia)
• Nitrogen (total Kjeldahl)
• Nitrogen (nitrate-nitrite)
• Phosphorus (total)
• Priority pollutant metals
• Priority pollutant organic compounds
• Total and fecal coliform bacteria
• Enterococcus bacteria.
22
-------
Effluent
Flow
Flow
Accurate flow measurement Is essential for proper operation and control
of wastewater treatment plants. Flow determinations are used for a variety
of purposes, including 1) provision of data with which plant operation
and performance can be evaluated, 2) provision of data for long-term planning
of treatment plant capacity relative to utilized control capacity, and
3) determination of compliance with permitted effluent mass-loading limits.
Flow can be measured jri situ using \ variety of methods. The two
major categories of methods are direct-discharge and velocity-discharge
(Metcalf and Eddy 1979). The direct-discharge methods are used most frequently
for measuring wastewater flows. These methods relate the rate of discharge
to one or two easily measured variables and employ such devices as weirs,
Parshall flumes, Venturi meters, and magnetic flow meters. Accuracy of
these direct-discharge methods generally ranges from +3 to +5 percent of
the flow rate (Hinrichs 1979). Any system that cannot measure wastewater
flow within +10 percent is considered unacceptable for NPOES compliance
(U.S. EPA 1977).
The preferred location for a flow-measuring device is at the end of
all treatment processes and downstream front all sidestreams. If the device
is located at the head of the treatment process, the flow must be corrected
for all return streams that originally discharged upstream from the flow-
measuring device. For continuous flow measurements, a system including
the primary flow device, a flow sensor, transmitting equipment, a recorder,
and possibly a totalizer are recommended.
Flow-measuring equipment should be routinely inspected, calibrated,
and maintained to ensure that accurate data are produced. At a minimum,
equipment should be serviced according to the frequency recommended by
the manufacturer. Servicing should be more frequent if the equipment does
not appear to be functioning properly. A common cause of improper equipment
operation is accumulation of foreign material in or on equipment components.
23
-------
Effluent
Flow
Flow should be reported as millions of gallons per day (MGD). Where
continuous monitoring of an effluent is required, flew measurements should
include the total daily flow and the peak daily flow on a 24-h basis.
-------
Effluent
PH
£H
Field Procedures--
Col 1ection--Samp1es for pH determination should be collected in poly-
ethylene or glass bottles having airtight screw caps. Because pH is unstable
and cannot be preserved, these samples should be analyzed as soon as possible
after collection. Because pH of waters not -,t equilibrium with the atmosphere
may change upon exposure to the atmosphere, sample containers should be
completely filled and tightly sealed after collection.
Prior to filling, each sample bottle and cap should be rinsed thoroughly
with sample water. This can be achieved by filling the bottle halfway,
sealing and shaking It, and rinsing the stopper as the wash water is discarded.
Process ing--Because pH cannot be preserved, samples should be analyzed
immediately after collection. If a short delay occurs, the samples should
be stored in the dark at 4° C and the storage time should be noted on the
log sheet.
Laboratory Procedures--
Analytical Procedures—Analytical procedures are given in U.S. EPA
Method 150.1 and APHA Method 423. As noted previously, pH samples should
be analyzed as soon as possible following collection.
Several potential sources of interference with pH measurements should
be avoided. Because the response of the electrode can be impaired if it
is coated with oily or particulate material, the electrode should be gently
blotted or washed with a detergent periodically. Treatment with hydrochloric
acid may be necessary to remove some kinds of film. Temperature can influence
pH measurements by altering electrode output and by changing the pH inherent
in the sample. The first source of temperature interference can be controlled
by using a pH meter having temperature compensation or by calibrating the
25
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Effluent
pH
meter at the temperature of the samples. Because the second source of
temperature interference cannot be controlled, the temperature at which
the pH determination of each, sample is made should be logged and reported.
It is recommendec that sample temperature never differ by more than 2° C
from that of the buffer solution.
When pH measurements are being made, it is critical that the sample
be stirr-Kl at a constant rate to provide drift-free (<0.1 pH units) measure-
ments. The rate of stirring should min-mize air transfer at the surface
of the sample. At least 30 sec should be allowed for each measuremen
to stabilize.
Calibration and Preventive Maintenance—Calibration procedures should
follow specifications given by the manufacturer of the pH meter. General
guidelines are given by U.S. EPA (1979b) and APHA (1985).
Primary buffer salts should be those of the National Bureau of Standards
(NBS). Secondary buffer salts can be prepared from the NBS salts or purchased
as a solution that has been calibrated to NBS salts. The pH meter should
be calibrated at a minimum of two points that bracket the expected pH value
of the samples and that are three or more pH units apart (U.S. EPA 1979b).
Prepare fresh buffer solutions at least every month to avoid erroneous
calibration due to mold growth or contamination.
Preventive maintenance procedures should follow specifications given
by the manufacturer of the pH meter. In general, verification of electrode
performance and meter performance is the only operator service recommended.
An electrode should be replaced when it no longer meets span requirements
and does not improve with rejuvenating procedures.
Quality Control Checks—The pH meter should be calibrated at the beginning
of each series of samples and after each group of 10 successive measurements.
It is recommended that duplicate pH determinations be made on at least
26
-------
Effluent
PH
10 percent of the total number of samples. As an independent check, a
U.TJ. EPA reference sample should be analyzed at a minimum of every 3 mo.
Corrective Action—If the pH meter does not appear to be operating
correctly, consult the manufacturer's troubleshooting guide. Common problems
include a dirty electrode, failure to fill the reference portion of the
electrode with internal solution, and inadequate stirring.
Data Quality and Reporting—A precision of +_ 0.02 pH unit and an accuracy
of 0.05 pH unit can be achieved under the best circumstances. However,
the '.imit of accuracy under most circumstances is ^0.1 pH unit (APHA 1985).
A precision of 0.1 pH unit is considered acceptable (U.S. EPA 1979b).
Measurements of pK are reliable only when the instrument has been
calibrated by standard buffers bracketing the-desired range. Samples having
a pH greater than 10 may require a special probe to correct for interference
from sodium ions.
It Is recommended that pH values be reported to the nearest 0-. 1 unit.
In addition, the ambient temperature at the time of measurement of each
sample should be reported to the nearest degree C. The results of all
determinations should be reported, including QA replicates. Any factors
that may have influenced sample quality should also be reported.
27
-------
Effluent
Temperature
Temperature
Field Procedures—
Col lection—Temperature can be measured using a mercury-filled Celsius
thermometer on samples collected in glass or plastic containers. The ther-
mometer should have a scale etched .jn capillary glass for O.io C increments
and a minimal thermal capacity to permit rapid equilibration. Temperature
can be measured in situ using a reversing thermome >r or a thermistor.
Of these two in situ instruments, the thermistor is more accurate, but
also more expensive.
Processing—Because temperature can change rapidly after a sample
is removed from ambient conditions, temperature determinations by thermometer
should be made immediately after sample collection.
Laboratory Procedures—
Analytical Procedures—Methods for making temperature measurements
are described in U.S. EPA Method 170.1 and in APHA Method 212. It is critical
that the measuring device be adequately immersed in the sample and allowed
to completely equilibrate (i.e., the temperature reading stabilizes) before
temperature is read.
Calibration and Preventive Maintenance—Each kind of temperature-measuring
instrument should be calibrated frequently against a National Bureau of
Standards (NBS)-certified thermometer that is used with its certificate
and correction chart. An NBS thermometer is recommended because some commercial
thermometers may be as much as 3° C in error (APHA 1985).
To prevent breakage, it is recommended that each thermometer be enclosed
in a metal case. If a mercury thermometer is broken, samples or bottles
in the vicinity of the exposed area may be contaminated by the mercury.
28
-------
Effluent
Temperature
Quality Control Checks—Each temperature-measuring instrument should
be calibrated against an NBS thermometer at least every week. It is recommended
that calibration should be conducted daily when a temperature violation
is suspected.
Corrective Action—If the temperature-measuring instrument cannot
be calibrated consistently against the NBS thermometer, it should be repaired
or replaced.
Data Quality and Reporting—Precision and accuracy have not been determined
for temperature measurements (U.S. EPA 1979b). Temperature measurements
should be reported to the nearest O.io C. The results of all determinations
should be reported, including QA replicates and calibration checks. Any
factors that may have influenced sample quality should also be reported.
29
-------
Effluent
Turbidity
Turbidity
Field Procedures--
Col lection--Turbidity samples can be collected in glass or plastic
containers. Samples should be removed from the sampler as soon as possible
after collection to minimize settling of suspended material within the
sample''. Sample containers and lids should be rinsed thoroughly with sample
water before samples are collected.
Process ing--Because turbidity samples cannot be preserved adequately,
they should be analyzed as soon as possible after collection. If a delay
occurs, samples should be held at 40 c for no more than 48 h to minimize
microbiological decomposition of solids. The length of delay should be
noted on the log sheet.
Laboratory Procedures —
Analytical Procedures—Two procedures are available for turbidity
measurements in effluent: the nephelometric method and the visual method
(APHA 1985). Although the results of both methods are comparable in a
general manner (U.S. EPA 1979b), they are not related directly (AHPA 1985)
and therefore should not be used interchangeably. Because of its greater
precision, sensitivity, and applicability over a wide turbidity range, the
nephelometric method is generally preferable to the v'isual method (APHA 1985).
The nephelometric method is based on the nephelometric turbidity unit
(NTU), and is described in U.S. EPA Method 180.1 and in APHA Method 214A.
For turbidities greater than 40 NTU, samples should be diluted with one
or more volumes of turbidity-free water until the turbidity falls below
40 NTU.
30
-------
Effluent
Turbidity
The visual method of turbidity determination is based on the Jackson
turbidity unit (JTU), and is described in APHA Method 214B. It currently
is the method specified by the California Ocean Plan. The lowest turbidity
that can be measured using a candle turbidimeter is 25 JTU (APHA 1985).
Samples with turbidity values less than 25 JTU must be estimated indirectly
by visual comparison with standards. Samples having turbidity values exceeding
1,000 JTU should be diluted with one or more volumes of turbidity-free
water until turbidity falls below 1,000 JTU.
Turbidity-free water is distilled wa:er passed through a membrane
filter having a pore size of 0.2 urn. Prior to analysis, each sample should
be shaken well to thoroughly disperse solids and resulting air bubbles
should be allowed to dissipate before the sample is analyzed.
Interference with turbidity measurements arises from several sources.
The presence of floating debris and coarse sediments that settle out rapidly
will give low readings. Therefore, turbidity readings should be made as
soon as possible after sample agitation. Finely divided air bubbles will
affect results in a positive direction. If present, the bubbles should
be allowed to dissipate before taking the reading. Finally, dissolved
substances in the sample, which absorb light, will reduce turbidity readings.
Any color in the sample should therefore be noted.
Calibration and Preventive Maintenance—It is recommended that standard
suspension of formazin be used to calibrate the nephelometer. Formazin
provides a more reproducible turbidity standard than do other materials
used in the past. The formazin standard suspension should be prepared
daily (APHA 1985). Commercially available standards, such as styrene divinyl-
benzene beads (trade name AMCO-AEPA-1), can be substituted for formazin
if they are demonstrated to be equivalent to freshly prepared formazin
(APHA 1985). Standards measured on the nephelometer should cover the range
expected for the samples. At least one standard should be run in each
instrument range to be used. The instrument should give stable readings
31
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Effluent
Turbidity
in all sensitivity ranges used. If a precalibrated scale is not supplied,
calibration curves should be prepared for each instrument range to be used.
Standards for visual comparisons based on JTU should be prepared from
natural turbid water or kaolin, and should be calibrated weekly using the
candle turbidimeter. The candle used in the turbidimeter should be made
of beeswax and spermaceti, and should burn within the limits of 114-i26
grains/h.
Quality Control Checks—The nepheloneter should be calibrated at the
start of each series of analyses and after each group of 10 successive
samples. Duplicate analyses should be conducted on at least 10 percent
of the total number of samples, using either method of turbidity determination.
Corrective Action—If the nephelometer wiM not stabilize in any of
the relevant ranges or if the instrument does not appear to be functioning
properly in any other aspect, the manufacturer's troubleshooting guide
should be consulted. Sample tubes that become scratched or etched should
be replaced.
Data Quality and Reporting—Limited precision data for the nephelometric
method indicate that standard deviations of measurements vary directly
with the level of turbidity (U.S. EPA 1979b). Accuracy data are not available
at present. The sensitivity of the nephelometer should allow detection
of a turbidity difference of 0.02 unit or less in waters having turbidities
less than 1.0 unit. Results should be reported in nephelometric turbidity
units (NTU). U.S. EPA Method 180.1 and APHA Method 214A describe the nearest
reporting units as a function of the range of values measured.
Results of analyses using visual comparisons should be reported in
Jackson turbidity units (JTU). APHA Method 214B describes the nearest
reporting units as a function of the range of values measured. The method
used to determine each JTU value (i.e., candle turbidimeter or bottle standards)
should be identified.
32
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Effluent
Turbidity
Results of all turbidity determinations should be reported, including
QA replicates. Any factors that may have influenced sample quality should
also be reported.
33
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Effluent
Total Suspended Solids
Total Suspended Solids
Field Procedures--
Col lection--Samples should be collected in glass or plastic bottles.
Nonrepresentative particulates, such as leaves, sticks, or rocks, should
be noted and excluded from the sample analysis.
Processing—Total suspended solids samples cannot be preserved adequc. ?ly
and should therefore be analyzed as soon as possible aftei col lection (A.'HA
1985). If a delay occurs, samples should be held at 40 C to minimize micro-
biological decomposition of solids. The length of delay should not exceed
7 days and should be noted on the log sheet.
Laboratory Procedures—
Analytical Procedures — Suspended solids determination should be made
according to procedures described in U.S. EPA Method 160.2 or APHA Method
209C. The drying temperature of the filtered residue can influence results
because temperature and length of heating affect weight losses due to volatili-
zation of organic matter, mechanically occluded water, water of crystallization,
and gasses from heat-induced chemical decomposition, as well as weight
gains due to oxidation (APHA 1985). Thus, drying temperature must be carefully
controlled and not allowed to deviate from the recommended range of 103-1050 C.
To avoid contamination, filters should be handled using forceps during
all steps from initial to final weight determinations. Filters should
be washed with distilled water and dried prior to initial weight determination.
Because glass-fiber filters are hygroscopic, they must be stored in a desiccator
when cooling. When filtering the samples, it is critical that the filter
is seated tightly on the surface of the filtration apparatus and that all
holes in the crucible are covered. To ensure complete removal of salts
after filtering the sample, the filter should be rinsed with a minimum
of three successive 20-mL portions of distilled water. It is recommended
34
-------
Effluent
Total Suspended Solids
that enough water be filtered to ensure that at least 5 mg of residue Is
collected. Because excessive residue on the filter may form a water-entrapping
crust, the sample size should be limited to that which yields less than
200 mg of residue (APHA 1985). Prolonged filtration times resulting from
filter clogging may produce high results due to'excessive solids capture
on the clogged filter. Therefore, filtering should be terminated before
any evidence of clogging is noted.
jlaMbration and Preventive Maintenance—The analytical balance should
be calib ated weekly using standard weights, according to the manufacturer's
instructions. It is recommended that the balance have a minimum accuracy
of 0.1 mg. The manufacturer's preventive maintenance procedures should
be followed carefully.
Quality Control Checks--For each weight determination, filters should
be run repeatedly through the drying/cooling cycle until the weight loss
is less than 4 percent of the previous weight or 0.5 mg, whichever is less
(APHA 1985). Duplicate analyses should be conducted on at least 10 percent
of the total number of samples. U.S. EPA reference samples should be analyzed
quarterly to check the overall accuracy of the method.
Corrective Action—If the analytical balance will not produce repeatable
measurements at 0.1 mg, the manufacturer's troubleshooting guide should
be consulted. If the filter becomes clogged during filtration, it should
be discarded and the analysis should be repeated using a clean filter.
To prevent clogging of the second filter, the volume of sample analyzed
should be reduced.
A filter blank should be carried through the preparation, drying,
and desiccation steps to monitor any changes in filter tare weight. If
the filter weight is not consistent after drying and cooling, the desiccant
should be checked. A color-indicating desiccant is recommended, so that
spent desiccant is easily detected. Also, the seal on the desiccator should
be checked and, if necessary, ground glass rims should be greased or "0"
35
-------
Effluent
Total Suspended Solids
rings replaced. Cooling should be closely timed, so that weighing times
are consistent between batches of samples, thereby minimizing the need
for multiple weighings.
Data Quality and Reporting—Precision of results varies directly with
the concentration of suspended matter and, at low levels, the ratio of
the weight of the suspended matter to the weight of the filter. There
are no procedures for determining the accuracy of field measurements of
suspended matter. Total suspended solids measure'. Tits should be reported
as mg/L to a minimum of two significant figures. Result., of all determinations
should be reported, including QA replicates. Any factors that may have
influenced sample quality should also be reported.
36
-------
Effluent
Settleable Solids
Settleable Solids
Field Procedures—
Collection—Settleable solids samples should be collected in 1-L glass
or plastic containers. The sample container selected for use should be
checked *.o ensure that material in suspension does not adhere to the container
walls. Nonrepresentative particulates, such as leaves, sticks, or rocks,
should be noted and excluded from sample analysis.
Processing—Settleable solids samples cannot be preserved adequately
and should therefore be analyzed as soon as possible after collection (APHA
1985). If a delay occurs, samples should be held at 40 C to minimize micro-
biological decomposition of solids. The length of delay should not exceed
48 h and should be noted on the log sheet.
Laboratory Procedures—
Analytical Procedures—Procedures used to determine settleable solids
concentrations are presented in U.S. EPA Method 160.5 and in APHA Method
209E. The sample should be well-mixed before introduction to the Imhoff
cone. It is critical that floating material is not included with settleable
material.
Calibration and Preventive Maintenance—Not applicable.
Quality Control Checks—Duplicate analyses should be conducted on
at least 10 percent of the total number of samples.
Corrective Actions—If suspended material is found to adhere to the
sides of the sample collection containers, a different type of container
should be used.
37
-------
Effluent
Settleable Solids
Data Quality and Reporting--The practical lower limit of measurement
defends on sample composition and generally is in the range of 0.1 to 1.0 mL/L
(APHA 1985). Precision and accuracy data are not available at present.
Settleable solids concentrations should be reported as mL/L to a minimum
of two significant figures. Results should be reported for all determinations,
including QA replicates. Any factors that may have influenced sample quality
sr.ould also be reported.
38
-------
Effluent
Floating Particulates
Floating Particulates
Field Procedures--
Col 1ection--A minimum of 5 L of sample should be collected in a glass
or plastic container. The container should be rinsed thoroughly with sample
water before sample collection, as no preservation techniques are available
for this variable.
Processing—For comparable results, samp >s must be treated uniformly
throughout sampling and handling. Analysis should be performed as soon
as possible after sample collection, as no preservation techniques are
available for this variable.
Laboratory Procedures--
Analytical Procedures--Floating particulates should be analyzed in
accordance with APHA Method 206A. At present, this method is tentatively
recommended by APHA. Because even slight differences 'in sampling and handling
can give large differences in measureable floating particulates, all samples
should be treated uniformly, preferably by adequately mixing them before
flotation. When mixing the sample, care should be taken to avoid extensive
air entrapment through formation of a large vortex. Because temperature
variations can affect results, all tests should be conducted at a constant
temperature, preferably 200 c.
Calibration and Preventive Maintenance—No calibration procedures
apply to the recommended method. To prevent oil and grease from sticking
to the analytical equipment, all internal surfaces should be coated with
TFE.
Quality Control Checks — It is recommended that duplicate analyses
be conducted on a minimum of 5 'percent of the total number of samples,
with an additional 5 percent of the samples checked for recovery.
39
-------
Effluent
Floating Particulates
Corrective Actions--If oil and grease appear to be sticking to surfaces,
7FE coatings should be renewed. If recovery drops below 90 percent, samples
snould be analyzed again and each step of the analysis carefully scrutinized.
Data Quality and Reporting--The minimum detectable concentration of
f'oating particulates using the recommended method is 1 mg/L. Precision
varies with the concentration cf suspended matter in the sample. A coefficient
of variation > ' 5.7 percent has been achieved using five replicate samples
(APHA 1985). Although there is no completely satisfactory procedure for
determining the accuracy of the method, approximate recovery can be determined
by running a second test for floatables on all water discharged throughout
the analytical procedures, except for the last 10 ml. Typical recoveries
exceed 90 percent (APHA 1985). Concentrations of floating particulates
should be reported in mg/L to the nearest 0.1 unit. Results for all deter-
minations should be reported, including QA replicates and recovery checks.
Any factors that may have influenced sample quality should also be reported.
40
-------
Effluent
Dissolved Oxygen (Winkler)
Dissolved Oxygen (Winkler Method)
Field Procedures—
Col lection—Prior to sample collection, the fixing reagents should
oe prepared and the dispensing apparatus should be filled. The accuracy
of the volumes being dispensed should be checked and no air should be trapped
in the system. It is recommended that 300-mL glass 800 bottles having
ground glass stoppers be used for the Winkler method. Becaus- chlorine
interferes with the Uinkler method, samples should be taken a a point
prior to chlorination.
Grab samples for dissolved oxygen should be collected very carefully
to avoid aeration and they should be preserved as soon as collected. Composite
sampling for dissolved oxygen is not possible. If possible, sample from
a tap by attaching a soft-walled rubber tube to the tap and extend the
tube to the bottom of the sample bottle. The tubing should be flushed
with sample water to remove air bubbles, and the bottle and stopper should
be rinsed at least three times with sample water. The bottle should be
filled slowly until at least half full, and then filled rapidly. At least
one full bottle volume of sample should overflow the bottle before the
tubing is removed, slowly. After the tubing is removed, the stopper should
be carefully put in place with a twisting motion while water is displaced
from the bottle. Once stoppered, the sample should be checked for air
bubbles. If bubbles are present, the sample should be discarded and a
new sample collected. Fixing solutions should be added to each sample
immediately after collection.
Processing—The stopper should be carefully removed from the bottle
without agitating the sample. Each fixing reagent should be added by gently
placing the tip of the pipet slightly below the surface of the sample and
gently pushing the plunger. The plunger should not be released until the
pipet has been removed from the sample. The pipet tip should be rinsed
with distilled water before being returned to the reagent bottle.
41
-------
Effluent
Dissolved Oxygen (Uinkler)
After the fixing reagents have been added, the bottle should be carefully
stoppered without introducing air bubbles. Excess fluid around the outside
of the stopper should be poured off and the sample bottle should be inverted
5-10 times to thoroughly disperse the precipitate. It is also a good practice
to invert bottles several times approximately 20 min after fixation, to
ensure thorough dispersion of the precipitate.
After allowing the precipitate to settle for 10-15 minutes, the :»< ipper
should be removed and sulfuric acid should be added ti the sample i.i the
same manner as the fixing reagents. The stopper should then be replaced
and the bottle Inverted until all of the precipitate has dissolved. If
the precipitate fails to dissolve, it should be allowed to settle again
and additional sulfuric acid should be added to the sample. It is critical
that all of the precipitate be dissolved before samples are stored. Also,
it is critical that samples not be allowed to stand longer than 8 h before
sulfuric acid is added, as erroneous measurements may result.
Preserved dissolved oxygen samples should be stored in the dark at
10-20° C. Samples should be analyzed as soon as possible after collection,
and storage time should not exceed 8 h. The length of storage should be
recorded on the log sheet.
Laboratory Procedures--
Analytical Procedures—The modified Winkler method is described in
detail in U.S. EPA Method 360.2 and in APHA Method 421B.
Calibration and Preventive Maintenance—Methods of standardizing the
thiosulfate solution are presented by U.S. EPA (1979b). It is recommended
that one person perform the standard and sample titrations because of subjec-
tivity in the color of the endpoint.
42
-------
Effluent
Dissolved Oxygen (Ulnkler)
Preventive maintenance is limited to ensuring that reagent dispensing
and titrating equipment is clean and functions properly.
Quality Control Checks—All standard titrations should be dupl icated.
It is recommended that duplicate analyses be conducted on at least 10 percent
of all samples. Reagent blanks should be run whenever a reagent is changed.
Corrective Action—If the results obtained by running duplicate standard
titration of the thiosulfate solution do not agree within HJ.05 tnL, the
titratior-j should be repeated until agreement is achieved. All reagent
dispensers should be checked for bubbles, and the amounts of reagents delivered
should be verified.
Data Quality and Reporting—Using the modified Winkler method, reproduc-
ibility for field samples is approximately 0.2 mg/L of dissolved oxygen
at the 7.5 mg/L level (U.S. EPA 1979b). Duplicate titrations made during
standardization of reagents should agree within ^0.05 ml. With careful
collection and treatment of samples, dissolved oxygen as low as 1 percent
of saturation can be measured. Dissolved oxygen concentrations should
be reported in mg/L to the nearest 0.1 unit. Results should be reported
for all determinations, Including QA replicates and reagent blanks. Any
factors that may have influenced sample quality should also be reported.
43
-------
Effluent
Dissolved Oxygen (Probe)
Dissolved Oxygen (Probe Method)
Field Procedures--
Col 1ection--Grab samples for dissolved oxygen should be collected
very carefully to avoid aeration. Composite sampling for dissolved oxygen
is not possible. If possible, sample from a tap by attaching a soft-walled
rubber tube to the tap and extend the tube to *• -e bottom of the sample
bottle. The tubing should be flushed with sample water to remove air bubbles,
and the bottle and stopper rinsed at least three times with sample water.
The bottle should be filled slowly until at least half full, and then filled
rapidly. At least one full bottle volume of sample should overflow the
bottle before the tubing is removed slowly. After the tubing is removed,
the stopper should be carefully put in place with a twisting motion while
water is displaced from the bottle. Once stoppered, the sample should
be checked for air bubbles. If bubbles are present, the sample should
be discarded and a new sample collected.
Processing—Because no reagents are used to preserve the oxygen samples,
analyses should be conducted immediately after collection. If a delay
occurs, it should be noted on the log sheet.
Laboratory Procedures—
Analytical Procedures—Detailed analytical procedures should be provided
by the manufacturer of the dissolved oxygen meter. General procedures
are listed in U.S. EPA Method 360.1 and APHA Method 421F.
jeveral precautions should be taken when making measurements with
a membrane electrode. First, constant turbulence should be provided by
a stirrer to ensure precise measurements. Second, adequate time should
be allowed for the instrument to warm up before measurements are started,
and, as individual samples are analyzed, the probe should be allowed to
44
-------
Effluent
Dissolved Oxygen (Prooe)
stabilize to sample temperature and dissolved oxygen. Third, reactive
gases, such as chlorine and hydrogen sulfide, pass through the membrane
probes and may interfere with the analysis or desensitize the probe. Finally,
broad variations in the kinds and concentrations of salts in samples can
influence the partial pressure of oxygen in samples and thereby affect
measurement accuracy.
Calibration and Preventive Maintenance—Cal ibration procedures should
follow the instructions given .y the manufacturer of the dissolved oxygen
meter. The meter generally can be calibrated using one of three methods:
Winkler titration, saturated water, or air. The air method is simplest
and quite reliable. Overall error is diminished when the probe and instrument
are calibrated under conditions of temperature and dissolved oxygen that
match those of the samples. Calibration can be disturbed by physical shock,
touching the membrane, or desiccation of the electrolyte.
Preventive maintenance procedures should follow the manufacturer's
reccmnendations. The oxygen probe should always be stored in a humid environ-
ment to prevent drying out and the need to frequently replace membranes.
Quality Control Checks—The instrument should be calibrated at the
beginning of each series of measurements and after each group of 10 successive
samples. Duplicate measurements should be made on at least 10 percent
of the total number of samples.
Corrective Action—If the dissolved oxygen meter does not appear to
be operating correctly, consult the manufacturer's troubleshooting guidelines
for remedial actions.
Data Quality and Reporting—Repeatability of dissolved oxygen measurements
using a membrane electrode should be 0.1 mg/L and accuracy should be *_ 1
percent (U.S. EPA 1979b). Sensitivity of the electronic readout meter
for the output from the dissolved oxygen probes should normally be 0.05 mg/L
45
-------
Effluent
Dissolved Oxygen (Probe)
(U.S. EHA 1979b). Dissolved oxygen concentrations should be reported in
mg/L to the nearest 0.1 umt. Results should be reported for all deter-
minations, including QA replicates. Any factors that may have influenced
sample quality should also be reported.
46
-------
Effluent
BOO
Biochemical Oxygen Demand (BOD)
Field Procedures--
Collect ion--BOD samples can be collected in glass or plastic containers.
Sample containers and caps should be rinsed thoroughly with sample water
before sample collection.
Processing--BOD samples should be analyzed • mediately after collection.
If a delay occurs, samples should he refrigerated at 40 C to minimize reduction
of BOD. Samples should not be stored for more than 48 h and the length
of storage should be recorded on the log sheet. Refrigerated samples should
be warmed to 200 C prior to analysis.
Laboratory Procedures--
Analytical Procedures—BOD concentrations should be determined according
to U.S. EPA Method 405.1 or APHA Method 507. Samples having more oxygen-
demanding materials than the amount of oxygen in air-saturated water should
be diluted to balance the oxygen demand and supply. If samples are diluted,
nutrient addition (I.e., nitrogen, phosphorus, trace metals) and pH buffering
of the dilution water are necessary to ensure that the sample is suitable
for bacterial growth. To prevent air from infiltrating the incubation
bottles, a water seal should be used. When samples are incubating, all
light should be excluded to prevent photosynthetic production of oxygen.
Samples containing caustic alkalinity or acidity should be neutralized
to pH 6.5-7.5 usfng sulfuric acid or sodium hydroxide. Samples containing
residual chlorine must be dechlorinated (e.g., using sodium thiosulfate).
Calibration and Preventive Maintenance—Dissolved oxygen concentrations
should be measured on all dilution water blanks and seed controls. The
dissolved oxygen uptake of the dilution water should not exceed 0.2 mg/L.
The dissolved oxygen uptake of seeded dilution water should be between
0.6 and 1.0 mg/L. A glucose-glutamic acid standard check solution should
47
-------
Effluent
BOO
be incubated with each batch of samples. Dissolved o.tygen measurements
should be calibrated according to accepted procedures (e.g., see descriptions
of the Winkler and probe methods in this document).
Quality Control Checks—The dilution water blank provides a quality
control on the dilution water as well as on the cleanliness of analytical
equipment (e.g., incubation bottles). Each sample should be analyzed at
a -ninimum of three different dilutions to ensure that dissolved oxygen
uptake is i the optimal range. Optimal dissolved oxygen uptake is at
least 2 mg/l after the incubation, with a residual dissolved oxygen of
at least 1 mg/l in the sample. Duplicate analyses should be conducted
on at least 10 percent of the total number of samples.
APHA (1985) should be consulted for methods of correcting for the
many kinds of interference that may accompany BOD analyses.
Corrective Action—If the dilution water blanks exceed 0.2 mg/L, clean-
liness of containers and water should be checked. Containers may require
1+1 HC1 rinse after detergent washing to remove any residual organic material.
Containers rinsed with acid should be thoroughly rinsed with distilled
water to prevent any acid carryover.
If a 2-percent dilution of the glucose-glutamic acid standard check
solution is outside the range of 200 +_ 37 mg/L, BOD determinations made
with the seed and dilution water should be rejected. The problem, which
could arise from numerous sources, may be identified by running a series
of dilution water blanks using different water sources with and without
seed, preparing a fresh solution of glucose-glutamic acid, changing the
seed, or preparing fresh reagents for the dilution water. The source of
the problem needs to be determined before performing any additional BOD
analyses.
Data Quality and Reporting — Precision data for spiked natural waters
indicate that standard deviations of +_0.7 and +26 mg/L can be achieved
48
-------
Effluent
BOO
for BOD concentrations of 2.1 and 175 mg/L, respectively (U.S. EPA 1979b).
There is no acceptable method for determining the accuracy of the BOO test.
BOD data should be reported as mg/L to the nearest 0.1 unit. Results for
all determinations should be reported, including QA replicates, dilution
*ater blanks, and glucose-glutamic acid standards. Any factors that may
have influenced sample quality should also be reported.
49
-------
Effluent
Total Chlorine Residual
Total Chlorine Residual
Field Procedures--
Col lection—Chlorine residual samples can be collected in glass or
plastic containers. During collection, samples should be protected from
strong light and excessive agitation.
Process ing--Because chlorine in aqueous solution is not stable. cMorine
content of samples will decrease rapidly after collection, part cularly
for weak solutions. In addition, both strong light (e.g., sunlight) and
agitation will accelerate the reduction of chlorine. There is no preservation
technique available. Chlorine-residual samples should therefore be analyzed
immediately after collection and protected from strong light and agitation
before and during the analysis.
Laboratory Procedures--
Analytical Procedures--Chlorine residual should be determined using
U.S. EPA Methods 330.1, 330.2, 330.3, 330.4, and 330.5 or APHA Methods
408A, 408B, 408C, 4080, and 408E. The methods should be consulted to determine
which are most appropriate for available equipment, expected effluent constit-
uents, and expected levels of precision and accuracy. The amperometric
method (i.e., U.S. EPA Method 330.1, APHA Method 408C) is the preferred
method because it Is not subject to interference from color, turbidity,
iron, manganese, or nitrite nitrogen (APHA 1985). This method is not as
simple as the colorimetric techniques and requires greater operator skill
to obtain the best results. However, sample color and turbidity may ir'erfere
with all colorimetric procedures. Because all methods of chlorine-residual
determination depend on the stoichiometric production of iodine, effluent
containing iodine-reducing substances may not be analyzed accurately by
any of these methods, especially where iodine remains in solution.
50
-------
Effluent
Total Chlorine Residual
Calibration and Preventive Maintenance—The methods should be consulted
for procedures related to preparation of standards and cal'oration.
Quality Control Checks — For all colorimetric procedures, a minimum
of one color and one turbidity blank should be run per batch to evaluate
these potential sources of interference. For all techniques, duplicate
analyses should be conducted on a minimum of 10 percent of the total njmber
of samples.
Corrective Action—There are many sources of •' terference with chlorine
residual tests. If a particular source of interference cannot be adequately
compensated for using one method, an alternate method should be used.
Data Quality and Reporting—The detection and accurate quantification
of total chlorine residual in effluent is routinely attainable, although
method detection limits, accuracy, and precision can vary depending on
the method used. The method should therefore be consulted to determine
expected detection limits, accuracy, and precision. Total chlorine residual
concentrations should be reported as mg/L to the nearest 0.01 unit. Results
should be reported for all determinations, including QA replicates and
color and turbidity blanks. Any factors that may have influenced sample
quality should also be reported.
51
-------
Effluent
Oil and Grease
Oil and Grease
Field Procedures—
Collection--Samples should be collected in glass bottles. Bottles
should first be washed with a warm aqueous detergent mixture, and then,
in sequence, thoroughly rinsed with hot tap water, rinsed at least twice
with distilled water, rinsed with 1,1,2-trichloro-l,2,2-trifluoroethane
(i.e., Freon or equivalent), and dried in a -.lean oven at >_105o C for 30
min. Bottle caps should be lined with TFE-coated cardboard inserts or aluminum
foil. Plastic containers are not acceptable.
Only grab samples, and not composite samples, should be collected
to limit loss of oil and grease on sampling equipment. The entire grab
sample should be submitted for analysis. To obtain a daily average concen-
tration, grab samples collected at prescribed time intervals can be submitted
for analysis. Headspace should be left in each sample container for addition
of acid and mixing.
Processing—Acidify the sample in the collection bottle to a pH <2
using sulfuric or hydrochloric acid. Samples should be stored in the dark
at 40 c. Recommended maximum holding time is 28 days (U.S. EPA 1984).
Laboratory Procedures—
Analytical Procedures—Oil and grease should be analyzed using procedures
described in U.S. EPA Method 413.1 or APHA Method 503A. Because asphaltic
materials are insoluble in Freon, the recommended method will give low
recoveries for samples containing such material. U.S. EPA Method 413.1
(gravimetric oil and grease) measures relatively nonvolatile hydrocarbons,
vegetable oils, animal fats, waxes, and soaps. The method is often used
for wastewater analyses since it requires minimal instrumentation and cali-
bration. Sulfur causes interference for the gravimetric method because
it will be extracted and included as oil and grease. Light hydrocarbons
52
-------
Effluent
Oil and Grease
that volatilize at temperatures below 700 c (e.g., gasoline through No.2
fuel oil) are lost during the solvent removal step.
Calibration and Preventive Maintenance—For gravimetric oil and grease
analyses, check accuracy of the analytical balance periodically (minimum
of once per week recommended) using Class S weights. A service contract
that includes scheduled preventive maintenance at least once per year is
recommended. Scratched, chipped, or cracked boiling flasks should be replaced.
For infrared oil and grease analys.', follow the manufacturer's preventive
maintenance procedures for the i.ifrared spectrophotometer. Cells used
for analysis should be checked for scratches each time they are used.
Scratched cells should not be used.
.Quality Control Checks—Duplicate samples should be collected and
performed at a minimum of every 10 samples to establish an estimate of
precision. Because samples should not be split after collection, separate
grab samples should be taken for analysis.
Distilled water spiked with a U.S. EPA performance sample should be
extracted and analyzed with every batch of samples to monitor recovery.
Bottles should be checked for cleanliness by analyzing distilled water
that has been acidified in a sample bottle. A solvent blank should accompany
each batch of samples.
Corrective Action—If oil and grease concentrations in procedural
blanks are greater than the detection limit, check the cleanliness of all
glassware. Always use separatory funnels with TFE stopcocks to avoid contam-
ination from stopcock grease. For gravimetric analyses, high results will
be obtained if any Freon or fumes remain in the flask after distillation.
If difficulty occurs with emulsions during solvent extraction, follow
procedures described in the methods. If the emulsion still fails to dissipate
53
-------
Effluent
Oil and Grease
after addition of salts, gently turn the separatory funnel to the horizontal
position and slowly rotate. Be careful to keep the stopper securely in
place.
If precision or recovery from spike results are poor, check adequacy
of extraction by increasing shaking time.
Data Quality and Reporting—The definition of oil and grease is based
on the procedures used, u less identical procedures are used, oil and
grease deter,.,inat-ions are ,iot intercomparable. Therefore, the method used
for analysis should always be specified.
Inter laboratory recovery results were 102+37 percent at 6.0 mg/L and
97+35 percent at 18.0 mg/L for the gravimetric method (U.S. EPA 1983a).
Measurements should be reported to a minimum of two significant figures
in mg/L. Detection limits are in the range of 5 mg/L for gravimetric oil
and grease determinations (U.S. EPA Method 413.1). Results should be reported
for all determinations, including QA replicates, spiked samples, and blanks.
Any factors that may have Influenced sample quality should also be reported.
54
-------
Effluent
Nitrogen (Ammonia)
Nitrogen (Ammonia)
Field Procedures--
Conection--Ammonia samples can be collected in glass or plastic bottles.
Each bottle and cap should be rinsed thoroughly with sample water prior
to sample collection.
Processing--Resu1 ts of ammonia analyses are most ^liable when they
are made on fresh samples. However, if analysis must be relayed, samples
can be stored for up to 28 days by acidification to pH<2 with sulfuric
acid and refrigeration at 40 c. The length of delay before analysis should
be recorded on the log sheet.
Laboratory Procedures--
Analytical Procedures—Manual distillation of effluent samples prior
to ammonia determinations is required by the U.S. EPA, unless data on file
demonstrate that distillation is not required. Distillation is recommended
due to the sensitivity of ammonia procedures to color and possible interferences
in the effluent. Following distillation at pH 9.5, ammonia concentration
can be determined by U.S. EPA Methods 350.1, 350.2, or 350.3 or APHA Methods
417A, 417B, 4170, or 417G. The methods should be consulted to determine
which is most appropriate for available equipment, expected concentrations,
and expected levels of precision and accuracy.
Calibration and Preventive Maintenance--Calibration procedures should
follow those specified in the method. If samples are being distilled,
standards should also be distilled prior to analysis to check for ammonia
contamination or loss during processing. Concentrations of the calibration
standards should bracket the sample concentrations. If a sample concentration
is outside the range of calibration, then an additional calibration standard
should be analyzed to check if the result is within the linear range of
55
-------
Effluent
Nitrogen (Ammonia)
the method. Alternatively, tne sample should be diluted to within the
calibration range and then reanalyzed.
Quality Control Checks—Duo! icate analyses snould be conducted on
a minimum of 5 percent of the total number of samples, with an additional
5 percent of the samples spiked and analyzed -or percent recovery. A blank
should be analyzed with each batch of samjles. A U.S. EPA performance
sample should be analyzed at 1 :ast once per quarter.
Corrective Act Ion--Contamination of ammonia samples .-an occur easily
due to the volatile nature of anmorna. To prevent possible cross-contamination,
reagents used for other analyses that contain ammonia (e.g., colorimetric
phenol) should be isolated from samples and standards used for ammonia
determinations. In addition, cleaning preparations that contain significant
quantities of ammonia (e.g., Pinesol, wax removers) should not be used
in the laboratory area where ammonia determinations are performed.
Contaminated glassware should be rinsed with 1+1 HC1 and then with
distilled water. To check for contamination, blanks should be analyzed
whenever a new reagent is prepared.
Data Quality and Reporting—Detection and accurate quantification
of ammonia in effluent is routinely attainable, although method detection
limits can vary widely because of methods or instrumentation. The analytical
method should be consulted to determine expected detection limits, precision,
and accuracy. Data should be reported in mg/L as N to a maximum of three
significant figures. Results should be reported for all determinations,
including QA replicates and spiked samples. Any factors that may have
influenced sample quality should also be recorded.
56
-------
Effluent
Nitrogen (Total Kjeldahl)
Nitrogen (Total Kjeldahl)
Field Procedures--
Co11ection--Kjeldahl nitrogen samples can be collected in glass or
plastic containers. Caps should be unlined, as paper liners and/or glue
may interfere with the analysis. Each container and cap should be rinsed
thoroughly with sample water prior to sample collection.
Processing—If possible, samples should be analyzed immediately after
collection. If Immediate analysis is not possible, samples can be stored
up to 28 days by acidification to pH<2 with sulfuric acid and refrigeration
at 40 c. The length of delay before analysis should be recorded on the
log sheet.
^
Laboratory Procedures--
Analytical Procedures—Approved test procedures for the analysis of
total Kjeldahl nitrogen include U.S. EPA Methods 351.1, 351.2, 351.3, or
351.4 and APHA Methods 417B, 4170, 417E, 420A, or 420B. The methods should
be consulted to determine which is most appropriate for available equipment,
expected concentrations, and expected levels of precision and accuracy.
Calibration and Preventive Maintenance--Calibration procedures should
follow those specified In the method. If samples are being digested and
distilled, standards should also be digested and distilled prior to analysis
to check for amnonia contamination or loss during distillation. Concentrations
in the calibration standards should bracket the sample --oncentrations.
If a sample concentration is outside of the range of calibration, then
an additional calibration standard should be analyzed to check if the result
is within the linear range of the method. Alternatively, the sample should
be diluted to within the calibration range and then reanalyzed.
57
-------
Effluent
Nitrogen (Total Kjeldahl)
Quality Control Checks--Duplicate analyses should be conducted on
a minimum of 5 percent of the total number of samples, with an additional
5 percent of the samples spiked and analyzed for percent recovery. A blank
should be analyzed with each batch of samples. A U.S. EPA performance
sample should be analyzed at least once per quarter.
Corrective Action—Because ammonia is a component of Kjeldahl nitrogen,
precautions against contamination chat were described for ammonia analyses
c ould be followed.
Data Quality and Reporting—The detection and accurate quantification
of Kjeldahl nitrogen in effluent is routinely attainable, although method
detection limits can vary widely because of methods or instrumentation.
The analytical method should be consulted to determine expected detection
limits, precision, and accuracy. Data should be reported in mg/L as N to
a maximum of three significant figures. Results should be reported for
all determinations, including QA replicates, blanks, and spiked samples.
Any factors that may have influenced sample quality should also be recorded.
58
-------
Effluent
Nitrogen (Nitrate and Nitrite)
Nitrogen (Nitra".e and Nitrite)
Field Procedures—
CpJJ_ec_tioin--Nitrate-nitrite samples can be collected in glass or plastic
containers. Prior to sample collection, each container and cap should
be rinsed thoroughly with sample water.
Processing--Nitr--, e-nurite samples should be analyzed immediately
after collection. If a Jelay occurs, samples can be stored for up to 24 h
by acidification to pH<2 with sulfuric acid and refrigeration at 40 C.
Samples must not be preserved using mercuric chloride because the mercuric
ion accelerates the degradation of the cadmium-reduction column (APHA 1985).
Laboratory Procedures—
Analytical Procedures—Approved test procedures for the analysis of
nitrate-nitrite include U.S. EPA Methods 353.1, 353.2, and 353.3 and APHA
Methods 418C and 418F. The methods should be consulted to determine which
is most appropriate for available equipment, expected concentrations, and
desired levels of precision and accuracy.
Calibration and Preventive Maintenance—Calibration procedures should
follow those specified In the method. Efficiency of each reduction column
should be checked by comparing a nitrite standard to a reduced nitrate
standard at the same concentration. This efficiency check should be made
a I the beginning and the end of each sample run and at a minimum frequency
of every 10 samples. Reactivate the copper-cadmium granules when reduction
falls below 75 percent.
Concentrations of the calibration standards should bracket the sample
concentrations. If a sample concentration is outside the range of calibration,
then an additional calibration standard should be analyzed to check if
59
-------
Effluent
Nitrogen (Nitrate and Nitrite)
tne result is within the linear range of the method. Alternatively, fhe
sample should be diluted to within the calibration range and reanalyzed.
Quality Control Checks--Duplicate analyses should be conducted on
a minimum of 5 percent of the total number of samples, with an additional
5 percent of the samples spiked f. analyze for percent recovery. A blank
-.-at nas been run through the reduction column should be analyzed with
each batch of samples. A U.S. EPA performance sample should be analyzed
at least once per quarter.
Corrective Action—Various components of the effluent can interfere
with the analysis. The method should be reviewed for ways to remove possible
interferences prior to analysis. Possible intereferences include suspended
solids, residual chlorine, oil and grease, and high concentrations of iron,
copper, or other metals.
The area where nitrate-nitrite analyses are performed should be well
isolated from exposure to nitric acid or nitric acid fumes.
Data Quality and Reporting—The detection and accurate quantification
of nitrate-nitrite in effluent are routinely attainable, although method
detection limits can vary because of methods or instrumentation. The analytical
method should be consulted to determine expected detection limits, precision,
and accuracy. Data should be reported in mg/L as N to a maximum of three
significant figures. Results of all determinations should be reported,
including QA replicates, blanks, and spiked samples. Any factors that
may have influenced sample quality should also be reported.
60
-------
Effluent
Phosphorus (Total)
Phcsphorus (Total)
Field Procedures--
Collection--Phosphorus samples may be collected in glass or plastic
containers. Containers should be rinsed with IN HC1 followed by several
r-.nses with distilled water. Detergents containing phosphate should never
be used on containers or labware that is to be used for phosphate analysis.
Sample containers and lids should be ri sed thoroughly with sample water
before sample collection.
Processing—Phosphorus samples can be stored up to 28 days before
analysis by acidification to pH<2 with sulfuric acid and refrigeration
at 40 c. Samples with low concentrations of phosphorus should not be stored
in plastic containers, as phosphates may adsorb onto the container walls.
Laboratory Procedures—
Analytical Procedures—Approved test procedures for the analysis of
total phosphorus in effluent are U.S. EPA Methods 365.1, 365.2, 365.3 and
365.4 and APHA Methods 424C, 424F, and 424G. The methods should be consulted
to determine which is most appropriate for available equipment, expected
concentrations, and desired levels of precision and accuracy.
Calibration and Preventive Maintenance—Cal ibration procedures should
follow those specified 1n the method. Concentrations of the calibration
standards should bracket the sample concentration. If a sample concentration
is outside the range of calibration, then an additional calibration standard
should be analyzed to check if the result is within the linear range of
the method. Alternatively, the sample should be diluted to within the
calibration range and reanalyzed.
Quality Control Checks—Duplicate analyses should be conducted on
a minimum of 5 percent of the total number of samples, with an additional
61
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Effluent
Phosphorus (Total)
5 percent of the samples spiked to analyze lor percent recovery. A blank
snould be analyzed with each batch of samples. A U.S. EPA performance
sample should be analyzed at least once per quarter.
Corrective Action—Because phosphorus contamination can occur from
a variety of sources, it is recommended that a clearly marked set of lab-
v«are be dedicated to only phosphorus analysis. This laoware should never
be exposed to phosphorus detergents or reagents containing phosphate.
Various components of the effluent can interfere wuh the analysis.
The method should be reviewed for ways to remove interferences or adjust
for interferences from components that cannot be removed. Silica and arsenic
are possible positive interferences, while hexavalent chromium and nitrite
can cause low recovery.
For highly colored or turbid samples, additional sample preparation
(e.g., further oxidation or filtration) may be required prior to color
aevelopment. In any case, blanks should be prepared by adding all the
reagents except the coloring reagents to the sample. Measure absorbance
in the sample blank at the wavelength used for the phosphorus determination
and subtract this absorbance value from the sample absorbance prior to
calculation of phosphorus concentration.
Data Quality and Reporting—Detection and accurate quantification
of total phosphorus in effluent is routinely attainable. Actual method
detection limits can vary because of methods or instrumentation. The analytical
method should be consulted to determine expected detection limits, precision,
and accuracy. Data should be reported in mg/L as P to a maximum of three
significant figures. Results of all determinations should be reported,
including QA replicates and spiked samples. Any factors that may have
influenced sample quality should also be reported.
62
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Effluent
Priority Pollutant Metals
P-iority Pollutant Metals
Field Procedures--
Col lection--The best containers for collection of trace metal samples
are made of quartz or 7FE. Because these containers are expensive, the
preferred sample container is made of polypropylene or linear polyethylene
v»ith a polyethylene cep (APHA 1985). Boros il icate glass containers may
be used, but do not use soft glass containers or container* .nth aluminum
or cardboard lined lids.
A minimum sample size of 100 mL is required for the analysis of all
priority pollutant metals, except mercury, which requires an additional
100 ml. To allow for duplicates, spikes, and potential reanalysis, a sample
size of 0.5-1 L is recommended for the entire list of priority pollutant
metals.
Possible problems during sample collection involve contamination from
the sampling device, airborne dust, cross-contamination from previous samples,
and loss of metals by adsorption on container walls or precipitation after
collection. Contamination can be minimized by avoiding the use of metal
when collecting effluent samples. Automatic sampling devices should be
free of metal parts in contact with the sample. Prior to use, containers
should be thoroughly cleaned with detergent solution, rinsed with tap water,
soaked in acid, and then rinsed with metal- free water. Any glass or plastic
parts associated with the sampling equipment should be cleaned in the same
manner. For quartz, TFE, or glass containers, use 1*1 HN03, 1+1 HC1, or
aqua regia (3 parts cone HC1 + 1 part cone HN03) for soaking. For plastic
material, use 1+1 HN03 or 1+1 HC1. Reliable soaking conditions are 24 h
at 700 c (APHA 1985). Oo not use chromic acid when preparing containers.
For metal parts, clean as stated for glass or plastic, except omit the
acid soak step of the cleaning procedure. All acids should be at leas*
reagent grade.
63
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Effluent
Priority Pollutant Metals
Processing—Acidify samples to pH<2 with nitric acid immediately after
sampling. Usually 1.5 ml concentrated HN03/L sample is sufficient. Check
that adequate acid has been added by testing an aliquot using pH paper
or a pH meter. (Do not insert any test materials into the sample container.)
Use high-purity acid for preservation.
Recommended maximum holding t ne for trace metal samples is 6 mo,
except for mercury samples, whicn should be held a maximum of 28 days prior
to analysis (U.S. EPA 1982).
Laboratory Procedures--
Analytical Procedures—Priority pollutant metals should be analyzed
according to the methods identified in Table IB (see Appendix A) of U.S. EPA
(1984). Prior to the analysis, the effluent sample must be digested using
tne acids specified in the procedure. The digestate can then be analyzed
by flame Atomic Absorption Spectrophotometry (AAS), graphite furnace AAS,
or Inductively Coupled Plasma (ICP), depending on the sample concentration
and required detection limit. Mercury analyses must be performed on a
separate sample aliquot by cold vapor AAS.
ICP can be used to screen samples for elements that are present in
relatively high concentrations or for those that may require more sensitive
analysis by graphite furnace AAS. Analysis by ICP can be subject to inter-
element interferences, while graphite furnace AAS can be subject to matrix
problems from acid or salt content of the samples. The detection limit
of the selected method should be adequate to allow determination of compliance
with water quality criteria.
Calibration and Preventive Maintenance—In general, all instruments
must be calibrated daily and each time the instrument is set up. Calibration
procedures should follow those for the specified method for each analysis.
Calibration standards must be prepared using the same concentrations of
acids as will result in the samples following sample preparation.
64
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Eff uent
Priority Pollutant Metals
After an instrument has been calibrated, the accuracy of the initial
calibration should be verified by the analysis of certified control solutions
at a frequency of once every 10 samples or every 2 h during an analysis
run, whichever is more frequent, and after the last analytical sample.
If a certified control solution is not available, then a standard should
be used. Analyte for this standard should be from a different source than
tnat for tne initial calibration. If the deviation of the continuing cal i-
bratio verification is greater than the calibration control limits specified
in the method, then the instrument must be recalibrated, and the preceding
10 samples reanalyzed.
All equipment should have scheduled routine preventive maintenance,
and a record of all maintenance should be kept in a logbook. Critical
spare parts should be kept on hand.
Quality Control Checks—To establish an estimate of precision, a minimum
of 5 percent of the total number of samples should be analyzed in duplicate
or one duplicate for each survey, whichever is more frequent. When more
tnan 20 samples are to be analyzed for one survey, the project manager
may choose to implement a program of triplicate analyses. The overall
percentage of replicates should be at least 5 percent. To establish an
estimate of recovery, samples spiked before digestion should be analyzed
at the same frequency as replicates. Spike concentrations to be added
should be approximately equal to the concentration found in the unspiked
sample. An acceptable range of spike concentrations is 0.5-5 times the
sample concentration.
A method blank should be carried through all digestion and analysis
steps at a minimum frequency of every 20 samples, or for each batch of
samples analyzed, whichever is more frequent. If the concentration of
the blank is less than the required detection limit, then no correction
of sample results is performed. If the blank contamination is extensive,
then the batch of samples associated with the blank should be reanalyzed.
65
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Effluent
Priority Pollutant Metals
In general, the blank should be 10 times less than the concentration of
the analyte in the sample. The data should be corrected by data users
for the blank values between the required detection limit and the control
1imit.
For ICP analysis, additional QC checks should include an interference
cneck sample to ;enfy interelement and background correction factors.
For graphite furnace AAS, additional QC checks should include duplicate
injections with the mean value repo. ed. Relative standard deviation of
the readings should be within control limits, or the sample must be rerun
at least once.
Corrective Action—If the concentration of the field or method blank
is greater than the required detection limit, then all steps in the sample
handling should be reviewed. Many trace metal contamination problems are
due to airborne dust. Keeping containers closed and rinsing all handling
equipment immediately prior to use minimizes dust problems.
Poor duplication may be caused by inadequate mixing of the sample
before removal of aliquot, inconsistent contamination, inconsistent digestion
procedures, or instrumentation problems.
Poor spike recovery may be caused for the same reasons as poor du-
plication. However, if duplicate results are acceptable, then poor spike
recovery may be caused by loss of analyte during digestion or by sample
matrix interferences during analysis. To check analyte loss during digestion
or for low recovery due to interferences during analysis, spike the sample
after digestion and compare the analysis to the predigestion spike. If
the results are different, then the digestion technique should be adjusted.
If the results are the same then dilute the sample by at least a factor
of five and reanalyze. If spike recovery is still poor, then standard
additions, matrix modifiers, or another method is required.
66
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Effluent
Priority Pollutant Metals
Data Quality and Reporting—Data should be reported to a maximum of
three significant figures as mg/L. These units are consistent with ODES
format. Detection limits can vary widely because of methods or instrumen-
tation. The analytical method should be consulted to determine expected
detection limits, precision, and accuracy. The data report should include
duplicate, spike, and blank results. The data should not be blank-corrected.
It is recommended that all new data included in the ODES database be blank-
corrected by the data users. The data nummary from the laboratory should
also include:
• Digestion procedures
t Instrument detection limits
• Detection limit for each element
• Blank associated with sample
• Deviations from the prescribed method
t Problems associated with analysis.
For a more thorough QA review, additional documentation (e.g., calibration
curves) may be requested.
67
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Effluent
Priority Pollutant Organic Compounds
Priority Pollutant Organic Compounds
Field Procedures--
Col lection--The priority pollutant organic compounds can be separated
into purgeable (volatile) and extractable (base/neutrals, acids, pesticides,
ani PCBs) compounds. Container preparation and collection techniques differ
fcr these two Croups.
If a volatile analysis is required, two separate 40-mL glass containers
should be filled leaving no headspace. The container, screw cap, and septum
should be washed with detergent, rinsed once with tap water, rinsed once
with distilled water, and dried at >105o c. Use of a solvent rinse will
interfere with the analysis. To obtain a sample with no headspace, fill
the vial to overflowing so that a convex meniscus forms at the top. With
the cap liner's TFE side down, place the cap carefully on the opening of
the vial, displacing the excess water. Once sealed, invert the bottle
to verify the seal by demonstrating the absence of air bubbles. Samples
for volatile analyses should be grabs only, because many of the volatile
compounds of interest may be lost while compositing.
For the extractable analyses, amber glass containers with TFE-lined
screw caps should be used. Clear glass may be used if the sample is kept
from light (i.e., stored in an ice chest or refrigerator). It is advised
that 4 L be collected to allow for reanalysis if necessary. Eight L should
be collected for samples to be analyzed as spikes or duplicates. The container,
lid, and liner should be washed with detergent, rinsed twice with tap water,
rinsed once with distilled water, and the liner and container rinsed once
with high-purity methylene chloride or acetone. Kiln drying of the container
at 4500 c for at least 1 h can be substituted for solvent rinse. There
is evidence that some priority pollutants (e.g., hexachlorobenzene) in
aqueous solution adsorb strongly to TFE lid liners, ."his may be a potential
problem if the effluent contains ".hese pollutants.
68
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Effluent
Priority Pollutant Organic Compounds
Automatic sampling equipment should be clean, and free of plastic
ana other sources of contamination. Ideally, a sample bottle containing
"organic-free water" carried through all processing and handling will serve
as a field blank. Because organic-free water is not readily available
outside of a laboratory, this may be impractical for all monitoring programs.
Alternatively, an empty sample jar carried through all processing and handling
will serve as a field blank. A solvent rinse of the bottle should be analyzed.
The solvent should be the same as that used for sample extraction. This
will serve as a check on contamination tha' may occur curing shipping and
storage.
Process ing--Samples should be stored in the dark at 40 c or on ice.
Freezing of the samples in transit should be avoided as the water expansion
causes broken vials. For volatile organic compounds, maximum recommended
holding time is 7 days. For extractable organic compounds, maximum recommended
holding time is 7 days prior to extraction, with all analyses completed
within 40 days of extraction.
Sample extracts should be stored frozen at <-10o c.
Laboratory Procedures--
Analytical Procedures--Priority pollutant organic compounds should
be analyzed according to the methods identified in Tables 1C and 10 (see
Appendix B) of U.S. EPA (1984). If samples with analyte levels that differ
by several orders of magnitude as determined by screening or experience
are being analyzed together, carryover can be a problem. This should be
kept in mind when reviewing data obtained by auto-injection. For volatile
organic compounds, samples that appear "clean" should be analyzed first.
Analysis of an appropriate solvent after a high-level sample can minimize
the chance of false positives. If separate injection syringes are used
for standards and samples, any bias between syringes should be accounted
for. They should be cleaned thoroughly between each use to avoid cross-
contamination.
69
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Effluent
Priority Pollutant Organic Compounds
Detection limits vary with specific compounds and methods but are
usually obtainable in the range of 5-10 ppb for base, neutral, and acid
compounds (U.S. EPA Method 625), 0.005-0.10 ppb for pesticide/PCB analysis
(U.S. EPA Method 608), and 1-10 ppb for volatiles (U.S. EPA Method 624).
Sample size, final volume, co-extractives and nature of effluent can all
affect the actual detection limit. Detection limits , equested by project
managers should take into account obtainable levels as well as required
301(hj program criteria.
Calibration and Preventive Maintenance—Before beginning analysis
of samples, a calibration curve bracketing the working range must be performed.
This calibration should be repeated after all major equipment disruptions.
Calibration checks of the 6C/MS system should be done at the beginning
and end of each day and at least every 12 h to verify that the instrument's
response is in control. Specific instrument tuning criteria (e.g., DFTPP,
BFB) are provided in each method (U.S. EPA 1984). Calibration of the GC/ECO
system should be done at the beginning of each day and verified at least
every 6 h. These are only recommended minimum frequencies. Depending
on the nature of the samples, it may be necessary to verify calibration
more frequently.
A routine QC check on each lot of analytical reagent used in extraction
can prevent undetected contamination problems. Each lot of alumina, silica
gel , sodium sulfate, or Florasil used should be monitored as a possible
source of contamination. The efficacy of adsorbants often varies between
lots and should be routinely monitored. Each lot of surrogate mixture
should also be checked for contaminants.
Equipment should be maintained and serviced routinely by experienced
chemists according to manufacturers instructions and good laboratory practices.
Logbook records should document maintenance for each measurement device.
70
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Effluent
Priority Pollutant Organic Compounds
Quality Control Checks--To monitor reproducitnlity and extraction
efficiency, duplicates and matrix spike analyses should be performed.
A frequency of 5 percent each or one each per survey (whichever is greater)
is the recornnended minimum. When more than 20 samples are to be analyzed
for one survey, the project manager may choose to implement a program of
triplicate analyses. The overall percentage of replicates should be at
least 5 percent. Method interferences can be caused by contaminated glassware,
reagents, solvents, or processing hardware. To check for ;ontamination,
one method blank should be processed after each group of Z samples or
with each batch, whichever is greater. Addition of known amounts of surrogate
compounds to each sample before purging or extraction will serve to monitor
preparation and analysis of samples. A spike prepared in the field can
be submitted to the laboratory as another measure of accuracy.
Corrective Action—When results of QC samples fall outside of established
limits, several courses of action are available. Contamination in the
lab reagent blank sample is cause for positive findings of the same compounds
in samples to be suspect. If contamination is extensive, reanalysis of
the entire associated group may be in order. Blank contamination should
be kept to less than 10 percent of sample values and preferably below the
method detection limits. Contamination found in the field blank should
be considered when looking at the associated sample data. Extensive contam-
ination of lab or field blank (>30 percent of sample values) should lead
to a detailed review of laboratory, sampling, transport, and storage pro-
cedures. Phthalates, methylene chloride, and toluene are common laboratory
contaminants that may be detected in blanks above the method detection
limit.
Poor duplication may be caused by inadequate mixing of the sample
before reroving aliquots, inconsistent contamination, inconsistent extraction
procedures, or instrument problems. If further replication of an analysis
produces poor results, a step-by-step examination of the method may be
necessary to determine the reason for the poor results. If the results
71
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Effluent
Priority Pollutant Organic Compounds
of the duplicates fall outside control limits, the laboratory should contact
the data user and discuss the possibility of further analyses.
Poor spike recovery may be caused for the same reasons as poor duplication
or by matrix effects. If the spiked compound is added at a concentration
much less than that found in the sample, recovery may be difficult to deter-
mine. This problem is difficult to avoid, as most environmental samples
cor:ain unknown concentrations of organic compounds. To check for analyte
loss curing processing, a step-by-step examinatior of c.ie method using
a spiked blank is necessary, with measurements of the .nalyte at each step.
Sample results that fall outside the established calibration curve
are suspect until linearity of response can be shown at that concentration,
or the extract diluted appropriately and reanalyzed. Extremely high concen-
trations of organic compounds may saturate the extraction capabilities
of the method and may necessitate re-extraction of a smaller sample size
or use of a more appropriate method.
If the instrument's continuing calibration (single point) falls outside
control limits, no samples should be analyzed until the calibration is
within control limits. The standard should be reinjected to confirm the
problem and to discount the possibility of operator error. If still outside
of control limits, the instrument should be recalibrated (multi-point)
and at least the previous sample reanalyzed and results compared. This
may indicate that reanalysis of all samples since last calibration is unneces-
sary.
Data Quality and Reporting—A data summary for each sample should
be submitted. All data should be reported in ug/L using two significant
figures. Data should not be blank-corrected. Spike recoveries, relative
percent difference between duplicates, and blank results (ng/sample) should
also be submitted. The following information is also needed for each sample
to allow independent QA review:
72
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Effluent
Priority Pollutant Organic Compounds
• Sample volume extracted
• Final volume of extract
• Amount of extract injected
• Instrument detection limits
• Detection limit f:r each compound
• Blank associated with sample
• Deviations from the prescribed method
t Problems associated with analysis.
For a more thorough QA review, additional documentation (e.g., chromatograms,
computer listings) may be requested.
73
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Effluent
Total and Fecal Coliform Bacteria
Total and Fecal Colifortn Bacteria
Field Procedures--
Col 1ection--Samples should be collected in clean, sterile polypropylene
or glass containers. The sample containers must ce resistant to sterilizing
conditions an-- to the solvent action of water. The container lids must
not produce bacteriostatic or nutritive compounds upon sterilization.
The sample containers must sea1 tightly. Containers with chips, cracks,
or etched marks should be discardeo.
Heat-resistant glass or plastic sample containers should be autoclaved
at 1210 c for 15 min. Alternatively, dry glass containers can be sterilized
in a hot-air oven at 1700 c for at least 2 h. For plastic containers that
are not heat-resistant, ethylene oxide gas sterilization is acceptable
(Bordner et al. 1978). Containers sterilized by gas should be stored at
least 12 h before use to ensure all gas has dissipated.
If the sample water has residual chlorine, sodium thiosulfate should
be added to neutralize the chlorine and thereby prevent continued bactericidal
action after sample collection. In this manner, the true microbial content
of the water at the time of sampling can be estimated more accurately.
If sodium thiosulfate must be added to a sample, it should be added to
the sample container prior to sterilization so that the final concentration
in the sample will be 100 mg/L. For a 120-mL container, 0.1 ml of a 10-
percent solution of sodium thiosulfate will neutralize a sample containing
as much as 15 mg/L of residual chlorine (APHA 1985).
If the sample water contains heavy metals in concentrations exceeding
0.01 mg/L, a chelating agent should be added to the sample container to
reduce metal toxicity. This is particularly important if samples are not
analyzed within 4 h after collection. APHA (1985) recommends using the
disodium salt of ethylenediaminetetracetic acid (EOTA), adjusted to pH
74
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Effluent
Total and Fecal Coliform Bacteria
6.5, and added to the sample contaiier before sterilization. For a 120-mL
container, addition of 0.3 ml of a 15-percent EDTA solution is considered
adequate (APHA 1985).
It is critical that samples are not contaminated during the collection
process. To avoid contamination, sterilized containers should be kept
sealed until they are used, containers should be filled without rinsing,
ana container lids should be replaced immediately after the samples have
been collected. When remo- id from containers, lids should be held face
down in one hand and not set «iown on any surface. Adequate headspace (at
least 2.5 cm) should be left in each sample container to facilitate mixing
prior to analysis.
Processing—Samples should be analyzed as soon as possible after col-
lection. If a delay occurs, samples should be held at 40 C for a maximum
of 6 h. The length of delay should be noted on the log sheet.
Laboratory Procedures--
Analytlcal Methods—Details of the membrane filter (MF) method and
tne most probable number (MPN) method are presented in Part III of Bordner
et al. (1978) and in Parts 908 and 909 of APHA (1985). Although the MF
method is more precise than the MPN method, it is also more sensitive to
interference from turbidity in samples. Because the MF technique usually
yields low and variable recovery from chlorinated waters, the MPN technique
should be used for chlorinated effluent.
Calibration and Preventive Maintenance—This information is reviewed
extensively in Part IV of Bordner et al. (1978) and in Part 902 of APHA
(1985).
Quality Control Checks — Qua! ity control checks for col i form analyses
are listed in detail in Part IV of Bordner et al. (1978) and in Part 902
of APHA (1985). The list includes:
75
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Effluent
Total and Fecal Coliform Bacteria
• Sterility checks on media, dilution and rinse water, glassware,
and membrane filters
• Duplicate analyses on 10 percent of samples and on at least
one sample per test run
• Colony verifications on a monthly basis.
Corrective Action—Procedures detailed in the relevant sections o^
Bordner et al. (1978) and APHA (1985) should be followed.
Data Quality and Reporting—Table 909:11 of APHA (1985) presents 95-
percent confidence limits for the MF method for coliform colonies of 1,
2, 3, 4, 5 and 10. The precision of the MPN method increases with increasing
nuntoer of replicates. With five tubes, each with 1 mL of sample, a completely
negative result is expected less than 1 percent of the time (APHA 1985).
Confidence limits (95 percent) for various MPN counts are given in Tables
908:111, 908:IV, and 908:V of APHA (1985).
Using the MF method, data should be reported as densities of col i forms
per 100 ml. Using the MPN method, data should be reported as MPN values
per 100 ml. Results should be reported for all determinations, including
QA replicates, sterility checks, and colony verifications. Any factors
that may have influenced sample quality should also be reported.
76
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Effluent
Enterococcus Bacteria
Eiterococcus Bacteria
Field Procedures--
Collection—Samples should be collected in clean, sterile polypropylene
or glass containers. The sample containers must be resistant to sterilizing
conditions and to the solvent action of water. The container lids must
not produce bacteriostatic or nutritive compounds upon sterilization.
Tne sample containers must seal tightly. Cntainers with chips, cracks,
or etched marks should be discarded.
Heat-resistant glass or plastic sample containers should be autoclaved
at 1210 C for 15 min. Alternatively, dry glass containers can be sterilized
in a hot-air oven at 1700 C for at least 2 h. For plastic containers that
are not heat-resistant, ethylene oxide gas sterilization is acceptable
(Bordner et al. 1978). Containers sterilized by gas should be stored at
least 12 h before use to ensure all gas has dissipated.
If the sample water has residual chlorine, sodium thiosulfate should
be added to neutralize the chlorine and thereby prevent continued bactericidal
action after sample collection. In this manner, the true microbial content
of the water at the time of sampling can be estimated more accurately.
If sodium thiosulfate must be added to a sample, it should be added to
the sample container prior to sterilization so that the final concentration
in the sample Mill be 100 mg/L. For a 120-mL container, 0.1 ml of a 10-
percent solution of sodium thiosulfate will neutralize a sample containing
as much as 15 mg/L of residual chlorine (APHA 1985).
If the sample water contains heavy metals in concentrations exceeding
0.01 mg/L, a chelating agent should be added to the sample container to
reduce metal toxicity. This is particularly important if samples are not
analyzed within 4 h after collection. APHA (1985) recommends using the
disodium salt of ethylenediaminetetracetic acid (EOTA), adjusted to pH
77
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Effluent
Enterococcus Bacteria
6.5, and added to the sample container before sterilization. For a 120-ml
container, addition of 0.3 ml of a 15-percent EDTA solution is considered
adequate (APHA 1985).
It is critical that samples are not contaminated during the collection
process. To avoid contamination, sterilized containers should be kept
sealed until they are used, containers should oe filled without rinsing,
ana container lids snould be replaced immediately after the samples have
uten collected. When removed from containers, lids should be heU face
down in one hand and not set down on any surface. Adequate headspact (at
least 2.5 cm) should be left in each sample container to facilitate mixing
prior to analysis.
Process ing--Samples should be analyzed as soon as possible after col-
lection. If a delay occurs, samples should be held at 40 C for a maximum
of 6 h. The length of delay should be noted on the log sheet.
Laboratory Procedures--
Analytical Methods—Methods for analyzing enterococcus bacteria are
currently being finalized by U.S. EPA.
Calibration and Preventive Maintenance—This information is reviewed
extensively in Part IV of Bordner et al. (1978) and in Part 902 of APHA
(1985).
Quality Control Checks—Qua! ity control checks for these analyses
are listed in detail in Part IV of Bordner et al. (1978) and in Part 902
of APHA (1985). The list includes:
• Sterility checks on media, dilution and rinse water, glassware,
and membrane filters
78
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Effluent
Enterococcus Bacteria
t Duplicate analyses on 10 percent of samples and on at least
on-» sample per test run
0 Colony verifications on a monthly basis.
Corrective Action — Procedures detailed in the relevant sections of
3c"dner et al. (1978) and APHA (1985) should be followed.
Data Quality and Reporting—Data should be reported according to the
specifications currently being finalized by U.S. EPA.
79
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MONITORING THE RECEIVING ENVIRONMENT
Recommended methods for measuring receiving-environment variables
during 301(h) monitoring programs are described in this section. The initial
major section includes general procedures for sampling preparation, sampling,
and sample handling. The section on sample handling contains tables that
sjmmarize the recommended coTaction specifications and analytical methods
for specific receiving-environment variables. The second major section
presents detailed procedures for measuring 21 recp n'ng ^.•.vironment variables.
GENERAL METHODS
Sampling Preparation
Prior to each field survey, a sampling and analysis plan should be
prepared. It should summarize all of the elements essential for conducting
the survey. The chief scientist or a designee should thoroughly review
the plan (including QA/QC criteria) before each cruise, and ensure that
its essential elements are understood by all members of the scientific
party. The review should focus on the completeness of the plan and the
clarity of its objectives. A complete sampling and analysis plan should
contain the following major elements:
• Identification of scientific party and the responsibilities
of each member
• Statement and prioritization of study objectives
• Description of survey area, including background information
and station locations
• Identification of variables to be measured and corresponding
required containers and preservatives
80
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t Identification of all sample splits or performance samples
to be submitted with the samples
• Brief description of sampling methods, including positioning
technique, sampling devices, replication, and any special
considerations (including handling and shipping hazardous
materials)
• Detailed cruise schedule, including time, date, and location
of embarkation and debarkation
0 Identification of onshore laboratories to which samples
should be shipped after cruise completion
• Survey vessel requirements (e.g., size, laboratory needs,
sample storage needs)
• Location and availability of an alternate survey vessel
t All special equipment needed for the survey (e.g., camera,
nets, communication devices)
• All pertinent QA/QC procedures.
It is essential that the study objectives and their prioritization
be understood by all members of the scientific party. This will ensure
that if modifications of the survey plan become necessary in the field,
their impact on the overall goals of the cruise can be evaluated adequately.
After the sampling plan has been reviewed, contingency plans should be
outlined. These plans should include potential problems and their solutions.
Possible solutions to some problems frequently encountered during offshore
surveys are listed in Table 4. Development of contingency plans can be
greatly assisted by reviewing cruise summary reports and consulting with
chief scientists of previous cruises.
81
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TABLE 4. EXAMPLES OF PROBLEMS FREQUENTLY ENCOUNTERED DURING
OFFSHORE SURVEYS AND RECOMMENDED SOLUTIONS TO EACH PROBLEM
Probh
Recommended Solution!s)
Sampling equipment fails or is lost overboard
Sampling efficiency reduced by poor * j'.
Sampling efficiency 'educed aecause of seasickness
Sampling delayed because vessel is inoperative
A sample is partially lost or slightly contam-
inated
Bottom sampler will not penetrate to a sufficient
depth after repeated casts
One or more water bottles fall to trip on
a particular cast
- Have necessary tools onboard to make repairs
- Maintain spare parts inventory for major
equipment
- Have back-up equipment onboard
- Have SCUBA equipment and divers onboard
for retrieval
- Know where nearest tools, back-up equipment.
or divers are located on shore
- Extend cruise length
- Reschedule cruise
- Ensure scientific crew is large enough
to compensate for reduced personnel
- See solutions to second problem
- Reschedule cruise
- Extend cruise length once vessel is repaired
- Charter an alternate vessel
- Discard and take another sample
Add weight to the sampler
Move a short distance before taking next
sample
Discard all samples from that cast and
take a new cast
82
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The captain of the survey vessel should be provided with a copy of
*.?ie survey plan to ensure that it is consistent with the equipment and
capabilities of the vessel. Modifications to the ship or cruise plan may
se required.
To ensure that all required sampling equipment and supplies are available
at the time of sampling, an equipment checklist should be constructed.
Spare parts and backup supplies should be included in the inventory. All
equipment should be inspected before the cruise, with sufficient time allowed
•o make necessary repairs or replacements. At the end of each cruise,
eauipment should be inspected, cleaned, and stored properly.
Sampling Procedures
Station Location-
Accurate navigation is essential to ensuring that stations can be
plotted and reoccupied with a high degree of certainty. Because compliance
with 301 (h) regulations usually requires that stations oe located at specified
distances relative to a fixed-point discharge (e.g., within-ZID, ZID-boundary)
or that stations be sampled seasonally or annually, failure to accurately
locate or relocate stations can strongly affect the results and conclusions
of a survey. For example, if a within-ZID station is positioned erroneously
outside the ZID, the resulting measurements may suggest that the environmental
impacts of the discharge are much less than they actually are. Alternatively,
if a station thought to be outside the ZID is located mistakenly within
the ZID, conditions would appear to be worse than they really are. As
an additional example, failure to accurately relocate a station may produce
results that suggest conditions are improving, when they actually may be
deteriorating.
Although a variety of navigation and/or position fixing systems are
available currently, factors such as price, availability, and accuracy
vary considerably among them (Tetra Tech 1986c). The station positioning
system selected for a given survey should be able to -neet all regulatory
requirements for accuracy and should, at a minimum, provide a high degree
83
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of precision (i.e., repeatable measurements). Positioning systems that
are precise but lack a high degree of accuracy may be used after actual
station locations are determined by accurate, independent means (i.e.,
"ground-truthed"). For bottom-related samples, all positioning systems
snould be used in conjunction with a fathometer to ensure that sampling
occurs at the proper water depth (allowing for tidal stage and any fathometer
corrections).
Water Sampling--
For most 301(h) monitoring programs, *ater column variables will be
sampled usi'i water bottles, in sit-j instrumentation, or a combination
of both of t ese techniques.
Water Bottles—Water bottle samplers are relatively simple devices
that generally consist of some type of cylindrical tube with stoppers at
each end and a closing device that is activated by a messenger or an electrical
signal. The most ccnmonly used samplers of this description are the Kemmerer,
Van Oorn, Niskin, and Nansen samplers. Each device samples a discrete
parcel of water at any designated depth. Frequently, multiple water samplers
are fixed on a rosette frame so that several depths can be sampled during
one cast or replicate samples can be taken at the same depth.
Prior to deployment, the stoppers of water bottle samplers are cocked
open on the sampling vessel. At this step, it is critical that the interior
of the sampler and stoppers remain free from contamination. All members
of the sampling team should therefore avoid touching the insides of the
sampler and stoppers, and all samplers and stoppers must be rinsed thoroughly.
After cocking, the sampler is lowered to a Designated depth. The
sampler must be open at both ends r>o that water is not trapped within the
device as it is being lowered through the water column. Once the sampler
reaches the desired depth, it should be allowed to equilibrate with ambient
conditions for 2-3 min before being closed.
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After equilibration, the closing device can be activated by messenger
or electrical signal, and the sampler can be retrieved. It is recommended
tnat at least two samplers be used simultaneously for each depth. A second
sampler will provide a backup to the primary sampler in case the latter
device misfires or won't trigger. This will eliminate the need for an
additional cast. A second sampler will also supplement the primary sampler
if the volume collected by the latter device is too small for all required
subsampl ing and rinsing. To ensure that all subsamples at a particular
depth are collected from the same water parcel, it is essential that they
all be taken from a single cast. Multiple casts using a single water sampler
will not meet this objective. Sample water must therefore be used conser-
vatively after collection.
Once the water sampler is brought on board the sampling vessel, the
stoppers should be checked immediately for complete seals. If a stopper
is not properly sealed, water from the sampled depth may have leaked out
upon retreival and been replaced by water from shallower depths. Because
this kind of contamination can bias results, the entire water sample should
be rejected.
Accepted water samples should be subsampled as soon as possible because
appreciable delay may result in unrepresentative subsamples. For example,
measurement of variables sensitive to biological alteration (e.g., dissolved
oxygen, turbidity, color, nutrients) or settlement within the water sampler
(e.g., total suspended solids, settleable solids, phytoplankton) can be
biased substantially by subsampling delays.
In Situ Instrumentation—A wide variety of instruments capable of
measuring water column variables in situ are available. Most are deployed
from the sampling vessel using a cable. Sensors housed within the instruments
measure the variables of interest and transmit data in the form of electrical
signals back to recorders on the survey vessel. The simplest instruments
measure conductivity (i.e., for conversion to salinity), temperature, and
water pressure (i.e., for conversion to depth). Additional sensors and
instrumentation can be included to measure a variety of additional water-
column variables such as dissolved oxygen, pH, transmissivity (i.e., an
85
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index of turbidity), oxidation-reduction potential, and specific ions (e.g.,
a/rnonia). Generally, the operating manuals supplied with these instruments
provide detailed descriptions of how to calibrate, operate, and maintain
the equipment. If a particular manual lacks sufficient detail, the manufacturer
snould be contacted for specific guidance.
Although the instrument operating manuals should be consulted for
specific instructions, several general procedures for operating j_n situ
instruments apply to all or most devices, and hava a direct influence on
data quality. When acquiring in situ instruments for use in marine and
estuarine waters the following features are highly recommended:
• Instruments should be of rugged constru"tion, corrosion-
resistant, and waterproof
0 Instruments should be capable of operating with acceptable
accuracy within the range of expected environmental condi-
tions
t Cables should be of adequate length and strength
• Electrical connectors should be easy to use, waterproof
when connected, and capable of being locked after connection
• External sensors should be protected by housings or other
means
• Instruments should be easy to calibrate on board the survey
vessel
• Ideally, a service center should be located nearby so mat
an instrument can be repaired rapidly, if necessary
• If sample contamination may be a problem, all sampling equipment
should be made of non-contaminating material.
86
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When using _l£ situ instruments it is critical that it be protected
from rough handling and adverse environmental conditions. The following
precautions should always be taken:
• Instruments should be transported in specially designed
shipping boxes
• Instruments should oe surrounded by a "birdcage" when being
deployed. Frequently, instruments are attached within a
rosette frame when they are used in conjunction with water
bottles. Caution should be taken to ensure that the "birdcage"
or rosette frame do not create sampling artifacts
• Instruments should be securely lashed down in a safe area
when on deck
0 Instruments should be rinsed with fresh or distilled water
after each submersion
• Optical surfaces should be cleaned with alcohol and lens
tissue after each submersion
• Instruments should be protected from direct sunlight and
excessive heat, as plastic components may be damaged by
heat
• External sensors and optical ports should be covered and
protected whenever the instrument is not being used.
When operating in situ instruments, the following procedures should
be followed to ensure that instruments are prepared, deployed, ana retrieved
properly:
• Instruments should be allowed to warm up for a sufficient
length of time prior to calibration or deployment
87
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• Instruments should oe field-calibrated at the beginning
of each day of sampling. All circuits should be tested
at the same time. Calibration should be conducted more
frequently if equipment malfunctioning is suspected
• Upon instrument deployment, the survey vessel should be
anchored or drifting slowly
• Instruments should be deployed relative to vessel construction
and sea conditions so that cables will not tangle in the
propel lor or rudder assemblies
• When measuring continuou: profiles, the lowering speed through
the water column should not exceed the equilibration rate
of the sensor having the slowest response time
• Excessive strain should not be placed on the cable(s), as
it could disrupt electrical connections.
Routine maintenance and inspection of in situ instruments should follow
the manufacturer1 s recommendations. General procedures include:
• All rubber parts of underwater connectors should be coated
with silicone grease to ensure proper lubrication
• Plugs should be inspected for bent or broken pins, which
may cause faulty connections and flooded cables
• Cables should be inspected for nicks, cuts, abrasions, or
other signs of physical damage
• Seals should be inspected and periodically cleaned and greased
to ensure a waterproof fit
• Oesiccant should be inspected and replaced with fresh or
reactivated desiccant when necessary.
-------
Factory inspection and recal ibration at reccrrniended intervals is essential
to ensure that an in situ instrument is functioning properly and will continue
to function properly during future cruises. It is recommended that factory
service be conducted at least once per year. Factory service should always
be conducted when instrument malfunctions cannot be corrected by following
the operating manual. Factory service may also be required when part of
an in situ system is replacec, as all components are not interchangeable
without factory recalibration.
Log Sheet — Variables that should be recorced on the water sampling
log sheet are:
• Geographic location
0 Date and time
• Weather conditions
Sea state
Sky state
Precipitation
• Station number
• Sample number
t Replicate number
t Position coordinates
• Total depth
• Sampled depth(s)
• Sampler description
Kind
Volume
Rosette, paired, single
• Kinds of subsamples
• Comments relative to sample quality
• Names of chief scientist and sampling team
• Vessel name.
89
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G'-ab Sampl ing—
Most sampling of benthic infauna and sediments for 301 (h) monitoring
p-ogr*Tis will be conducted using grab samplers. Benthic organisms may
be used for a variety of purposes, such as 1) comparing species composition
ar.d abundances between potentially impacted and reference areas (spatial
trends) or between sampling periods (temporal trends), and 2) evaluating
bioaccumulation of toxic substances in body tissues. Sediments may be
used for such purposes as 1) comparing characteristics of grain sizes and
concentrations of conventional and toxic substances between potentially
impacted and reference drees or between sampling periods, and 2) relating
sediment characteristics to observed distributions of benthic infauna.
To be capable of collecting infauna and sediments suitable for quantitative
comparisons, the grab must be appropriate for the sediment and depth conditions
in the study area, and must be operated properly following standardized
procedures. Holme (1971) discusses the details of grab selection and oper-
ation. A thorough reading of that reference is highly recommended.
Grab Selection—Numerous grab samplers are available for collecting
benthic infauna (see Holme 1971). This variety exists because field conditions
(e.g., weather, water depth, sediment type) are not uniform, and because
the structure (e.g., species composition, abundance) and function (e.g.,
trophic relationships, energy flow) of benthic communities vary spatially
and temporally. The grab selected for use should be able to perform reliably
and efficiently under the anticipated field conditions and given the anticipated
types of benthic communities that will be sampled.
To be reliable, a grab must collect useful samples consistently within
its design undermost anticipated field conditions. That is, each time
the grab is deployed there should be a high probability of collecting a
quantitative sample. Matching sampler design with the predominant sediment
types is especially important for achieving a high degree of reliability.
The grab should also collect representative samples of the populations
that comprise the infauna (i.e., be efficient). To be efficient, the "bite"
into the sediments should be deep enough to collect most of the organisms
within the jaws of the grab. Moreover, the "bow wake" associated with
90
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trie descending grab should be as small as possible to minimize blowing
away the organisms at the sediment-water interface immediately before the
grab contacts the sediment.
The installation of mesh screens instead of solid plates on the tops
of the grab jaws will help minimize the "bow wake", and will keep organisms
from escaping once the grab jaws are closed. The screen mesh apertures
should be the same as those used for sieving samples in the field. The
screens should be hinged to provide easy access to the surface of the sample
so that grab penetration can be estimated, the condition of the surface
sediments can be observed, and sediment subsamples can be collected. The
exterior of the hinged screens hould be fitted witn rubber flaps that
close when the grab is being retrieved, but open when the grab is being
deployed. Although these flaps may increase the bow wake slightly (even
when they open on descent), they will help minimize winnowing of the surficial
sediments when the grab is retrieved.
In addition to selecting the proper kind of sampler, it is important
to choose a grab that samples an appropriate area of bottom. This should
be done prior to the major sampling effort using data from historical studies
or a preliminary survey. There are two recommended ways to determine the
appropriate area to sample (Gonor and Kemp 1978):
1. Choose a sample unit size that collects a representative
complement of the species present, or
2. Choose a sample unit size for which the variance-to-mean
ratio for major community variables is low.
The first criterion is met by plotting the number of species collected
as a function of different sample unit sizes. -At the point on the curve
where the number of species approaches an asymptote, the sample unit size
is considered sufficiently large to collect a representative complement
of species. The second criterion is met by calculating the variance-to-
mean ratios of various sample unit sizes for a given variable. A low ratio
is desirable for accurate and precise estimates of mean values and for
91
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statistical comparisons of mean values among stations. The variance-to-
r*an ratio changes with sample unit size because benthic organisms are
attributed patchily in most cases (Elliott 1971). When the sample unit
s:ze is much smaller than the patch size, the ratio generally is low.
As the sample unit size increases to about that of the patch, the ratio
usjally reaches its highest value. The ratio then declines with further
increases in sample unit size.
In summary, the ideal benthic grab sampler is reliable, efficient,
large enough to collect a representative complement of species, *nd small
erough to yield low variance-to-mean ratios for fiajor community variables
(e.g., to* 1 numbers of species, total numbers of individuals). Unfortunately,
S'-ch a sai pier has yet to be invented. However, consideration of the forgoing
factors should result in the selection of a dependable sampler that yields
information adequate to meet the objectives of the monitoring program.
In most coastal and estuarine areas, a 0.1-m2 (I.l-ft2) van Veen grab and
a 0.1-m2 (I.l-ft2) Smith-Mclntyre grab are adequate in most silty and sandy
substrates. However, preliminary tests should be conducted for various
samplers in all areas that have not been sampled previously, and in areas
wf.ere the sediments are predominantly clay or gravel.
Grab Deployment and Retrieval—The grab sampler should be attached
to a hydraulically operated cable and should be preceded by a ball-bearing
swivel (Figure 3). The swivel and all shackles connecting the cable to
the grab should have a load capacity at least five times that of a loaded
grab. It may be desirable to keep the shackles and swivel small, however,
as it may be necessary for the shackles to travel through the block at
tne end of the davit when retrieving the grab. A snatch block.at the end
of the davit in Heu of a standard block may also facilitate grab deployment
and retrieval. For safety reasons, a safety pin should be used to prevent
the grab f^om triggering accidentally. Also, the grab should be deployed
with a minimum of swinging when out of the water. Use of handling lines
(Figure 3) clipped to the hydrowire is an effective method for minimizing
grab swinging.
92
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DAVIT
SNATCH BLOCK
RAIL OF SHIP
Figure 3. Deployment of a grab sampler.
HANDLING UNE
WITH SNAP HOOKS
V
93
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The grab sampler should be lowered and raised through the water column
at a speed of approximately 20 m/min (66 ft/min) using the vessel's hydraulic
system. This lowering rate generally allows adequate penetration of the
aev ice into the sediment while minimizing the chance that the sampler will
be flipped over by the weight of excess wire cable falling faster than
the parachuting sampler. If adequate penetration cannot be achieved using
a controlled descent, the sampler can be allowed to free fall the last
1 or 2 m. However, extra attention should be paid to evaluating surface
disturbance under sjch conditions. Flipping the grab sampler due to excess
caole weight occurs most frequently when sampling at depths in excess of
200 m (656 ft). At shallower depth., the weight of the cable is generally
lighter than the weight of the sampling device. When more than 20C i (656 ft)
of cable is deployed, care must also be taken >.o ensure that the cable
does not pull itself off the reel.
After the grab has contacted the bottom, it should be retrieved slowly
at first to permit the grab jaws to close properly. After the jaws are
closed, a retrieval speed of 20 m/min (66 ft/min) should be maintained.
When the grab approaches the water surface (i.e., when first sighted),
trie winch should be stopped to permit the handling lines to be clipped
onto the cable. The grab can then be raised slowly, and the handling lines
can be used to minimize swinging of the grab. Excessive swinging is unsafe
and may also compromise the quality of the sample. When brought on board,
the grab and sample should be lowered into a waist-high stand designed
to receive it.
Once the sampler Is brought aboard, the hinged screens can be opened
and the sample examined. The sediment should be characterized using the
following qualitative physical characteristics: sediment texture; sediment
color; presence, type, and strength of odor; grab penetration depth [to
nearest 0.5 cm (0.2 in)]; degree of leakage (if any); sediment surface
disturbance; and obvious abnormalities, such as wood debris or large quantities
of mollusc fragments. Samples showing excessive leakage or disturbance
of the sediment surface should be rejected (Figure 4). It is also recommended
that samples be rejected if they do not achieve the following minimum pene-
tration depths in various types of sediment:
94
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ACCEPTABLE IF MINIMUM
PENETRATION REQUIREMENT MET
UNACCEPTABLE (WASHED. ROCK
CAUGHT IN JAWS)
UNACCEPTABLE (CANTED
WITH PARTIAL SAMPLE)
UNACCEPTABLE (WASHED)
Figure 4. Examples of acceptable and unacceptable grab ..~..,/ies
-------
• Cobble/pebble substrate cannot be sampled adequately using
conventional grab samplers
• 4 cm (1.6 in) for coarse sand/gravel
• 5 cm (2.0 in) for medium sand
• 7 cm (2.7 in) for fine sand
• 10 cm (3.9 in) for sandy silt and silty sand
t 10 cm (3.9 in) for si it
• 10 on (3.9 in) for clay.
See Shepard (1954, 1963) for definitions and size ranges of the foregoing
granulometric terms.
The acceptable penetration depths are based on the vertical distri-
bution of organisms found in these different sediment types. Generally,
they are sufficient to insure that at least 95 percent of the organisms
and species that would have been encountered to depths of 20 cm (7.9 in)
are captured. All of the foregoing grab sample characteristics should
be recorded on the field log sheet.
Log Sheet—Variables that should be recorded on the grab sampling
log sheet are:
• Geographic location
• Date and time
• Weather conditions
Sea state
Sky state
Precipitation
96
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• Station number
• Replicate number
• Position coordinates
• Depth
• Sampler description
Kind
Area sampled
t Number and kinds of subsamples
t Sediment characteristics
Texture
Color
P or
S'ructures
Debris
Degree of leakage
Surface disturbance
• Penetration depth
• Sieve mesh size
• Comments relative to sample quality
t Names of chief scientist and sampling team
• Vessel name.
Trawl Sampling--
Most sampling of demersal fishes and epibenthic macroinvertebrates
for 301(h) monitoring programs will be conducted using small otter trawls.
Sampled organisms may be used for a variety of purposes, such as 1) quantitative
comparison of abundances between potentially impacted and reference areas
(spatial trends) or between sampling periods (temporal trends), 2) determination
of prevalences of grossly visible external abnormalities, and 3) evaluation
of bioaccumulation of contaminants in tissue (Tetra Tech 1985a). To be
capable of collecting data suitable for quantitative comparisons, the otter
trawl must be designed in a specific manner and operated using standardized
procedures. Ideally, at least one member of the trawling team should have
ample experience in using this device. Mearns and Allen (1978) provide
the most comprehensive description of how small otter trawls should be
97
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cesigned and used for conducting biological surveys in coastal waters.
A thorough reading of that document is highly recommended.
Trawl Design—To ensi re proper operation, it is essential that the
otter trawl be constructed by an experienced manufacturer using a proven
net design. The net should be large enough to capture adequate numbers
of fishes, but small enough to be handled easily by a crew of two or three
persons on a medium-size [i.e., 9-15 m (30-50 ft)] vessel. Past experience
with 301(h) data collection has shown that a net with headrope length of
approximately 7.6 m (25 ft) is an acceptable size.
The footrope of the Uer trawl must be weighted to ensure that fishes
living close to the bottom are adequately sampled. This capability is
especially critical for sampling flatfishes (i.e., PIeuronectiformes), as
many of these individuals lie partly buried in bottom sediments when resting.
Mearns and Allen (1978) recommend using a chain rather than lead weights
on the footrope because its effect on the digging characteristics of the
net is easier to adjust.
Floats must be attached to the headrope of the otter trawl to hold
the mouth of the net open when sampling. When sampling at depths greater
than 15-30 m (50-100 ft), it is essential that floats be designed to withstand
high ambient pressure without deforming. Floats made of styrofoam or cork
tend to compress under high pressure, whereas hollow plastic floats tend
to burst.
Otter boards should be fitted with a minimum of four towing chains
to allow fine-scaled adjustment of the angle at which the boards lean when
towed. A proper angle of attack is critical to holding the mouth of the
net open when towing. The wear pattern on the shoe of the otter board
can indicate whether a board is operating properly. In addition, the manner
in which the boards spread the net immediately following submersion also
indicates whether they are functioning correctly. Mearns and Allen (1978)
describe how to adjust the otter boards.
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The net bridles should be flexible (e.g., braided nylon) and attached
to the hydrowire with a two- or three-way swivel. This arrangement minimizes
the twisting forces on the net assembly and allows it to take the shape
and orien-ation for which it was designed. Mearns and Allen (1978) recommend
that the length of the bridles be approximately three times the headrope
length to ensure that the otter boards spread properly.
Trawl Operation--Before sampling a transect, it is advisable to slowly
make a pass along the isobath of interest and monitor its course with a
fathometer. If the isobath is highly irregular and requires the vessel
to change course frequently, an alternate transect should be considered.
To allow the otter trawl to sample a a uniform depth in a consistent manner,
the transect should be nearly straigl1: or broadly curved.
After a sampling transect has been selected, the trawl assembly should
be readied for deployment. The net should be laid out on the deck and
scanned for rips. All lines should be untangled and their attachment points
checked. The vessel should begin maneuvering toward the transect, and
the net should be lowered into the water a sufficient distance from the
sampling transect to ensure that it will touch bottom at the beginning
of the transect. Immediately after the otter boards enter the water, the
winch should be stopped and the boards allowed to spread. If the boards
do not open the mouth of the net properly, the assembly should be retrieved
and adjusted. After the otter boards spread properly, the trawl assembly
should be lowered at a slow and steady speed while the vessel is underway.
If the descent is erratic, or tension is reduced on the hydrowire, the
otter boards may cross, requiring the trawl assembly to be retrieved and
redeployed.
After the net touches bottom, a sufficient length of hydrowire (i.e.,
scope) should be payed out to ensure that the net is pulled from a horizontal,
rather than vertical position. If the scope is insufficient, the net will
tend to leave the bottom and inadequately sample fishes residing at the
sediment-water interface. In general, required scope declines with increasing
depth because the additional weight of the hydrowire enhances the horizontal
99
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component of the towing forces. Based on general guidance given by Mearns
anc! Allen (1978), the following scope-to-depth ratios are reconmended:
• 5:1 for depths < 20 m (66 ft)
• 4:1 for depths of 20-75 m (66-246 ft)
• 3.5:1 for depths of 75-200 m (246-656 ft)
• 3:1 for depths of 200-350 m (656-1,148 ft)
• 2:i for depths > 350 m (1,148 ft).
It is critical that all stations at a particular depth be sampled using
the same scope. Failure to use a consistent scope can result in variable
sampling efficiencies among stations.
After sufficient scope has been payed out, the winch'should be locked.
Sampling begins at that point. When sampling, the vessel should proceed
at a constant speed of approximately 2.5 kn (4.2 ft/sec) relative to the
seabed. If the speed is not held constant, the shape of the net mouth
will vary and the transect will not be sampled consistently throughout
its length. Gibbs and Mathews (1982) found that an 8-m (26-ft) otter trawl
fishes most efficiently when towed at about 2.5 kn. Faster speeds caused
the footrope to leave the bottom, whereas slower speeds caused the spread
of the trawl to decline and the otter boards and footrope to dig excessively
into soft substrates.
When a surface current or substantial wave action occurs, the vessel
should point into the current or waves when sampling providing this can
be accomplished within the constraints of the study plan. It is strongly
recommended that fishing effort be based on transect length rather than
time of trawling. As shown in Figure 5, a 5-min haul at a constant vessel
speed of 2.5 kn (relative to the water surface) will cover a distance of
0-385 m (0-1,262 ft), depending upon the speed of the opposing surface
100
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CO
cc
UJ
UJ
UJ
UJ
i — i — i — i — i — i
0.2 0.4 as as uo 1.2 1.4 is 18 2.0 2.2 2.4 2.6
CURRENT SPEED (KNOTS)
Figure 5. Transect length as a function of current
speed. Net is towed at 2.5 knots for 5
minutes into an opposing surface current.
101
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c-rrent. It is therefore very difficult to sample a standardized area
o- the seafloor when fishing effort is based on time of trawling.
When a transect has been covered completely, the winch should be engaged.
Sampling ends at that point. The net should be retrieved at a constant
speed as the vessel moves forward slowly. Constant tension should be maintained
on the net to prevent captured fishes from escaping. Ideally, the net
assembly should break the water in its fishing orientation.
After the net assembly is retrieved and secured, the catch can be
c-ocessed. Qualr.y assurance procedures related to catch processing are
given in later sections of this report concerning bioaccunulatior and assemblage
cnaracteristics of demersal fishes and epibentnic macroinvertetr-ates.
Log Sheet—The variables that should be recorded on the trawl sampling
log sheet are:
0 Geographic location
• Date
• Weather conditions
Sea state
Sky state
Precipitation
• Station number
• Replicate number
• Depth
• Vessel heading
0 Vessel speed
• Surface current
Speed
Direction
• Hydrowire length
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• Position coordinates and times
When net begins descent
When winch Is locked
When net begins ascent
• Sampler description
Kind
Headrope length
Body mesh size
Cod-end liner mesh size
0 Number and kinds of subsamples
• Comments relative to sample quality
• Names of chief scientist and sampling team
• Vessel name.
Sample Handling
After sample collection, proper sample handling ensures that changes
in the constituents of interest are minimized and guards against errors
when shipping and analyzing samples. Recommended sample size, containers,
preservation, and storage requirements for each offshore variable are summarized
in Table 5. Recommended laboratory methods for measuring receiving-environment
variables are listed in Table 6. These requirements and methods are also
noted in subsequent sections. These requirements and methods should be
reviewed in advance by the laboratory personnel to ensure that sample sizes,
containers, preservatives, and all other specifications are consistent
with laboratory needs and capabilities.
Field Procedures--
It is important throughout any sampling and analysis program to maintain
integrity of the sample from the time of collection to the point of data
reporting. Proper chain-of-custody procedures allow the possession and
handling of samples to be traced from collection to final disposition.
Documents needed to maintain proper chain-of-custody include:
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TABLE 5. RECOMMENDED SAMPLE SIZES, CONTAINERS, PRESERVATION,
AND HOLDING TIMES FOR OFFSHORE SAMPLES
,
Measurement
Receiving waterC
PH
Salinity
Temperature
Color
Turbidity
Total suspended solids
Settleable solids
Floating particulates
Dissolved oxygen
Probe
Winkler
Biochemical oxygen
demand
Oil and grease
Nitrogen
Atnmonia-N
Total Kjeldahl-N
Nitrate * Nitrite-N
Phosphorus (total)
Total and fecal
coli form bacteria
Enterococcus bacteria
Chlorophyll a
n i n imum
Sample
Size'
25 mL
200 mL
1 L
SO mL
100 mL
1-4 L«
1 L
5 L
300 mL
300 mL
1.000
1.000
400 mL
500 mL
100 mL
SO mL
250-500
250-500
2-4 L«
Container*
P.G
P.G
P.G
P.G
P.G.
P.G
P.G
P.G
6 bottle &
top only
G bottle &
top only
P.G
6 only
P.G
P.G
P.G
P.G
mL P.G
mL P.G
P.G
Preservative
None
None
None
Cool, 40c
Cool . 40c
Cool . 4<>c
Cool. 40C
None
None
Fix on site; store
in dark
Cool. 4<>c
Cool, 40c
H2S04 to pH<2
Cool , 40C H2SOd to
pH<2
Cool. 40c H2SOa to
pH<2
Cool, 40C H2S04 to
pH<2
Cool, 40c H2SO« to
pH<2
Cool . 40 c
0.0081 Na2S203?
Cool, 40 c
0.0081 Na 282039
Freeze at -20°
Maximum
Holding
Time
Analyze immediately13
Indefinite
Measure immediately*1
48 h
48 h
7 days
48 h
Analyze Immediately"1''
Analyze immediately4
8 h
48 h
28 days
28 days
28 days
28 days
28 days
6 h
6 h*
21 days*
Phytoplankton
1 L
P.G
in the dark
in a desiccator
101 formal in
Indefinite"
104
-------
TABLE S. (Continued)
Sediment/ In fauna
Grain size
Total solids
Total volatile solids
Total organic carbon
Biochemical oxygen demand
Chemical oxygen demand
Oil and grease
Su If ides
Total
Water soluble
100 g
SO g
SO g
50 g
SO g
25 g
50 g
50 g
10 g
P.G
P.G
P.G
P.G
P.G
P.G
G only
P,G
P.G
Cool. 40 c
Freeze
Freeze
Freeze
Cool, 40 c
Cool, 40 c
Cool 40 C;
or freeze
Cool , 40 c
zinc acetate
Cool, 40 c
SAOB
6
-------
TABLE 6. RECOMMENDED METHODS FOR MEASURING RECEIVING-ENVIRONMENT VARIABLES
Method Reference
Variable
Receiving Water
pH
Salinity
Temperature
Color
Turbidity
Transmissivity
Total suspended solids
Settleable solids
Floating particulates
Dissolved oxygen
Probe
Uinkler
Biochemical oxygen demand
Oil and grease
Nitrogen
Ammonia-N
Total Kjeldahl-N
U.S. EPAa
150.1
-
170.1
110.3
180.1
-
160. 2f
160.5
-
360.1
405.1
413.1
413.2
350.1
350.2
350.3
351.1
351.3
APHAb
423
-
212
2048
214A
-
209Cf
209E
206A9
42 IF
507
503A
503B
41 7A
41 7B
4170
4176
41 7B
4170
41 7E
42 OA
420B
Otherc
ln_ situd
Salinometere
In situd
In situd
-
-
In situd
-
-
-
In situd
SFrTcTTand
and Parsons
(1972)
-
-
In situd
106
-------
TABLE 6. (Continued)
Nitrate+nitrite-N
Phosphorus (total)
Total coliform bacteria
Fecal coliform bacteria
Enterococcus bacteria
Chlorophyll £
Phytoplankton
Sediment
Grain size
Total solids
Total volatile solids
Total organic carbon
Biochemical oxygen demand
Chemical oxygen demand
Oil and grease
Sulfides
Total
Water soluble
353.2
353.3
365.1
365.2
365.3
418C
418F
42 4C
42 4F
42 4G
908A"
909Ai
908Ch
909C1
p. H4h.j
p. 1081.J
p. 132" .J
p. 1241 ,j
U.S. EPAk
Strickland
and Parsons
(1972)
Stofan and
Grant (1978)
Plumb (1981)
Plumb (1981)
Plumb (1981)
Plumb (1981)
Plumb (1981)
Plumb (1981)
Plumb (1981)
Plumb (1981)
Green and
Schnitker
(1974)
107
-------
TABLE 6. (Continued)
Priority pollutant metals - - Tetra Tech
(1986a)
Priority pollutant organic compounds - - Tetra Tech
(1986a)
Infauna - - Present
document
Bioaccumulation (tissue)
Priority pollutant metals - - Tetra Tech
(1986b)
' loricy pollutant organic compounds - - Tetra Tech
(1986b)
a Methods recommended in U.S. EPA (1979b).
b Methods recommended in APHA (1985).
c Methods recommended in sources other than U.S. EPA (1979b) or APHA (1985)
when no methods were recommended in the latter two sources.
d This variable can be measured using an in situ instrument. The operating
manual for the instrument should provide all necessary information for
proper instrument calibration and measurement of this variable.
« The instruction manual for the salinometer should provide all necessary
information for instrument calibration and salinity determination.
f A 0.40- or 0.45-um membrane filter should be used instead of the glass-
fiber filter recommended in the method.
9 This method is tentatively recommended by APHA.
n This method can be used whether or not chlorine is present.
i This method can be used only when chlorine is absent.
J Page reference of this method in Bordner et al. (1978).
k U.S. EPA is currently finalizing a recommended method for this variable.
108
-------
Field logbook -- All pertinent information on field activities
and sampling efforts should be recorded in a bound logbook.
The field supervisor should be responsible for ensuring
that sufficient detail is recorded in the logbook. The
logbook should enable someone else to completely reconstruct
the field activity without relying on the memory of the
field crew. All entires should be made in indelible ink,
with each page signed and dated by the author, and a line
drawn through the unused portions of any page. All corrections
should consist of permanent line-out deletions that are
initialed by the chief scientist. At a minimum, entries
in a logbook should include:
Date and time of starting work
Names of field supervisor and team members
Purpose of proposed sampling effort
Description of sampling site, including information
on any photographs that may be taken
Location of sampling site
Details of actual sampling effort, particularly deviations
from standard operating procedures
Field observations
Field measurements (e.g., pH, temperature, flow)
Field laboratory analytical results
Sample identification
109
-------
Type and number of sample bottles collected
Sample handling, packaging, labeling, and shipping
information (including destination).
Chain-of-custody procedures should be maintained with the
field logbook. While being used in the field, the logbook
should remain with the field team at all times. Upon completion
of the sampling effort, the logbook should be kept in a
secure area. All logged information should be summarized
and submitted to U.S. EPA after sampling is completed.
• Sample labels -- Sample labels mi'-.t be waterproof and must
be securely fastened to the outside and/or placed inside
each sample container (depending on the kind of sample)
to prevent misidentification of samples. Labels must contain
at least the sample number, preservation technique, date
and time of collection, location of collection, and signature
of the collector. Labels should be marked with indelible
ink. Abbreviated labels may also be placed on the cap of
each jar to facilitate sample identification.
t Chain-of-custody records -- A chain-of-custody record (Figure 6)
must accompany every group of samples. Each person who
has custody of the sample must sign the form and ensure
that the samples are not left unattended unless secured
properly.
• Custody seals -- Custody seals (Figure 7) are used to detect
unauthorized tampering with the samples. Sampling personnel
should attach seals to all shipping containers sent to the
laboratory by common carrier. Gummed paper seals or custody
tape should be used so that the seal must be broken when
the container holding the samples is opened.
110
-------
SERA
i
i
t
(
(nind Sum R«gl
kgcney SMC
:HAIN OF CUSTODY RECORD
on 10
Sbrth AVWHM
d« WA 98101
PROJECT
LAB *
STATION
DATE
TIME
SAMPLERS: <5«»iw«
SAMPLE TYPE
i
•I
M
»
\
%
i
NUMBt*
Of
CONTAINfNS
i
REMARKS
RELINQUISHED BY: **— RECEIVED BY: a****
RELINQUISHED BY: /SPOT* RECEIVED BY: rspww*
RELINQUISHED BY:
-------
O.S. ENVIRONMENTAL PROTECTION AGENCY
CLP Sample Management Office
P.O. Bos SIS - Alexandria, Virginia 22313
Phone: 703/337-2*90 - FTS/337-2490
1.
2.
J.
7. _
S. .
9. .
10. .
II. .
I*. .
13. .
1%.
«*•
'*•
17.
'«•
19.
20.
SPECIAL ANALYTICAL SERVICE
PACKING LIST
EPA CUSTODY SEAL
SA5 Number
Sampling Office:
Sampling Contact:
(name)
(phone)
Sampling DateUh
Date Shipped!
Site Name/Code:
Ship Tot
Attro
Par Lab Use Only
Date Samples RecM:
Received Byi
Sample Sample Oesc. iption Sample Condition on
Numbers Le., Analysis, Matr' Zoncentratioa Receipt at Lab
For Lab Use Only.
EMYIROHUEMTAL PROTECPOM ACEKCY
MMFUHO.
UQMATUIIB
PNMTMAM8 AH9 nTl«Oi»»eea *a*i*mTi*al*mt
•
I
*
•
IS
Figure 7. Examples of a sample analysis request form (above)
and a custody seal (below). Equivalent material
should be used for 301(h) monitoring.
112
-------
For further information regarding proper chain-of-custody procedures, consult
the policies and procedures manual for the National Enforcement Investigations
Center (NEIC; U.S. EPA 1978).
Sample Shipment--
All preserved samples should oe shipped immediately after completion
of sampling. This minimizes the number of people handling samples, and
protects sample quality and security. Guidance for shipping hazardous
materials can be found in U.S. Department of Transportation (1984). As
samples are prepared for shipping, the following should be kept in mind:
« Shipping ,ontainers should be in good shape and capable
of withstanding rough treatment during shipping.
• Samples should be packed tightly
Dividers must separate all glas-s containers
Empty space within shipping boxes should be filled so that
jars are held securely.
• All containers must be leak-proof. If a container is not
leak-proof by design, the interior should be lined with
two heavy-duty plastic bags and the tops of bags should
be tied once samples are inside. Adequate absorbent material
should be placed in the container in a quantity sufficient
to absorb all of the liquid.
t All samples should be accompanied by a sample analysis request.
Variables to be analyzed by the laboratory, and total number
and kind of samples shipped for analysis should be listed
on the request sheet. An example sample analysis request
form is illustrated in Figure 7. The laboratory should
acknowledge receipt of shipment by signing and dating the
form, and returning a copy to the designated QA coordinator.
113
-------
• A chain-of-custody record for each shipping container should
be filled out completely and signed.
• The original chain-of-custody record and sample analysis
request should be protected from damage ana placed inside
the shipping box. A copy of each should be retained by
the shipping party.
• The custody seal should be attached so that :he shipping
box cannot *ie opened without breaking the seal.
f For shipping containers carrying liquid samples
A "This End Up" label should be attached to each side to
ensure that jars are transported in an upright position
A "Fragile-Glass" label should be attached to the top
of box to minimize agitation of samples.
• Shipping containers should be sent by a carrier'that will
provide a delivery receipt. This will confirm that the
contract laboratory received the samples and serve as a
backup to the chain-of-custody record.
• All shipping charges should be prepaid by the sender to
avoid confusion and possible rejection of package by the
contract laboratory.
Laboratory Procedures--
At the laboratory, ona person should be designated custodian of all
incoming samples. An alternate should also be designated to serve in the
custodian's absence. The custodian should oversee the following activities:
114
-------
• Reception of samples
• Maintenance of chain-of-custody records
• Maintenance of sample-tracking logs
• Distribution of samples for laboratory analyses
• Sending samples to outside laboratories
• Supervision of labeling, log ke<=Ding, data reduction, and
data transcription
• Storage and security of all samples, data, and documents.
Upon reception of samples, a designated laboratory person should fill
out the chain-of-custody record, indicating time and date of reception,
number of samples, and condition of samples. All irregularities indicating
that sample security or quality may have been jeopardized (e.g., evidence
of tampering, loose lids, cracked jars) should be noted on the sample analysis
request form and returned to the client-designated QA coordinator.- In
addition, a designated laboratory person should initiate and maintain the
sample tracking log that will follow each sample through all stages of
laboratory processing and analysis.
Minimum information required in a sample-tracking log includes:
• Sample identification number
0 Location and condition of storage
• Date and time of each sample removal and return to storage
• Signature of person removing and returning the sample
115
-------
• Reason for removal from storage
• Final disposition of sample.
All logbooks, labels, data sheets, tracking logs, and custody records
should have proper identification numbers and be accurately filled out.
All information should be written in ink. Corrections should be made by
drawing a line through the error and entering the correct information.
Corrections should be signed and dated. Accuracy of all data reductions
ard transcriptions should be verified at least twice. All samples and
documents should be properly stored within the laboratory until the client
a-.-.-horues their removal. Security and confidentiality of all stored material
should be maintained at all times. Before releasing analytical results,
all information on sample tags, data sheets, tracking logs, and custody
records should be cross-checked to ensure that data pertaining to each
sample are consistent throughout the record.
Originals of the following documents should be sent to the client:
• Chain-of-custody records
• Sample-tracking logs
• Data report sheets
0 Quality control records.
Copies of all forms should be retained by the laboratory in case originals
are lost in transit.
When replicate analyses are conducted as QA/QC checks, it is recomiended
that they be distributed relatively evenly throughout the full sequence
of samples analyzed, rather than being concentrated at some point (e.g.,
beginning, end) of the sequence. This precaution will enhance the probability
that if problems develop within part of a sequence, they will be detected.
116
-------
S-ipboard Laboratory Analyses
Depending upon the size and capabilities of the survey vessel, many
cf the receiving-environment variables described in this document can be
analyzed on board. In general, the laboratory procedures described in
tms document are applicable to both shipboard and land-based laboratories.
Tnis consistency is important to ensuring that analytical results will
te comparable regardless of which kind of laboratory generates them.
Although most laboratory procedures are similar between shipboard
via land-based laboratories, a number of additional factors must be considered
wnen analyzing samples at sea. These factors relate primarily to the remoteness
cf the shipboard laboratory from land-based support and th-» movement and
limited space of the survey vessel. The major considerations are:
• The design of the laboratory should be efficient, with convenient
equipment locations and adequate storage space.
• The vessel should be equipped with an uninterruptible power
supply that is adequate for operation of all scientific
instruments.
t The laboratory should be well-ventilated to remove any toxic
vapors created by chemicals.
0 The temperature of the laboratory should be controlled,
especially if variations in ambient temperature can influence
particular analyses.
• Adequate lighting is necessary, especially for analyses requiring
color discrimination (e.g., titration endpoints). Fluorescent
lights of the daylight type are recommended (U.S. Navy 1968).
• The laboratory should have adequate water purification apparatus
or be capable of storing water purified on shore.
117
-------
• For storing many kinds of samples, adequate refrigeration
and freezing capabilities are desirable.
t The laboratory should never be used as a general passageway
or lounge.
• The laboratory should be off-limits to unauthorized personnel.
• Adequate safety and first aid equipment should be on board,
preferably including an overhead quick-pull safety shower.
a Extreme care must be taken when handling sample (for quality
purposes) and hazardous reagents (for safety purposes},
as vessel movement can sometimes be unpredictable.
• Backup supplies and instruments should be on board so that
sampling can continue if a piece of equipment is broken
or will not operate properly. A continuously updated inventory
tracking system is useful for maintaining backup equipment.
a All equipment should be properly secured to compensate for
predictable and unpredictable vessel movements. Specially
designed racks are useful for this purpose.
0 Instruments should be checked and calibrated before sailing
so that problems requiring land-based assistance can be
solved quickly.
• Whenever possible, plastic containers should be used instead
of glass because plastic is less susceptible to breakage.
• Instruments having digital displays are preferred over those
using analog displays.
a Pre-printed data sheets should be used to ensure that all
required information is recorded.
118
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RECEIVING WATER ANALYSES
QA/QC procedures are presented in this section for the following 21
receiving water variables:
• pH*
t Salinity*
• Temperature*
• Color
• Transparency
• Turbidity (transmissivity*)
• Total suspended solids
• Settleable solids
• Floating particulates
• Dissolved oxygen (Winkler method)
• Dissolved oxygen (Probe method)*
• Biochemical oxygen demand (BOO)
• Oil and grease
• Nitrogen (ammonia)*
• Nitrogen (total Kjeldahl)
• Nitrogen (nitrate and nitrite)
t Phosphorus (total)
t Total and fecal coliform bacteria
• Enterococcus bacteria
• Chlorophyll £
• Phytoplankton.
Samples to be analyzed for these variables generally will be collected
using water-bottle samplers. Those variables followed by an asterisk (*)
may also be measured using in situ instruments. Operation of water-bottle
samplers and in situ instruments are described in the preceding general
methods section.
119
-------
Receiving Water
PH
2H
Field Procedures--
Col 1ection--Samples for pH determination should be collected in poly-
ethylene or glass bottles with airtight screw caps. Because pH is unstable
and cannot be preserved, these samples should be collected immediately
after the sampler is brought on deck. Only dissolved oxygen samples should
be collected before pH samples. Because pH of waters not at equilibrium
with the atmosphere may change upon pxposure to the atmosphere, sample
containers should be completely filled and tightly sealed during collection.
Prior to sample collection, each sample bottle and cap should be rinsed
thoroughly with sample water. This can be achieved by filling the bottle
halfway, sealing and shaking it, and rinsing the stopper as the wash water
is discarded.
A piece of soft-walled rubber tubing should be attached to the outlet
valve of the sampler. This tubing should then be inserted to the bottom
of the sample bottle and at least one full volume allowed to overflow the
bottle. With the water still flowing, the tubing should be withdrawn slowly
from the sample bottle. Contamination of the sample with air bubbles should
be avoided.
After the tubing has been removed from the sample bottle, the stopper
should be put in place carefully to avoid trapping air bubbles. Once stoppered,
the sample should be checked for bubbles. If they are present, the sample
should be discarded, and a new one taken. The stopper on each accepted
sample should be double-checked to ensure a tight seal.
Process ing--Because pH cannot be preserved, samples should be analyzed
immediately after collection. If a short delay occurs, the samples should
be stored in the dark at 4° c and the storage time should be noted on the
log sheet.
120
-------
Receiving Water
PH
Laboratory Procedures--
Analytical Procedures--Ana1ytica1 procedures are given in U.S. EPA
Method 150.1 and APHA Method 423. As noted previously, pH samples should
be analyzed as soon as possible following collection.
Several potential sources of interference with pH measurements should
be avoided. Because the response of the electrode can be impaired if it
is coated with oily or iarticuiate material, the electrode should be gently
blotted 01- washed periodically with a detergent. Treatment with hydrochloric
acid may be necessary to remove some kinds of film. Temperature can influence
pH measurements by altering electrode output and changing the pH inherent
in the sample. The first source of temperature interference can be controlled
by using a pH meter with temperature compensation or by calibrating the
meter at the temperature of the samples. Because the second kind of temperature
interference cannot be controlled, the temperature at which the pH determination
of each sample is made should be logged and reported. It is recommended
that sample temperatures never differ by more than 2° C from that of the
buffer solution.
When pH measurements are being made, it is critical that the sample
be stirred at a constant rate to provide drift-free (<0.1 pH units) measure-
ments. The rate of stirring should minimize air transfer at the surface
of the sample. At least 30 sec should be allowed for each measurement
to stabilize.
Calibration and Preventive Maintenance--Calibration procedures for
the pH meter should follow specifications of the manufacturer. General
guidelines are given by U.S. EPA (1979b) and APHA (1985).
Buffer salts can be purchased as a solution that has been calibrated
to National Bureau of Standards salts. The pH meter should be calibrated
at a minimum of two points that bracket the expected pH value of the samples
121
-------
Receiving Water
pH
and that are three or more pH units apart (U.S. EPA 1979b). Prepare fresh
buffer solutions at least every month to avoid erroneous calibration as
a result of mold growth or contamination.
Preventive maintenance procedures should follow specifications given
by the manufacturer of the pH meter. In general, verification of electrode
performance and meter performance is the only operator service recommended.
An electrode should be replaced when it no longer meets span requirement
and does not improve with rejuvenating procedures.
Qua! ity Control Checks—The pH meter should be calibrated at the beginning
of each series of samples and after each group of 10 successive measurements.
It is recommended that duplicate pH determinations be made on at least
10 percent of the total number of samples. As an independent check, a
U.S. EPA reference sample should be analyzed at a minimum of every 3 mo.
Corrective Action—If the pH meter does not appear to be operating
correctly, consult the manufacturer's troubleshooting guide. Some common
problems include a dirty electrode, failure to fill the reference portion
of the electrode with internal solution, and inadequate stirring.
Data Quality and Report ing--A precision of +_ 0.02 pH unit and an accuracy
of 0.05 pH unit can be achieved under the best circumstances. However,
the limit of accuracy under most circumstances is ^0.1 pH unit (APHA 1985).
A precision of 0.1 pH unit is considered acceptable (U.S. EPA 1979b).
Measurements of pH are reliable only when the instrument has been
calibrated by standard buffers bracketing the desired range. Samples having
a pH greater than 10 may require a special probe to correct for "sodium"
error. However, pH values as high as 10 are not likely to be encountered
in most coastal waters.
It is recommended that pH values be reported to the nearest 0.1 unit.
In addition, the ambient temperature at the time of measurement of each
122
-------
Receiving Water
PH
sample should be reported to the nearest degree C. Results of all determina-
tions should be reported, including QA replicates. Any factors that may
have influenced sample quality should also be reported.
123
-------
Receiving Water
Salinity
Salinity
Field Procedures--
Col1ection--Two kinds of bottles are acceptable for collecting salinity
samples. The first is made of borosilicate glass and has an airtight septum
stopper. The second is made of low-pressure polyethylene and has polyethylene
inserts in the screw caps. Soft glass bottles, high-pressure polyethylene
bottles, and ground glass stoppers snould not be used.
Prior to sampling, the collection bottles should be "seasoned" by
filling them with seawater. Bottles should remain upside down in the case
until the sample is taken. Bottles with chipped edges or loose caps should
not be used.
Each1 collection bottle and cap should be rinsed at least three times
with sample water before the sample is collected. No salt crystals should
remain on the bottle or stopper. The bottle and stopper should not be
contaminated by contact with any surface. If contamination occurs, the
rinsing step should be repeated.
After the bottle and stopper have been rinsed thoroughly, the bottle
should be filled to approximately 90 percent of the bottle volume with
sample and sealed. The stopper should be double-checked for a tight fit.
The external label on each bottle should be filled out completely.
Processing—No reagents are necessary to preserve the salinity samples.
Bottles should be stored upright after samples have been collected. If
necessary, properly sealed salinity samples can be stored indefinitely
before analysis (Strickland and Parsons 1972).
124
-------
Receiving Water
Salinity
liooratory Procedures--
Analytical Procedures--The salinometer manufacturer should provide
a detailed description of how to use the instrument. To avoid heating
t^e sample bottle, it should oe propped up rather than gripped Dy hand
d-jring the salinity determination. The sample cell bowl should be filled
slowly to avoid introducing bubbles. The cell should be rinsed thoroughly
with distilled water between successive samples.
Calibration and Preventive Maintenance--Calibration prod lures should
follow the specifications given by the manufacturer of the salinometer.
It is recommended that the primary standard be Copenhagen seawater and
tnat all secondary standards be based on this primary standard. Secondary
standards should consist of filtered seawater collected from the open ocean
at a depth of at least 50 m (APHA 1985) and should be periodically checked
against the primary standard to guard against contamination or drift.
The secondary standards should be equilibrated to the temperature of the
samples before calibration begins. Secondary standards should be stored
in glass containers and protected from evaporation and light.
Preventive maintenance procedures should follow the salinometer manufac-
turer's recommendations. These include periodic cleaning of the sample
cell bowl, greasing the threads on the water-trap jar, tightening all water
connections, checking the temperature circuit calibration, and lubricating
tne pump, pump motor, and stirrer motor. It is critical that the sample
cell bowl be kept clean. Normally, the bowl should be cleaned daily.
However, if the sample water is very dirty, hourly cleaning may be necessary.
Quality Control Checks—Two standard sample determinations should
be made before the start of each series of samples. In addition, one standard
sample should be analyzed after each group of 10 successive samples, to
monitor instrument drift. It is recommended that duplicate determinations
be made for at least 10 percent of the samples analyzed.
125
-------
Receiving Water
Salinity
Corrective Action—If the salinometer does not appear to be operating
properly, the manufacturer's troubleshooting guide should be consulted.
Several common problems include failure of the null indicator to show a
deflection, failure of the thermometer circuit, excessive salinity balance
drift, inability to fill the sample cell completely, and failure of the
stirrer to operate properly.
Data Quality and Reporting — It is recommended that salinity determinations
be made using an induction salinometer. Other, less common procedures
include the hydrometric and argentometric methods see Methods 210B and
210C Of APHA 1985).
A precision of +0.1 ppt is possible using an induction salinometer.
Conductivity measurements should be converted to salinity values using
standard tables corrected for temperature. If possible, salinity concentrations
should be reported in ppt to the nearest 0.01 unit. Results of all deter-
minations should be reported, including QA replicates and standards. Any
factors that may have influenced sample quality should also be reported.
126
-------
Receiving Water
Temperature
Temperature
Field Procedures--
Col 1 ection--Temperature can be measured using a mercury-filled Celsius
thermometer on samoles collected in glass or plastic containers. The ther-
mometer should have a scale etched on capillary glass for 0.1° C increments
and a minimal thermal capacity to permit rapid equi1 ibration. Temperature
can be measured -in situ using a reversing thermometer or a thermistor.
Of these two _i£ situ instruments, the then-istor is more accurate, but
also more expensive.
Processing—Because temperature cen change rapidly after a sample
is removed from ambient conditions, temperature determinations by thermometer
should be made immediately after sample collection.
Laboratory Procedures—
Analytlcal Procedures—Methods for making temperature measurements
are described in U.S. EPA Method 170.1 and in APHA Method 212. It is critical
that the measuring device be adequately immersed in the sample and allowed
to completely equilibrate (i.e., the temperature reading stabilizes) before
temperature is determined.
Calibration and Preventive Maintenance—Each kind of temperature-measuring
instrument should be calibrated frequently against a National Bureau of
Standards (NBS) certified thermometer that is used with its certificate
and correction chart. An NBS thermometer is recommended because some commercial
thermometers may be as much as 30 C in error (APHA 1985).
To prevent breakage, it is recommended that each thermometer be enclosed
in a metal case. If a mercury thermometer is broken, samples or containers
in the vicinity of the exposed area may be contaminated by the mercury.
127
-------
Receiving Water
Temperature
Quality Control Checks—Each temperature-measuring instrument should
be calibrated against a N8S standard thermometer at least every week.
It is recommended that calibration be conducted daily when a temperature
violation is suspected.
Corrective Action--If the temperature-measuring instrument cannot
be calibrated consistently against the N8S thermometer, it should be repaired
or replaced.
Data Quality and Reporting--Pre. ision and accuracy have not been determined
for temperature measurements (U.S. EPA 1979b). If possible, temperature
measurements should be reported to the nearest O.Olo C. Results of all
determinations should be reported, including QA replicates and standard
checks. Any factors that may have influenced sample quality should also
be reported.
128
-------
Receiving Water
Color
Color
Field Procedures—
Col lection—Color samples shoula be collected from the water sampler
In glass or plastic bottles. Bottles should be cleaned thoroughly before
use. Because biological activity may change color characteristics, samples
should be coMected and analyzea as soon as possible after the sampler
is Drought on board. It is recommended that color samples be collected
immediately after dissolved r tygen and pH samples.
Processing—Because no reagents can be added to preserve color, samples
should be analyzed as soon as possible after collection. If a delay occurs,
samples should be held at 4o c for no more than 24 h. The length of delay
should be recorded on the log sheet.
Laboratory Procedures—
Analytical Procedures—Color should be determined using the spectrophoto-
metric methods described in U.S. EPA Method 110.3 and in APHA Method 204B.
The recommended method of color analysis is very sensitive to turbidity.
However, the optimum filter media to remove turbidity without removing
color has not been found. Recommended methods include filtration through
a 0.45-um filter and centrifugation. The spectrophotometric method is
also sensitive to sample pH.
Calibration and Preventive Maintenance—For the spectrophotometric
method, the instrument should be set to read 100-percent transmittance
on a distilled water blank. All determinations should be made within a
narrow spectral band.
Preventive maintenance procedures for the spectrophotometer shoula
follow the manufacturer's recommendations.
129
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Receiving Water
Color
Quality Control Checks — It is recommended that duplicate color deter-
minations be made on at least 10 percent of the total samples analyzed.
Corrective Action — If the spectrophotometer does not appear to be
operating properly, the manufacturer's troubleshooting guide should be
consulted.
Data Quality and Reporting—A: present, precision and accuracy data
are not avail t'.e for the spectrophotometric method (U.S.- EPA 1979b) .
"esults of th'S method should be reported at pH 7.6 and at the origina"1
pH in terms of dominant wavelength hue (to the nearest 1.0 nm), luminance
(to nearest 0.1 percent), and purity (to nearest 1.0 percent). In addition,
the kind of spectrophotometer, number of selected ordinates (10 or 30),
and the spectral band width (nm) should also be reported. Results of all
determinations should be reported, including QA replicates. Any factors
that may have influenced sample quality should also be reported.
130
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Receiving Water
Transparency
Transparency
Transparency is measured using a Secchi disk. This device is a circular
plate with a standard diameter of 30 cm (U.S. Navy 1968). The top side
of the disk should be white. A ring attached at the center of the disk
allows a graduated line to be secured. A 2-4 kg weight should be attached
centrally to the underside of the disk to ensure that the device will sink
rapidly and vertically. The deployment line should be made of material
that will not stretch substantially after repeated use (e.g., braided dacron).
Transparency measurements should be made by Ijwering the Sec;hi disk
from the shaded side of the survey vessel until the disk is barely perceptible.
This depth should be recorded to the nearest 0.5 m and the disk should
continue to be lowered until it is no longer visible. The disk should
then be raised slowly until it is again barely visible.. This second depth
should also be recorded. The average of the two depth readings (i.e.,
downward and upward) should be reported to the nearest 0.5 m as the measured
transparency value.
Because Secchi disk readings are dependent upon the available illumination,
they vary with time of day, cloud formation, and cloud cover. Secchi disk
readings also vary with the observer because of interpersonal differences
in visual ability. Thus, to standardize these readings, repeated measurements
should be made by one Individual under similar conditions of illumination.
Because these criteria are not always achievable, associated meteorological
data at the time of measurement and the name of the person making the deter-
minations should be included on the log sheet with the Secchi disk readings.
131
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Receiving Water
Turbidity
Turbidity
Field Procedures —
Col lection—Turbidity samples can be collected in glass or plastic
containers. Samples should be removed from the sampler as soon as possible
after collection to minimize settling of suspended material within the
sampler. Sample containers and lids should be rinsed thoroughly with sample
water before samples are collected. Turbidity can be estimated in situ
as transmissivity using a transmissometer. A discussion of such . • situ
instrumentation is provided earlier in the general methods section.
Processing—Because turbidity samples cannot be preserved adequately,
they should be analyzed as soon as possible after collection. If a delay
occurs, samples should be held at 40 C for no more than 48 h to minimize
microbiological decomposition of solids. The length of delay should be
noted on the log sheet.
Laboratory Procedures—
Analytical Procedures—The nephelometric method is described in U.S. EPA
Method 180.1 and in APHA Method 214A. For turbidities greater than 40 NTU,
samples should be diluted with one or more volumes of turbidity-free water
until the turbidity falls below 40 NTU. Turbidity-free water is distilled
water passed through a membrane filter with a 0.2-um pore size. Samples
should be shaken well to thoroughly disperse solids, and resulting air
bubbles should be allowed to dissipate before the sample is analyzed.
Interference with turbidity measurements arises from several sources.
Because the presence of floating debris and coarse sediments that settle
out rapidly will give low readings, readings should be made as soon as
possible after sample agitation. Finely divided air bubbles will affect
results in a positive direction. If present, the bubbles should be allowed
to dissipate before a reading is made. Finally, dissolved substances that
132
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Receiving Water
Turbidity
absorb light in the sample will reduce turbidity readings. Any color in
the sample should therefore be noted.
Calibration and Preventive Maintenance—It is recommended that a standard
suspension of formazin be used to calibrate the nephelometer. Formazin
provides a more reproducible turbidity standard than do other materials
used in the past. The formazin standard suspension should be prepared
daily (APHA 1985). Commercially available standards such as styrene divinyl-
benzene beads (trade name AMCO-AEPA-1), can be substituted for formazin
if they ar«- demonstrated to be equivalent to freshly prepared formazin
(APHA 1985). Standards measured on the nephelometer should cover the rar.c-e
expected for the samples. At least one standard should be run in each
instrument range to be used. The instrument should give stable readings
in all sensitivity ranges used. If a precal ibrated scale is not supplied,
calibration curves should be prepared for each instrument range to be used.
Quality Control Checks — The nephelometer should be calibrated at the
start of each series of analyses and after each group of 10 successive
samples. Duplicate analyses should be conducted on at least 10 percent
of the total number of samples.
Corrective Action—If the nephelometer will not stabilize in any of
the relevant ranges or if the instrument does not appear to be functioning
properly in any other aspect, the manufacturer's troubleshooting guide
should be consulted. Sample tubes that become scratched or etched should
be replaced.
Data Quality and Reporting—Because the nephelometric method of turbidity
measurement is more sensitive than the visual method, the former is recom-
mended. Limited precision data indicate that standard deviations of measure-
ments vary directly with the level of turbidity (U.S. EPA 1979b). Accuracy
data are not available at present. The sensitivity of the nephelometer
should allow detection of a turbidity difference of 0.02 unit or less in
waters with turbidities less than 1.0 unit. Results should be reported
133
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Receiving Water
Turbidity
in nephelometric turbidity units (NTU). U.S. EPA Method 180.1 describes
the nearest reporting units as a function of the raige of values measured.
Results of all determinations should be reported, including QA replicates.
Any factors that may have influenced sample quality should also be reported.
134
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Receiving Water
Total Suspended Solid-?
Total Suspended Solids (TSS)
Field Procedures—
Collection—Samples should be collected in glass or plastic bottles.
Samples should be collected soon after the sampler is brought on board
to minimize settling of suspended material within the sampler. Nonrepresen-
tative particulates such as leaves and sticks should be noted, and then
excluded from the sample.
Processing—Total suspended solids samples cannot be preserved adequately
and should therefore be analyzed as soon as possible after collection (APHA
1985). If a delay occurs, samples should be held at 40 C to minimize micro-
biological decomposition of solids. The length of delay should not exceed
7 days and should be noted on the log sheet.
Laboratory Procedures—
Analytical Procedures—Suspended solids determination should be made
according to procedures described in U.S. EPA Method 160.2 and APHA Method
209C. However, a 0.40- or 0.45-um membrane filter should be used to remove
suspended solids instead of the glass fiber filter specified in the U.S. EPA
and APHA methods. These filters are the ones used most commonly, for oceano-
graphic work in coastal waters.
The drying temperature of the filtered residue can influence results
because temperature and time of heating affect weight losses due to volatili-
zation of organic matter, mechanically occluded water, water of crystallization,
and gasses from heat-induced chemical decomposition, as well as weight
gains due to oxidation (APHA 1985). Thus, drying temperature must be carefully
controlled and not allowed to deviate from the reccrrmended range of 103-1050 C.
To avoid contamination, filters should be handled with forceps during
all steps from initial to final weight determinations. Filters should
135
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Receiving Water
Total Suspended So>ids
a" ways be stored in a desiccator when cooling. When filtering the samples,
it is critical that the filter is seated tightly on the surface of the
filtration apparatus and that all holes in the crucible are covered. To
ensure complete removal of salts after filtering the sample, the filter
snould be rinsed with a minimum of three successive 20-nt portions of distilled
water. It is recommended that enough water be filtered to ensure that
at least 5 mg of residue is collected. Because excessive residue on the
filter rjiay form a water-entrapping crust, the sample size should be limited
to that which yields less than 200 mg of residue (APHA 1985). Prolonged
filtration times resulting -on filter clogging may produce high results
due to excessive solids cap':ure on the clogged filter. Therefore filtering
should be terminated before any evidence of clogging is noted.
Calibration and Preventive Maintenance—The analytical balance should
be calibrated weekly using standard weights according to the manufacturer's
instructions. It is recommended that theibalance have a minimum accuracy
of 0.1 mg. The manufacturer's preventive maintenance procedures should
be followed carefully.
Quality Control Checks—For each weight determination, filters should
be run repeatedly through the drying/cooling cycle until the weight loss
is less than 4 percent of the previous weight or 0.5 mg, whichever is less
(APHA 1985). Duplicate analyses should be conducted on at least 10 percent
of the total number of samples. A filter blank should be taken through
the preparation, drying, and desiccation steps for each batch of samples
to monitor changes in filter tare weight. U.S. EPA reference samples should
be analyzed quarterly to check the overall accuracy of the method.
Corrective Action—If the analytical balance will not produce repeatable
measurements within 0.1 ng, the manufacturer's troubleshooting guide should
be consulted. If the filter becomes clogged during filtration, it should
be discarded and the analysis should be repeated using a clean filter.
To prevent clogging of the second filter, the volume of sample analyzed
should be reduced.
136
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Receiving Water
Total Suspended Solids
If the weight of the filter blank is not consistent after drying and
cooling, the desiccant should be checked. A color-indicating desiccant
is recommended, so that spent desiccant is easily detected. Also, the
seal on the desiccator should be checked and, if necessary, ground glass
rims should oe greased or "0" rings replaced. Cooling time should be closely
monitored, so weighing times are consistent between batches of samples,
thereby minimizing the need for multiple weighings.
Data Quality and Reporting—Precision of res- its varies directly with
the concentration of suspended matter and, at lo/ levels, the ratio of
the weight of the suspended matter to the weight of the filter. There
are no procedures for determining the accuracy of field measurements of
suspended matter. Total suspended solids measurements should be reported
as mg/L to a minimum of two significant figures. Results of all determinations
should be reported including QA replicates and filter blanks. Any factors
that may have influenced sample quality should also be reported.
137
-------
Receiving Water
Settleable Solids
Settleable Solids
Field Procedures—
Col 1 ect ion—Settleable solids samples should be collected in 1-L glass
or plastic containers. The sample container selected for use should be
cnecked to ensure that material in suspension does not adhere to the container
walls. Samples should be collected soon after the sampler is brought on
board, to minimize settlement of suspended material within the sampler.
Nonreprf ;ent?tive particulates such as leaves and sticks should be noted
and then excluded from the sample.
Process ing--Settleable solids samples cannot be preserved adequately
and should therefore be analyzed as soon as possible after collection (APHA
1985). If a delay occurs, samples should be held at 40 C to minimize micro-
biological decomposition of solids. The length of delay should not exceed
48 h and should be noted on the log sheet.
Laboratory Procedures—
Analytical Procedures—Procedures used to determine settleable solids
concentrations are presented in U.S. EPA Method 160.5 and in APHA Method
209E. The sample should be well-mixed before introduction to the Imhoff
cone. It is critical that floating material is not included with settleable
material.
Calibration and Preventive Maintenance—Not applicable.
Quality Control Checks--Duplicate analyses should be conducted on
at least 10 percent of the total number of samples.
Corrective Actions—If suspended material is found to adhere to the
sides of the sample collection containers, a different type of container
should be used.
138
-------
Receiving Water
Settleable Solids
Gata Quality and Reporting — The practical lower limit of measurement
depends on sample composition and generally is in the range of 0.1 to 1.0 mL/L
(APHA 1985). Precision and accuracy data are not available at present.
Settleable solids concentrations should be reported as mL/L to a minimum
of two significant figures. Results should be reported for all determinations,
including QA replicates. Any factors that may have influenced sample quality
should also be reported.
139
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Receiving Water
Floating Participates
F". aati-iq Particulates
Field Procedures—
Col lee tion—A minimum of 5 L of sample snould be collected in a glass
or plastic container. The container should be rinsea thoroughly with sample
water before sample collection.
Processing — For comparable results, samples must be treated uniformly
throughout sampling and handling. Analyses should be conduct-d as soon
as possible after sample collection.
Laboratory Procedures--
Analytical Procedures—Floating particulates should be analyzed in
accordance with APHA Method 206A. At present, this method is tentatively
recommended by APHA. Because even slight differences in sampling and handling
can give large differences in measureable floating particulates, all samples
should be treated uniformly, preferably by adequately mixing them before
flotation. When mixing the sample, care should be taken to avoid extensive
air entrapment through formation of a large vortex. Because temperature
variations can affect results, all tests should be conducted at a constant
temperature, preferably 20° C.
Calibration and Preventive Maintenance—No calibration procedures
apply to the recommended method. To prevent oil and grease from sticking
to the analytical equipment, all internal surfaces should be coated with
TFE.
Quality Control Checks —It is recommended that duplicate analyses
be conducted on a minimum of 5 percent of the total number of samples,
with an additional 5 percent of the samples checked for recovery.
140
-------
Receiving Water
Floating Participates
Corrective Actions—If oil and grease appear to be sticking to surfaces,
TFE coatings should be renewed. If recovery jrops below 90 percent, samples
should be reanalyzed and each step of the analysis carefully scrutinized.
Data Quality and Reporting—The minimum detectable concentration of
floating particulates using the recommended method is 1 mg/L. Precision
varies with the concentration of suspended matter in the sample. A coefficient
of variation of 5.7 percent has been achieved using five replicate samples
(APHA 1985). Although there is no completely satisfactory procedure for
determining thp iccuracy of the method, approximate recovery can be determined
oy running a .econd test for floatables on all water discharged throughout
the analytical procedures, except for the last 10 ml. Typical recoveries
exceed 90 percent (APHA 1985). Concentrations of floating particulates
should be reported in mg/L to the nearest 0.1 unit. Results of all determi-
nations should be reported, including QA replicates and recovery checks.
Any factors that may have influenced sample quality shou.ld also be reported.
141
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Receiving Water
Dissolved Oxygen (Winkler)
Dissolved Oxygen (Winkler Method)
Field Procedures--
Col1ection--Prior to sample collection, the fixing reagents should
be prepared and the dispensing aoparatus should be filled. The accuracy
of the volumes being dispensed should be checked and no air should be trapped
in the system. It is recc-mended that 300-mL glass BOO bottles with ground
glass stoppers be used for the Winkler method.
Oxygen samples must be :he first ones collected from the sampler and
they should be collected immediately after the sampler is brought on board.
It is recommended that a piece of soft-walled rubber tubing be connected
to the discharge valve of the sampler to prevent air bubbles from contaminating
the sample during collection. The tubing should be .soaked in seawater
prior to use to prevent air bubbles from collecting inside.
After being attached to the sampler, the tubing should be flushed
with sample water to remove air bubbles. The sample bottle and stopper
should then be rinsed thoroughly with sample water. After rinsing, the
tubing should be inserted to the bottom of the sampling bottle. The bottle
should be filled slowly until at least half full, and then filled rapidly
thereafter. At least one full bottle volume of sample should overflow
the bottle before the tubing is removed. After the tubing is removed slowly,
the stopper should be carefully put in place with a twisting motion while
water is displaced from the bottle. Once stoppered, the sample should
be checked for air bubbles. If bubbles are present, the sample should
be discarded and a new sample collected. Acceptable samples should be
fixed as soon as possible after collection.
Processing—The stopper should be carefully removed from the bottle
without agitating the sample. Each fixing reagent should be added by gently
placing the tip of the pi pet slightly below the surface of the sample and
gently pushing the plunger. The plunger should not be released until the
142
-------
Receiving Water
Dissolved Oxygen (Winkler)
pi pet has been removed from the sample. The pipet tip should be rinsed
with distilled water before being returned to the reagent bottle.
After the fixing reagents have been added, the bottle should be carefully
stoppered without introducing air bubbles. Excess fluid around the outside
of the stopper should be poured off and the sample bottle should be inverted
5-10 times to thoroughly disperse the precipitate. It is also good practice
to invert bottles several times approximately 20 min after fixation to
ensure thorough dispersion of the precipitate.
After allowing the precipitite to settle for 10-15 min, the stopper
should be removed and sulfuric acid should be added to the sample in the
same manner as the fixing reagents. The stopper should then be replaced
and the bottle inverted until all of the precipitate has dissolved. If
the precipitate fails to dissolve, it should be allowed to settle again
and additional sulfuric acid should be added to the sample. It is critical
that all of the precipitate be dissolved before samples are stored. Also,
it is critical that samples not be allowed to stand longer than 8 h before
sulfuric acid is added, as erroneous measurements may result.
Preserved dissolved oxygen samples should be stored in the dark at
10-200 c. Samples should be analyzed as soon as possible after collection,
and storage time should not exceed 8 h. The length of storage should be
recorded on the log sheet.
Laboratory Procedures--
Analytical Procedures—The recommended modified Winkler method for
saline water is described in detail in Strickland and Parsons (1972).
Calibration and Preventive Maintenance—Methods of standardizing the
thiosulfate solution are presented by Strickland and Parsons (1972). It
is recommended that one person perform the standard and sample titrations
because of subjectivity in the color of the endpoint.
143
-------
Receiving Water
Dissolved Oxygen (Winkler)
Preventive maintenance is limited to ensuring that reagent disoensing
and titrating equipment is clean and functions properly.
Quality Control Checks—All standard titrations should be duplicated.
It is recommended that duplicate analyses be conducted on at least 10 percent
of the -otal samples. Four replicate reagent blanks should be run once
a week during a cruise, or whenever a reagent is changed.
Corrective Action--! the results obtained by running duplicate standard
titrations of the thiosulfate solution do not agree within ^0.05 ml, the
titrations should be repeated until agreement is achieved. All reagent
dispensers should be checked for bubbles, and the amounts of reagents delivered
should be verified.
Data Quality and Reporting—Using the modified Winkler method, repro-
ducibility for field samples is approximately 0.2 mg/L of dissolved oxygen
at the 7.5 mg/L level (U.S. EPA 1979b). Duplicate titrations made during
standardization of reagents should agree within ^0.05 ml. With careful
collection and treatment of samples, dissolved oxygen as low as 1 percent
of saturation can be measured. Dissolved oxygen concentrations should
be reported in mg/L to the nearest 0.1 unit. Results should be reported
for all determinations, including QA replicates and reagent blanks. Any
factors that may have influenced sample quality should also be reported.
144
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Receiving Water
Dissolved Oxygen (Probe)
Dissolved Oxygen (Probe Method)
Field Procedures--
Conection--0xygen samples should be the first ones collected from
the sampler and they should be collected immediately after the sampler
is brought on board. It is recommended that a piece of soft-walled rubber
tubing be connected to the aischarge valve of the sampler to prevent air
bubbles from contaminating the sample during collection. The tubing should
be soaked in seawater prior to use to prevent air bu». >les from collecting
inside.
After being attached to the sampler, the plastic or rubber tubing
should be flushed with sample water to remove air bubbles. The sample
bottle and stopper should then be rinsed thoroughly with sample water.
After rinsing, the tubing should be inserted to the bottom of the sampling
bottle. The bottle should be filled slowly until at least half full, and
then filled rapidly thereafter. At least one full bottle volume of sample
should overflow the bottle before the tubing is removed. After the tubing
is slowly removed, the stopper should be carefully put in place with a
twisting motion while water is displaced from the bottle. Once stoppered,
the sample should be checked for air bubbles. If bubbles are present,
the sample should be discarded and a new sample collected.
Processing—Because no reagents are used to preserve the oxygen samples,
analyses should be conducted Immediately after collection. If a delay
occurs, it should be noted on the log sheet.
Laboratory Procedures—
Analytical Procedures—Detailed analytical procedures should be provided
by the manufacturer of the dissolved oxygen meter. General procedures
are listed in U.S. EPA Method 360.1 and APHA Method 421F.
145
-------
Receiving Water
Dissolved Oxygen (Probe)
Several precautions should be taken when making measurements with
a membrane electrode. First, constant turbulence should be provided by
a stirrer to ensure precise measurements. Second, adequate time should
be allowed for the instrument to warm up before measurements are started
and, when individual samples are analyzed, for the probe to stabilize to
sample temperature and dissolved oxygen. Third, reactive gases, such as
chlorine and hydrogen sulfide, pass through the -"lembrane probes and may
interfere with the analysis or desensitize the probe. Finally, broad variations
in th kinds and concentrations of salts in samples can influence the partial
press re of oxygen in samples and thereby affect measurement accurscy.
Calibration and Preventive Maintenance—Calibration procedures should
follow the instructions given by the manufacturer of the dissolved oxygen
meter. The meter generally can be calibrated using one of three methods:
Winkler titrati'on, saturated water, or air. The air method is simplest
and quite reliable. Overall error is diminished when the probe and instrument
are calibrated under conditions of temperature and dissolved oxygen that
match those of the samples. Calibration can be disturbed by physical shock,
touching the membrane, or desiccation of the electrolyte.
Preventive maintenance procedures should follow the manufacturer1 s
reconnendations. The oxygen probe should always be stored in a humid environ-
ment to prevent drying out and the need to frequently replace membranes.
Quality Control Checks—The instrument should be calibrated at the
beginning of each series of measurements and after each group of 10 successive
samples. Duplicate measurements should be made on at least 10 percent
of the total number of samples.
Corrective Action--If the dissolved oxygen meter does not appear to
be operating correctly, consult the manufacturer's troubleshooting guidelines
for remedial actions.
146
-------
Receiving Water
Dissolved Oxygen (Probe)
Data Quality and Reporting—Repeatability of dissolved oxygen measurements
using a membrane electrode should be 0.1 mg/L and accuracy should be +1 percent
(J.S. EPA 1979b). Sensitivity of the electronic readout meter for the
output from the dissolved oxygen probes should normally be 0.05 mg/L (U.S. EPA
1979b). Dissolved oxygen concentrations should be reported in mg/L to
tne nearest 0.1 unit. Results should be reported for all determinations,
including QA replicates. Any factors that may have influenced sample qual ity
snould also be reported.
147
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Receiving Water
BOD
Biochemical Oxygen Demand (BOD)
Field Procedures--
Conection--BOD samples can be collected in glass or plastic containers.
Sample containers and caps should be rinsed thoroughly with sample water
before sample collection.
3rocessing--BOD samples should be analyzed immediately after collection.
If a delay occurs, samples should be held at 40 c to min lize reduction
of BOO. Samples should not be stored for more than 48 h and the length
of storage should be recorded on the log sheet. Refrigerated samples should
be warmed to 2DO C prior to analysis.
Laboratory Procedures--
Analytical Procedures--BQD concentrations should be determined according
to U.S. EPA Method 405.1 or APHA Method 507. Generally seawater samples
require no dilution. Samples having more oxygen-demanding materials than
the amount of oxygen in air-saturated water should be diluted to balance
the oxygen demand and supply. If samples are diluted, nutrient addition
(i.e., nitrogen, phosphorus, trace metals) and pH buffering of the dilution
water are necessary to ensure that the sample is suitable for bacterial
growth. To prevent air from infiltrating the incubation bottles, a water
seal should be used. When samples are incubating, all light should be
excluded to prevent photosynthetic production of oxygen. Samples containing
residual chlorine must be dechlorinated (e.g., using sodium thiosulfate).
Calibration and Preventive Maintenance—Dissolved oxygen concentrations
should be measured on all dilution water blanks and seed controls if appli-
cable. Generally, seawater samples require no dilution of seed. Therefore
to monitor the method, a glucose-gultamic acid standard check solution
should be incubated with each batch of samples. Dissolved oxygen measurements
148
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Receiving Water
BOO
snould be calibrated according to accepted procedures (e.g., see descriptions
of the Winkler and probe methods in this document).
Quality Control Checks—The dilution water blank and the glucose-glutamic
acid standard provide quality control on the dilution water as well as
on the cleanliness of analytical equipment (e.g., incubation bottles).
Each sample should be analyzed in triplicate to monitor precision. Optimal
uatake is at least 2 mg/L after the incubation, with a residual-oxygen
concentration of at least 1 mg/L in the sample. Duplicate analyses should
be conducted on a* '-east 10 percent of the total number of samples.
Corrective Action--APHA (1985) should be consulted for methods of
correcting the many kinds of interference that may accompany BOO analyses.
If the dilution water blanks exceed 0.2 mg/L, cleanliness of containers
and water should be checked. Containers may require 1+1 HC1 rinse after
detergent washing to remove any residual organic material. Any containers
rinsed with acid should be thoroughly rinsed with distilled water to prevent
acid carryover.
If a 2-percent dilution of the glucose-glutamic acid standard check
solution is outside the range of 200 +. 37 mg/L, BOD determinations made
with the seed and dilution water should be rejected. Several methods used
to determine the problem include running a series of dilution water blanks
using different water sources with and without seed, preparing a fresh
solution of glucose-glutamic acid, changing the seed, or preparing fresh
reagents for the dilution water. The source of the problem should be determined
before additional BOD analyses are performed.
Data Quality and Reporting — Precision data for spiked natural waters
indicate that standard deviations of +0.7 and j+26 can be achieved for BOO
concentrations of 2.1 and 175 mg/L, respectively (U.S. EPA 1979b). There
is no acceptable method for determining the accuracy of the BOD test.
BOD data should be reported as mg/L to the nearest 0.1 unit. Results should
149
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Receiving Water
800
be reported for all determinations, including QA replicates, dilution water
clanks, and glucost-gl utamic acid standards. Any factors that nay have
influenced sample q-jal ity should also be reported.
150
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Receiving Water
Oil and Grease
Oil and Grease
Field Procedures--
Conection--Samp1es should be collected in glass bottles. Bottles
should first be washed with a warm aqueous detergent mixture, and then,
in sequence, thoroughly rinsed with hot tap water, rinsed at least twice
with distilled water, rinsed witn 1,1,2-trichloro-l,2,2-trifluoroethane
(i.e., Freon or equivalent), ard dried in a clean oven at ^i05o C for 30
min. Bottle caps should be lined with ' :E-coated cardboard inserts or aluminum
foil. Plastic containers are not acceptable. Headspace should be left
in each sample container for addition of acid and mixing.
Processing—Acidify the sample in the collection bottle to a pH <2
using sulfuric or hydrochloric acid. Samples should be stored in the dark
at 40 c. Recommended maximum holding time is 28 days (U.S. EPA 1984).
The length of storage should be recorded on the log sheet.
Laboratory Procedures--
Analytical Procedures--0il and grease should be measured according
to U.S. EPA Methods 413.1 and 413.2 or APHA Methods 503A and 5038. Because
asphaltic materials are insoluble in Freon, recommended methods will give
low recoveries for samples containing such material. The gravimetric method
for oil and grease measures relatively nonvolatile hydrocarbons, vegetable
oils, animal fats, waxes, and soaps. The method is often used for wastewater
analyses because it requires minimal instrumentation and calibration.
Sulfur causes interference for the gravimetric method because it will be
extracted and included as oil and grease. Light hydrocarbons that volatilize
at temperatures below 70<> C (e.g., gasoline through No. 2 fuel oil) are
lost during the solvent removal step.
The infrared oil and grease procedure is more precise at lower concen-
trations (1-10 mg/L) than the gravimetric oil and grease procedure. The
151
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Receiving Water
Oil and Grease
infrared method will give higher results than the gravimetric method if
tne sample contains volatile components, and will give lower results if
t-.e sample is high in sulfur content. A limitation to the infrared determi-
nation is the method of standardization, which requires a prepared reference
oil. The reference oil may not be comparable to the type of oil and grease
in the samples, resulting in inaccurate results.
Calibration and Preventive Maintenance—For gravimetric oil and grease
analyses, check accuracy of the analytical balance periodically (minimum
of once per week rec nmended) using Class S weights. A service contract
tnat includes schedulec preventive maintenance at least once per year is
recommended. Scratched, chipped, or cracked boiling flasks should be replaced.
For infrared oil and grease analyses, follow the manufacturer1 s preventive
maintenance procedures for the infrared spectrophotometer. Cells used
for analysis should be checked for scratches each time they are used.
Scratched cells should not be used. For infrared analyses, calibration
should be performed as specified in U.S. EPA Method 413.2, Section 6.4.
Quality Control Checks--Duplicate samples should be collected and
performed at a minimum of every 10 samples to establish an estimate of
precision. Because samples should not be split after collection, separate
grab samples should be taken for analysis.
Distilled water spiked with a U.S. EPA performance sample or reference
oil should be extracted and analyzed every 20 samples to monitor recovery.
Bottles should be checked for cleanliness by analyzing distilled water
that has been acidified in a sample bottle. A solvent blank should accompany
each batch of samples. A procedural blank should be run with each batch
of samples to monitor reagent contamination or procedural problems.
Corrective Action—If oil and grease concentrations in procedural
blanks are greater than the detection limit, check the cleanliness of all
glassware. Always use separatory funnels with TFE stopcocks to avoid contam-
ination from stopcock grease. For infrared analyses, severe interferences
152
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Receiving Water
Oil and Grease
will result if Freon contacts any material containing plasticizers (e.g.,
plastic tubing). For gravimetric analyses, high results will be obtained
if any Freon or fumes remain in the flask after distillation.
If difficulty occurs with emulsions during solvent extraction, follow
procedures described in the methods. If the emulsion still fails to dissipate
after addition of salts, gently turn the separatory funnel to the horizontal
position and slowly rotate. Be careful to keep the stopper securely in
place.
If precision or recovery from spike results is p or, check adequacy
of extraction by increasing shaking time.
Data Quality and Reporting--The definition of oil and grease is based
on the procedures used. Unless identical procedures are used, oil and
grease determinations are not intercomparable. Therefore, the method used
for analysis should always be specified.
Objectives for precision and accuracy are indicated below:
0 Reproducibility for the gravimetric method using distilled
water spikes is ^18-percent coefficient of variation (U.S. EPA
1983). These results probably overestimate the precision
possible with effluent samples. No precision data are reported
by the U.S. EPA for the infrared method.
• Inter laboratory recovery results were 102+37 percent at
6.0 mg/L and 97+35 percent at 18.0 mg/L for the gravimetric
method (U.S. EPA 1983).
Measurements should be reported to a minimum of two significant figures
in mg/L. Detection limits are in the range of 5 mg/L for gravimetric oil
and grease determinations (U.S. EPA Method 413.1), and 0.2 mg/L for infrared
determinations (U.S. EPA Method 413.2). Results of all determinations
153
-------
Receiving Water
Oil and Grease
should be reported, including QA replicates, blanks, spiked samples, and
reference measurements. Any factors that may have influenced sample quality
should be reported.
154
-------
Receiving Water
Nitrogen (Ammonia)
N'trogen (Ammonia)
Field Procedures--
Conection--Ammonia samples can be collected in glass or plastic bottles.
Each bottle and cap should be rinsed thoroughly with sample water prior
to sample collection.
Processing--Resuits of ammonia analyses are most reliable when they
.•-e made on fresh samples. However, if analysis must ae delayed, samples
can be stored for up to 28 days by acidification to pH<2 with sulfuric
acid and refrigeration at 40 C. The length of delay before analysis should
be recorded on the log sheet.
Laboratory Procedures--
Analytical Procedures—Manual distillation of effluent samples prior
to ammonia determinations is required by the U.S. EPA, unless data on file
demonstrate that distillation 1s not required. Distillation is recommended
due to the sensitivity of amnonia procedures to color and possible interferences
in the effluent. Following distillation at pH 9.5, ammonia concentration
in saline water can be determined by U.S. EPA Methods 350.1, 350.2, or
350.3 or APHA Methods 417A, 417B, 4170, or 417G. The methods should be
consulted to determine which is most appropriate for available equipment,
expected concentrations, and expected levels of precision and accuracy.
Calibration and Preventive Maintenance—Calibration procedures should
follow those specified in the method. If samples are being distilled,
standards should also be distilled prior to analysis to check for ammonia
contamination or loss during processing. Concentrations of the calibration
standards should bracket the sample concentrations. If a sample concentration
is outside the range of calibration, then an additional calibration standard
should be analyzed to check if the result is within the linear range of
155
-------
Receiving Water
Nitrogen (Ammonia)
t.-e method. Alternatively, the sample should be diluted to within the
calibration range and then reanalyzed.
Quality Control Checks--Duplicate analyses should be conducted on
a minimum of 5 percent of the total number of samples, with an additional
5 percent of the samples spiked and analyzed for percent recovery. A blank
snould be analyzed with each batch of samples. A U.S. EPA performance
sample should be analyzed at least once per quarter.
Corrective Action—Contamination of ammoma sample? can recur easily
due to the volatile nature of ammonia. To prevent possible cro s-contamination,
reagents used for other analyses that contain ammonia (e.g., colorimetric
phenol) should be Isolated from samples and standards used for ammonia
determinations. In addition, cleaning preparations that contain significant
quantities of ammonia (e.g., Pinesol, wax removers) should not be used
in the laboratory area where ammonia determinations are performed.
Contaminated glassware should be rinsed with 1+1 HC1 and then with
distilled water. To check for contamination, blanks should be analyzed
whenever a new reagent is prepared.
Data Quality and Reporting—Detection and accurate quantification
of ammonia in receiving water is routinely attainable, although method
detection limits can vary widely because of methods or instrumentation.
The analytical method should be consulted to determine expected detection
limits, precision, and accuracy. Data should be reported in mg/L as N
to a maximum of three significant figures. Results should be reported
•or all determinations, including QA replicates and spiked samples. Any
factors that may have influenced sample quality should also be recorded.
156
-------
Receiving Water
Nitrogen (Total Kjeldah!)
Nitrogen (Total Kjeldahl)
Field Procedures--
Collection--Kjeldah! nitrogen samples can be collected in glass or
plastic containers. Caps should be unlined, as paper and/or glue may interfere
with the analysis. Each container and cap should be rinsed thoroughly
with sample water prior to sample collection.
Processing--I1 possible, samples should be analyzed immediately after
collection. If immediate analysis is not possible, samples can be stored
up to 28 days by acidification to pH<2 with sulfuric acid and refrigeration
at 40 C. The length of delay before analysis should be recorded on the
log sheet.
Laboratory Procedures--
Analytical Procedures—Approved test procedures for the analysis of
total Kjeldahl nitrogen in saline water include U.S. EPA Methods 351.1
and 351.3 and APHA Methods 417B, 417D, 417E, 420A, or 420B. The methods
should be consulted to determine which is most appropriate for available
equipment, expected concentrations, and expected level's of precision and
accuracy.
Calibration and Preventive Maintenance—Calibration procedures should
follow those specified in the method. If samples are being digested and
distilled, standards should also be digested and distilled prior to analysis
to check for ammonia contamination or loss during distillation. Concentrations
in the calibration standards should bracket the sample concentrations.
If a sample concentration is outside of the range of calibration, then
an additional calibration standard should be analyzed to check if the result
is within the linear range of the method. Alternatively, the sample should
be diluted to within the calibration range and then reanalyzed.
157
-------
Receiving Water
Nitrogen (Total Kjeldahl)
Quality Control Checks--Duplicate analyses should be conducted on
a .-ninimum of 5 percent of the total number of samples, with an additional
5 percent of the samples spiked and analyzed for percent recovery. A blank
snould be analyzed with each batch of samples. A U.S. EPA performance
sample should be analyzed at least once per quarter.
Corrective Action—Because ammonia is a component of Kjeldahl nitrogen,
precautions against contamination that were described for anmonia analyses
snould be followed.
Data Quality and Reporting—The detection and accurate quantification
of Kjeldahl nitrogen in receiving water is routinely attainable, although
method detection limits can vary widely because of methods or instrumentation.
The analytical method should be consulted to determine expected detection
limits, precision, and accuracy. Data should be reported in mg/L as N to
a maximum of three significant figures. Results should be reported for
all determinations, including QA replicates, blanks, and spiked samples.
Any factors that may have influenced sample quality should also be recorded.
158
-------
Receiving Water
Nitrogen (Nitrate and Nitrite)
Nitrogen (Nitrate and Nitrite)
Procedures--
Conection—Nitrate-nitrite samples can be collected in glass or plastic
containers. Prior to sample collection, each container and cap should
be rinsed thoroughly with sample water.
Pr oc ess ing--Ni irate- nitrite samples should be analyzed immediately
after collection. If a delay occurs, sa iples can be stored for up to 24 h
by acidification to pH<2 with sulfuric acid and held at 40 C. Samples
must not be preserved using mercuric chloride because the mercuric ion
accelerates the degradation of the cadmium-reduction column (APHA 1985).
Laboratory Procedures--
Analytical Procedures—Approved test procedures for the analysis of
nitrate-nitrite in saline waters include U.S. EPA Methods 353.2 and 353.3
and APHA Methods 418C and 418F. The methods should be consulted to determine
which is most appropriate for available equipment, expected concentrations,
and desired levels of precision and accuracy.
Calibration and Preventive Maintenance—Calibration procedures should
follow those specified in the method. Efficiency of each reduction column
should be checked by comparing a nitrite standard to a reduced nitrate
standard at the same concentration. This efficiency check should be made
at the beginning and the end of each sample run and at a minimum frequency
of every 10 samples. Reactivate the copper-cadmium granules when reduction
falls below 75 percent.
Concentrations of the calibration standards should bracket the sample
concentrations. If a sample concentration is outside the range of calibration,
then an additional calibration standard should be analyzed to check if
159
-------
Receiving Water
Nitrogen (Nitrate and Nitrite)
the result is within the linear range of tne method. Alternatively, the
sample should be diluted to within the calibration range and reanalyzed.
Quality Control Checks--Dup1icate analyses should be conducted on
a minimum of 5 percent of the total number of samples, with an additional
5 percent of the samples spiked and analyzed for percent recovery. A blank
fiat has been run through the reduction column should be analyzed with
each batch of samples or every 20 samples. A U.S. EPA performance sample
should be analyzed ? least once per quarter.
Corrective Action--Various components of the effluent can interfere
with the analysis. The method should be reviewed for ways to remove possible
interferences prior to analysis. Possible intereferences include suspended
solids, residual chlorine, oil and grease, and high concentrations of iron,
copper, or other metals.
The area where nitrate-nitrite analyses are performed should be well
isolated from exposure to nitric acid or nitric acid fumes.
Data Quality and Reporting—The detection and accurate quantification
of nitrate-nitrite in receiving water are routinely attainable, although
method detection limits can vary because of methods or instrumentation.
The analytical method should be consulted to determine expected detection
limits, precision, and accuracy. Data should be reported in mg/L as N
to a maximum of three significant figures. Results of all determinations
should be reported, Including QA replicates, blanks, and spiked samples.
Any factors that may have influenced sample quality should also be reported.
160
-------
Receiving Water
Phosphorus (Total)
Pnosphorus (Total)
Field Procedures--
Co llect ion—Phosphorus samples may be collected in glass or plastic
containers. Containers should be rinsed with IN HC1 followed by several
rinses with distilled water. Detergents containing phosphate should never
be used on containers or labware that is to be used for phosphate analysis.
Sample containers and lids should be rinsed thoroughly rith sample water
before sample collection.
Processing--Phosphorus samples can be stored up to 28 days before
analysis by acidification to pH<2 with sulfuric acid and refrigeration
at 40 c. Samples with low concentrations of phosphorus should not be stored
In plastic containers, as phosphates may adsorb onto the'container walls.
Laboratory Procedures--
Analytlcal Procedures—Approved test procedures for the analysis of
total phosphorus in receiving water are U.S. EPA Methods 365.1, 365.2,
and 365.3 and APHA Methods 424C, 424F, and 424G. The methods should be
consulted to determine which is most appropriate for available equipment,
expected concentrations, and desired levels of precision and accuracy.
Calibration and Preventive Maintenance—Calibration procedures should
follow those specified in the method. Concentrations of the calibration
standards should bracket the sample concentration. If a sample concentration
is outside the range of calibration, then an additional calibration standard
should be analyzed to check if the result is within the linear range of
the method. Alternatively, the sample should be diluted to within the
calibration range and reanalyzed.
Quality Control Checks—Duplicate analyses should be conducted on
a minimum of 5 percent of the total number of samples, with an additional
161
-------
Receiving Water
Phosphorus (Total)
5 percent of the sanoies spiked and analyzed for percent recovery. A blank
should be analyzed with each batch of samples. A U.S. EPA performance
sample should be analyzed at least once per quarter.
Corrective Action — Because phosphorus contamination can occur from
a variety of sources, it is recommended that a clearly marked set of lab-
ware be dedicated to only phosphorus analysis. This labware should never
be exposed to pnosphorus detergents or reagents containing phosphate.
Various components of the effluent can interfere witt the analysis.
The method should be reviewed for vays to remove interferences or adjust
for interferences from components that cannot be removed. Silica and arsenic
are possible positive Interferences, while hexavalent chromium and nitrite
can cause low recovery.
For highly colored or turbid samples, additional sample preparation
(e.g., further oxidation or filtration) may be required prior to color
development. In any case, blanks should be prepared by adding all the
reagents except the coloring reagents to the sample. Measure absorbance
in the sample blank at the wavelength used for the phosphorus determination
and subtract this absorbance value from the sample absorbance prior to
calculation of phosphorus concentration.
Data Quality and Reporting—Detection and accurate quantification
of total phosphorus in receiving water is routinely attainable. Actual
method detection limits can vary because of methods or instrumentation.
The analytical method should be consulted to determine expected detection
limits, precision, and accuracy. Data should be reported in mg/L as P
to a maximum of three significant figures. Results of all determinations
should be reported, including QA replicates and spiked samples. Any factors
that may have influenced sample quality should also be reported.
162
-------
Receiving Water
Total and Fecal Coliform Bacteria
Total and Fecal Coliforro Bacteria
Field Procedures--
Con ection--Samples should be collected in clean, sterile polypropylene
or glass containers. The sample containers must be resistant to sterilizing
conditions and to the solvent action of water. The container lids must
not produce bacteriostatic or nutritive compounds upon sterilization.
The sample containers must seal tightly. Containers with chips, cracks,
or etched marks should be discarded.
Heat-resistant glass or plastic sample containers should be autoclaved
at 1210 c for 15 min. Alternatively, dry glass containers can be sterilized
in a hot-air oven at 1700 c for at least 2 h. For plastic containers that
are not heat-resistant, ethylene oxide gas sterilization is acceptable
(Bordner et al. 1978). Containers sterilized by gas should be stored at
least 12 h before use to ensure all gas has dissipated.
If the sample water has residual chlorine, sodium thiosulfate should
be added (to a concentration of 0.008 percent) to neutralize the chlorine
and thereby prevent continued bactericidal action after sample collection.
In this manner, the true microbial content of the water at the time of
sampling can be estimated more accurately. If sodium thiosulfate must
be added to a sample, it should be added to the sample container prior
to sterilization.
If the sample water contains heavy metals in concentrations exceeding
0.01 mg/L, a chelating agent should be added to the sample container to
reduce metal toxicity. This is particularly important if samples are not
analyzed within 4 h after collection. APHA (1985) recommends using the
disodium salt of ethjlenediaminetetracetic acid (EOTA), adjusted to pH
6.5, and added to the sample container before sterilization. For a 120-mL
container, addition of 0.3 ml of a 15-percent EDTA solution is considered
adequate (APHA 1985).
163
-------
Receiving Water
Total and Fecal Coliform Bacteria
It is critical that samples are not contaminated during the collection
process. To avoid contamination, sterilized containers should be kept
sealed until they are used, containers should be filled without rinsing,
and container lids should be replaced immediately after the samples have
been collected. When removed from containers, lids should be held face
dcwn in one hand and not set down on any surface. Adequate headspace (at
least 2.5 cm) should be left in each sample container to facil itate mixing
prior to analysis.
Processing—Samples should be analysed as soon as possible after col-
lection. If a delay occurs, samples should be held at 40 C for a maximum
of 6 h. The length of delay should be noted on the log sheet.
Laboratory Procedures--
Analytical Methods—Details of the membrane filter (MF) method and
the most probable number (MPN) method are presented in Part III of Bordner
et al. (1978) and in Parts 908 and 909 of APHA (1985). Although the MF
method is more precise than the MPN method, it is also more sensitive to
interference from turbidity in samples. Because the MF technique usually
yields low and variable recovery from chlorinated waters, the MPN technique
should be used when samples contain chlorine residual.
Calibration and Preventive Maintenance—This information is reviewed
extensively in Part IV of Bordner et al. (1978) and in Part 902 of APHA
(1985).
Quality Control Checks--Quality control checks for total and fecal
coliform bacteria analyses are listed in detail in Part IV of Bordner et al.
(1978) and in Part 902 of APHA (1985). The list includes:
• Sterility checks on media, dilution and rinse water, glassware,
and membrane filters
164
-------
Receiving Water
Total and Fecal Coliform Bacteria
• Duplicate analyses on 10 percent of samples and on at least
one sample per test run
• Colony verifications on a monthly basis.
Corrective Action—Procedures detailed in the relevant sections of
Bordne: et al. (1978) and APHA (1985) should be followed.
Data Qua! ity and eport ing—Table 909:11 of APHA (1985) presents 95-percent
confidence limits for - ne MF method for coliform colonies of 1, 2, 3, 4,
5 and 10. The precision of the MPN method increases with increasing number
of replicates. With five tubes, each with 1 ml of sample, a completely
negative, result is expected less than 1 percent of the time (APHA 1985).
Confidence limits (95 percent) for various MPN counts are given in Tables
908:111, 908:IV, and 908:V of APHA (1985).
Using the MF method, data should be reported as densities of coliforms
per 100 mL. Using the MPN method, data should be reported as MPN values
per 100 ml. Results of all determinations should be reported, including
QA replicates, sterility checks, and colony verifications. Any factors
that may have influenced sample quality should also be reported.
165
-------
Receiving Water
Enterococcus Bacteria
Enterococcus Bacteria
Field Procedures--
Col lection—Samples snould be collected in clean, sterile polypropylene
or glass containers. The sample containers must be resistant to sterilizing
conditions and to the solvent action of water. The container lids must
not produce bact :riostatic or nutritive compounds upon sterilization.
The sample containers must seal tightly. Containers with chips, cracks,
or etched marks should be discarded.
Heat-resistant glass or plastic sample containers should be autoclaved
at 121Q C for 15 min. Alternatively, dry glass containers can be sterilized
in a hot-air oven at 1700 C for at least 2 h. For plastic containers that
are not heat-resistant, ethylene oxide gas sterilization is acceptable
(Bordner et al. 1978). Containers sterilized by gas should be stored at
least 12 h before use to ensure all gas has dissipated.
If the sample water has residual chlorine, sodium thiosulfate should
be added to neutralize the chlorine and thereby prevent continued bactericidal
action after sample collection. In this manner, the true microbial content
of the water at the time of sampling can be estimated more accurately.
If sodium thiosulfate must be added to a sample, it should be added to
the sample container prior to sterilization so that the final concentration
in the sample will be 100 mg/L. For a 120-mL container, 0.1 nt of a 10-percent
solution of sodiun thiosulfate will neutralize a sample containing as much
as 15 mg/L of residual chlorine (APHA 1985).
If the sample water contains heavy metals in concentrations exceeding
0.01 mg/L, a chelating agent should be added to the sample container to
reduce metal toxicity. This is particularly important if samples are not
analyzed within 4 h after collection. APHA (1985) recommends using the
disodium salt of ethylenediaminetetracetic acid (EOTA), adjusted to pH
166
-------
Receiving Water
Enterococcus Bacteria
6.5, and added to the sample container before sterilization. For a 120-mL
ccnta-'ner, addition of 0.3 ml of a 15-percent EDTA solution is considered
acequcte (APHA 1985).
It is critical that samples are not contaminated during the collection
process. To avoid contamination, sterilized containers should be kept
sealed until they are used, containers should be filled without rinsing,
and container lids should be replaced immediately after the samples have
been collected. When removed from containers, lids should be held face
down in one hand and not set down on any s rface. Adequate headspace (at
least 2.5 cm) should be left in each sample container to facilitate mixing
prior to analysis.
Process ing--Samples should be analyzed as soon as possible after col-
lection. If a delay occurs, samples should be held at 40 C for a maximum
of 6 h. The length of delay should be noted on the log sheet.
Laboratory Procedures--
Analytical Methods—Methods for analyzing enterococcus bacteria are
currently being finalized by U.S. EPA.
Calibration and Preventive Maintenance—This information is reviewed
extensively in Part IV of Bordner et al. (1978) and in Part 902 of APHA
(1985).
Quality Control Checks—Quality control checks for these analyses
are listed in detail in Part IV of Bordner et al. (1978) and in Part 902
of APHA (1985). The list includes:
• Sterility checks on media, ailution and rinse water, glassware,
and membrane filters
167
-------
Receiving Water
Enterococcus Bacteria
• Duplicate analyses on 10 percent of samples and on at least
one sample per test run
• Colony verifications on a monthly basis.
Corrective Action--Procedures detailed in the relevant sections of
Bordner et al. (1973) and APHA (1985) should be followed.
Data Quality and Reporting — Data should be reported according to the
specifications v. -rently being finalized by U.S. EPA.
163
-------
Receiving Water
Chlorophyll ^
Chlorophyll a
Field Procedures—
Col lection—Chlorophyll ^ samples can be collected in glass or plastic
containers. Because acids will decompose chlorophyll a_ to phaeophyton,
it is critical that the water sampler and sample containers remain free
o* acids (including acids frcm fingerprints). In addition, sample containers
should be rinsed three times before sample collection.
Processing—Chlorophyll ^ samples should oe filtered immfcdiately after
collection. Two or three drops of magnesium chloride suspension should
be added to the sample before filtration to prevent the sample from becoming
acidic. Filters can be stored for a few weeks by holding them in the dark
in a desiccator at -200 c (Strickland and Parsons 1972). However, storage
usually leads to low results and makes the extraction of chlorophyll more
difficult. It is therefore recommended that filters be extracted imnediately
after filtration. If filters are stored, the length of delay until analysis
should be recorded on the log sheet.
Laboratory Procedures--
Analytical Procedures—Chlorphyll £ determinations using the fluorometric
method are described in detail in Section IV.3.IV of Strickland and Parsons
(1972) and as Method 1002G2 of APHA (1985). All work with chlorophyll
extracts should be conducted in subdued light to avoid degradation. Opaque
containers or containers wrapped in aluminum should be used to protect
samples from light. Glass fiber filters are recommended because they are
inexpensive and result in practically no blank (Strickland and Parsons
1972). To improve extraction efficiency, a cell-grinding steo should be
included prior to extraction. A TFE/glass grinder should be used for glass
fiber filters.
169
-------
Receiving Water
Chlorophyll £
Calibration and Preventive Maintenance—The fluorometer should be
calibrated spectrophotometrically using samples from the same standard
chlorophyll solution. This solution should have a known concentration
of chlorophyll a^ that was extracted from marine phytoplankton. Strickland
and Parsons (1972) recommend that a mixed culture of equal amounts (by
pigment) of Skeletonema costatum, Coccolithus huxleyi i, and Peridinium
trochoidium be used as the sources of chlorophyll. If natural phytoplankton
populations are used, phaeo-pigments may be present.
A series of dilutions of the standard chlorophyll solution shu Id
be made so that concentrations of 2, 6, 20, and 60 ug/'L are achieved.
Readings of each dilution should be made at sensitivity settings of Ix,
3x, lOx, and 30x.. This will allow derivation of calibration factors to
convert fluorometric readings in each sensitivity level to chlorophyll ^
concentrations (for details, see APHA 1985).
Quality Control Checks—To correct for scatter, the fluorometer should
be zeroed against a cuvette of 90-percent acetone for each level of sensi-
tivity. Duplicate analyses should be conducted on at least 10 percent
of the total number of samples.
Corrective Action—If the fluorometer does not appear to be functioning
properly, the manufacturer1 s troubleshooting guide should be consulted.
Scratched or etched cuvettes should be replaced.
Data Quality and Reporting—It is recommended that chlorophyll £ concen-
trations be determined using the fluorometric method rather than the spectro-
photometr ic method, because the former technique is more sensitive, requires
less sample, and can be used for in vivo measurements (APHA 1985). Precision
of the recommended method varies as a function of the amount of pigment
being measured. For chlorophyll £ concentrations exceeding 0.5 mg/m3,
a precision of +8 percent is possible. The sensitivity of detection has
been estimated as 0.01 mg/m3 for a 2-L sample (Strickland and Parsons 1972).
Chlorophyll ^ concentrations should be reported as mg/m3 to the nearest
170
-------
Receiving Water
Chlorophyll a
0.01 unit. Results of all determinations should be reported, including
QA replicates. Any factors that may have influenced sample quality should
also be reported.
171
-------
Receiving Water
Phytoplankton
Phytoplankton
Field Procedures--
Pre-Collection Preparation--A solution of 100-percent buffered formalin
should be prepared by adding 2 g of sodium borate per 98 ml of undiluted
formalin. Sample jars should have a capacity of approximately 1 L and
should have lined plastic or polyethylene lids. Each jar should be marked
at the point where it is 90 percent full.
Coliection--When the sampler is retrieved, it should be shaken to
disperse the organisms. (Note: swirling is not effective for dispersing
plankton.) Each sample should then be drained into a marked 32-oz jar until
the water level reaches the 90-percent full mark.
Process ing--Because preservation renders many phytoplankton forms
taxonomically Intractable and indistinguishable from detritus, it is highly
desirable to analyze live material (Stofan and Grant 1978). Unfortunately,
this is not practical for most routine monitoring programs. Hence, procedural
recommendations given herein address preserved samples.
It is recommended that formalin be used to fix the phytoplankton samples.
However, other fixatives are available if formalin proves unsatisfactory
(see Stofan and Grant 1978). Once a fixative is selected, it should be
used exclusively so as not to bias the data.
After each sample has been collected, the jar should be filled to
the top with 100-percent buffered formalin and the cap should be screwed
on tightly. The sample jar should then be inverted several times to mix
the contents.
Samples should be stored upright in an opaque box to minimize exposure
to sunlight and in a cool area to minimize exposure to high temperatures.
Samples should also be stored in a stable part of the ship to minimize
172
-------
Receiving Water
Phytoplankton
agitation. If special racks are not available to keep the shipping boxes
from moving, they should be lashed down.
Laboratory Procedures--
Analytical Procedures—As with most biological field samples, handling
is detrimental to specimen quality, and hence, data quality. Handling
should therefore be kept to a minimum.
Several methods of : >ecimen identification and enumeration may be
used (Stofan and Grant 19/8). The most common is the Utermohl method,
by which phytoplankters are identified using an inverted microscope and
a fixed, settled sample. Contents of the sample jar are first resuspended
by gently shaking in an up-and-down motion. An aliquot Is then poured
into a graduated vertical counting chamber and allowed to settle. Phytoplank-
ters are then identified as they rest on the bottom of the chamber. Scanning
electron microscopy and Nomarski illumination for light microscopy can
be used to aid taxonomic identifications.
If possible, phytoplankters should be identified and enumerated from
an unconcentrated sample. This 1s highly desirable because sample handling
is minimized. However, in nutrient-poor waters the phytoplankton community
may be sparsely distributed, necessitating sample concentration. Several
concentrating procedures are commonly used (see Stofan and Grant 1978).
If sample concentration is necessary, tests should be conducted to see
which method Is the least harmful to the specimens. The selected method
should then be evaluated for efficiency by comparing it with replicate
aliquots of unconcentrated samples. After being accepted, the method should
be the only one used for concentrating samples. Data generated from concen-
trated samples should always be evaluated in light of the results of the
efficiency tests recommended above.
All phytoplankers should be identified to the species level and enumerated,
and the data should be recorded on a log sheet. After the identifications
173
-------
Receiving Water
Phytoplankton
h = ve been completed and the data have been recorded, each sample aliquot
trat was examined should be archived. Each aliquot should be stored in
a separate vial that is labeled internally with the following information:
survey area; station number; replicate number (if any); latitude and longitude,
Loran C, or other positioning information; date of collection; and collection
depth. Each vial should be sealed tightly with a polyseal cap, and all
vials from the same sample should be stored in a common container filled
^th preservative. To prevent evaporation of the fixative, vial and container
lids can be sealed with plastic tape. Although preserving and archiving
the sample aliquots is recommendeo for reference and verification purposes,
it should be remembered that many of the organisms will deteriorate over
time because of the caustic properties of the fixative.
Calibration and Preventive Maintenance—The inverted microscope should
be serviced annually unless the manufacturer recommends otherwise.
Taxonomic identifications should be consistent within the given laboratory,
and with the identifications of other workers. To that end, at least three
individuals of each taxon should be sent for verification to recognized
experts in museums and/or academic institutions. The verified specimens
snould then be photographed, and the photographs should be placed in the
permanent reference museum. Continued collection of a verified species
does not require additional expert verification. Participation of the
laboratory staff in a regional taxonomic standardization program (if available)
is required to ensure regional consistency and accuracy of identifications.
The photographic reference collection should be started and completed as
early as possible during the project.
Photographs of phytoplankton taxa are recommended because of the diffi-
culties involved in preserving phytoplankton specimens. The photographs
should be recorded on 35-mm color slide film using a high-quality photomicro-
scopy apparatus. Each slide should be coded, and an accessory logbook
of information should be maintained. The logbook should contain the following
Information: survey name; station number; replicate number (if any); latitude
174
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Receiving Water
Phytoplankton
and longitude, Loran C, or other positioning information; date of collection;
kind of sampler; and name of collector. In addition, the name of the taxonomic
expert who verified the identification, institutional affiliation, and
tne date of verification should be recorded.
A computer listing of each species' name, the person who verified
the identification, date of verification, location of the photograph in
tne collection, status of the photograph if it has been loaned to outside
exoerts, and references to pertinent literature should be maintained by
tne laboratory performing identification? for the monitoring program.
This listing should be available near the col ection. Reference photographs
are invaluable, and should be retained at the location where the identifications
were performed, in the offices of the funding agencies, or at a museum
with long-term storage capabilities. In no instance should this portion
of the collection be destroyed. A single person should be identified as
the curator of the collection and be responsible for its integrity.
Quality Control Checks--"New" taxonomists should have all of their
identified organisms verified within the laboratory, until their accuracy
reaches 95 percent. Thereafter, all identifications in one of every 10
samples should be checked for accuracy.
The most recent taxonomic literature should be used whenever possible.
Citations for all taxonomic references should be appended to the data report
for each survey.
Corrective Action— If a sample is found in which one or more species
are misidentified, all previous unchecked samples should be examined for
those specific errors. Errors should be rectified when located.
Data Quality and Reporting Requirements--Al1 phytoplankton should
be identified to the lowest possible taxon, preferably to the species level.
In cases where the identity of a species is uncertain, a species number
may be used (e.g., Chaetoceros sp. 1). Data for each sample should be
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Receiving Water
Phytoplankton
resorted as number of individuals per liter for each species. Results
of all determinations should be reported, including taxonomic verifications.
Any factors that may have influenced sample quality should also be reported.
176
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SEDIMENT/INFAUNAL ANALYSES
QA/QC procedures are presented in this section for the following 11
sediment/infauna variables:
• Grain size
t Total solids/water content
• Total volatile sol ids
• Total organic carbon
• Biochemical oxygen demand
• Chemical oxygen demand
• Oil and grease
t Sulfides
0 Priority pollutant metals
• Priority pollutant organic compounds
• Infauna.
Samples to be analyzed for these variables generally will be collected
using a bottom grab. Operation of a bottom grab is discussed earlier in
the general methods section.
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Sediment
Grain Size
Grain Size
Field Procedures--
Col lection--Grain size samples can be collected in glass or plastic
containers.
Processing — Grain size samples should be stored in a refrigerator
&~ 40 c to minimize the effects of bacterial growth. They may be held
for up to 6 mo in this manner. Samples must not be frozen or dried p -ior
to analysis, as either of these processes may change the particle-size
distribution.
Laboratory Procedures--
Analytical Procedures—Grain size should be measured according to
the sieving and pipet procedures described in Plumb (1981). A wet sample
is first sorted into coarse and fine fractions by wet sieving through a
63-urn (i.e., 4-phi) sieve. Particles having a diameter greater than 63
urn (i.e., -sand and gravel) are sorted by dry sieving through a graded series
of screens. Particles having a diameter less than 63 urn (i.e., silt and
clay) are sorted by their different settling velocities during the pipet
analysis.
An aliquot of the wet-sediment sample should be analyzed for total
solids to estimate the dry weight of the aliquot used for grain size analysis.
This estimated weight can be compared with the summed weight of all fractional
determinations to evaluate the efficiency of the method.
It is critical that each sample be thoroughly homogenized in the laboratory
before aliquots are removed. Laboratory homogenization should be conducted
even if samples were homogenized in the field.
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Sediment
Grain Size
If organic matter is oxidized prior to size analysis, the true particle
size will be determined. If organic matter is not oxidized prior to analysis,
tne apparent particle size will be determined. Because results from these
two techniques can differ, one method should be selected and used for all
grain-size determinations.
At a minimum, it is recommended that each sample be analyzed for percent
gravel, sand, silt, and clay. However, if the fine-grained fraction comprises
less th*n 1 percent of the total sample weight, the pipet analysis is not
recommer ed and the weight of the combined silt-clay fractions should be
reported. In very sandy sediments, the sand fraction may be subdivided
further. If this is desired, it is recommended that the fraction be sieved
at 1-phi increments to yield five subfractions.
Calibration and Preventive Maintenance—The analytical balance, drying
oven, sieve shaker, and temperature bath should be inspected each time
they are used, and calibrated weekly at a minimum. The manufacturer's
instructions should be consulted for calibration of the analytical balance
and for preventive maintenance procedures for the analytical balance, drying
oven, sieve shaker, and temperature bath. It is recommended that a calibrated
dial thermometer be used in the drying oven to determine average temperature.
Quality Control Checks—Several procedures are critical to the collection
of high quality particle size data. Most important to the dry sieve analysis
is that the screens are clean before conducting the analysis, and that
all of the sample is retrieved from them. To clean a screen, it should
be inverted and tapped on a table, while making sure that the rim hits
the table evenly. Further cleaning of brass screens may be performed by
gentle scrubbing with a stiff bristle nylon brush. Stainless steel screens
may be cleaned with a nylon or brass brush.
The most critical aspect of the pipet analysis is knowledge of the
temperature of the silt-clay suspension. An increase of only 1° C will
increase the settling velocity of a particle 50 urn in diameter by 2.3 percent.
179
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Sed imen t
Grain Size
It is generally recommended that the pipet analysis be conducted at a constant
temperature of 200 c. However, Plumb (1981) provides a table to correct
for settling velocities at other temperatures. Thorough mixing of the
silt-clay suspension at the beginning of the analysis is also critical.
A perforated, plexiglass disc plunger is very effective for this purpose.
Replicate sieve and pipet analyses should be conducted on at least
10 percent of the total number of samples analyzed. If >20 samples are
sorted, it is recommended that at least one tripiic te en'lysis be performed.
Corrective Action--An analysis should be repeated if weighing errors
exceed 5 percent of the original sample weight. If the mass of sediment
used for pipet analysis exceeds 25 g (0.9 oz), a subsample should be used
as described by Plumb (1981). Silt-clay samples in excess of 25 g (0.9 oz)
may give erroneous results because of electrostatic interactions between
the particles. Silt-clay samples less than 5 g yield a large experimental
error in weighing relative to the total sample weight. Other reasons for
erroneous results include hindered settling and flocculation.
Data Quality and Reporting — Ideally, the summed weight of all sample
fractions should differ from the original sample weight by less than 1 percent
(Plumb 1981). Weights of the six gravel/sand fractions, the total silt
fraction, and the total clay fraction should be reported to the nearest
0.0001 g. See Shepard (1963) for size ranges of the textural classes.
Results of all determinations should be reported, including QA replicates.
Any factors that may have influenced sample quality should also be reported.
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Sediment
Total Solids/Water Content
Total Solids/Water Content
Field Procedures—
Collection--Samples can be collected in glass or plastic containers.
Processing — Samples should be stored frozen and can be held for up
to 6 mo *:n that condition.
Laboratory Procedures—
Analytical Procedures—Total solids should be determined using the
procedure described in Plumb (1981). The general procedure is to weigh
a wet sample, dry it at 103-1050 C, reweigh the sample, and determine the
percentage of wet weight accounted for by the dry weight. Unrepresentative
material should be removed from the sample prior to analysis and noted
on the laboratory log sheet.
Because total solids content is operationally defined by the drying
temperature, it is essential that this temperature be held constant at
the specified value. It is also critical that each sample be thoroughly
homogenized in the laboratory before a subsample is taken for analysis.
Laboratory homogenization should be conducted even if samples were homogenized
w the field. Ignition of the evaporating dish prior to analysis is critical
to ensuring that the container is free from contaminants.
Calibration and Preventive Maintenance—The analytical balance, drying
over, and muffle furnace should be inspected and calibrated weekly, at
a minimum.
Quality Control Checks—Duplicate analysis should be conducted on
at least 10 percent at the total number of samples.
Corrective Action—None applicable.
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Sedimen t
Total Solids/Water Content
Data Quality and tteporting--Total solids content or, alternatively,
water content should be reported as a percentage of the wet weight of the
sediment sample to the nearest O.i unit. Results of all determinations
snould be reported, including QA replicates. Any factors that may have
influenced sample quality should also be reported.
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Sediment
Total Volatile Solids
Total Volatile Solids (TVS)
Field Procedures--
Collection—Samples can be collected in glass or plastic containers.
Processing—Samples can be stored frozen and can be held for up to
6 mo in that condition.
Laboratory Procedures--
Analytical Procedures—Methods for volatile solids analysis are given
by Plumb (1981). The basic procedure is to dry a sediment sample at 103-1050 C,
weigh it, and then combust it at 5500 C. The difference in mass between
the combusted sample and the desiccated sample estimates the mass of volatile
organic matter, unrepresentative material should be removed from the sample
prior to analysis and noted on the laboratory log sheet.
Because total volatile solids is operationally defined by the drying
and ignition temperature, it is essential that these temperatures be held
constant at their specified levels. It is also critical that each sample
be thoroughly homogenized in the laboratory before a subsample is taken
for analysis. Laboratory homogenization should be conducted even if samples
were homogenized in the field. Evaporating dishes (or crucibles) must
be ignited at 550<> C before being used for total volatile solids analysis.
This step ensures that the dishes are free from volatile contaminants.
Dried and combusted samples should be cooled in a desiccator and held there
until they are weighed. If a desiccator is not used, the sediment will
accumulate ambient moisture and the sample weight will be overestimated.
A color-indicating desiccant is recommended so that spent desiccant can
be detected easily. Also, the seal on the desiccator should be checked
periodically and, if necessary, the ground glass rims should be greased
or the "0" rings should be replaced.
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Sediment
Total Volatile Solids
Calibration and Preventive Maintenance--The analytical balance, drying
oven, and muffle furnace should be inspected and calibrated weekly, at
a minimum. The muffle furnace thermocouples and thermometers should be
inspected before each use and calibrated as needed.
Quality Control Checks—Duplicate analyses should be conducted on
at least 10 percent of the total number of samples.
Corrective Action—None appl -cable.
Data Quality and Reporting—Total volatile solids content should oe
reported as a percentage of the dry weight of the uncombusted sediment
sample to the nearest 0.1 percent. Results should be reported for all
determinations, Including QA replicates. Any factors that may have influenced
sample quality should also be reported.
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Sediment
Total Organic Carbon
Total Organic Carbon (TOG)
Field Procedures--
Collection—Samples can be collected in glass or plastic containers.
Processing—Samples should be stored frozen and can be held for up
to 6 mo in that condition. Excessive temperatures should not be used to
tnaw samples.
Laboratory Procedures--
Analytical Procedures—The method for total organic carbon analysis
is given by Plumb (1981). The recommended procedure is to first acidify
the samples to remove inorganic carbon, and then convert all organic carbon
compounds to carbon dioxide by catalytic combustion.
Because inorganic carbon (e.g., carbonates, bicarbonates, free C02)
will interfere with total organic carbon determinations, samples must be
treated to remove inorganic carbon before being analyzed. Samples may
be combusted in one of several different models of induction furnace.
All combustions should be performed in a single furnace (even if two furnaces
are the same model), as difrsrent furnaces may give slightly different
results. It is also critical that each sample be thoroughly homogenized
in the laboratory before a subsample is taken for analysis. Laboratory
homogenization should be conducted even 1f samples were homogenized in
the field. Dried samples should be cooled in a desiccator and held there
until they are weighed. If a desiccator is not used, the sediment will
accumulate ambient moisture and the sample weight will be overestimated.
A color-indicating desiccant is recommended so that spent desiccant can
be detected eaily. Also, the seal on the desiccator should be checked
periodically and, if necessary, the ground glass rims should be greased
or the "0" rings should be replaced.
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Sediment
Total Organic Carbon
Calibration and Preventive Maintenance—The induction furnace used
for the total organic carbon analyses should be inspected and calibrated
daily. The analytical balance and drying oven should be inspected and
calibrated weekly, at a minimum.
Quality Control Checks—Duplicate analyses should be conducted on
at least 10 percent of the total number of samples. An NBS traceable standard
reference material should be analyzed daily to monitor performance. A
method blank should be analyzed with each batch of samples.
Corrective Action—If the induction furnace does not appear to be
operating properly, follow the manufacturer's instructions for troubleshooting
and repair.
Data Quality and Reporting—Total organic carbon should be reported
as a percentage of the dry weight of the unacidified sediment sample to
the nearest 0.1 unit. Results should be reported for all determinations,
including QA replicates. Any factors that may have influenced sample quality
should also be reported.
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Sediment
Biochemical Oxygen Demand
S'.ochemical Oxygen Demand (BOD)
Field Procedures--
Con ection—Samples can be collected in glass or plastic containers.
Processing--BOD samples should be analyzed immediately after collection.
If a delay occurs, samples should be stored at 40 C, and can be held for
ua to 7 days in that condition. Samples should be kept field moist and
air contac' should be prevented to minimize oxidation. Refrigerated samples
should be v irmed to 200 c prior to analysis.
Laboratory Procedures--
Analytical Methods—BOD should be determined using the method described
by Plumb (1981). This procedure is similar to U.S. EPA Method 405.1 and
APHA Method 507 for water and waste water and specifies an incubation period
of 5 days at 20° C.
It is critical that each sample be thoroughly homogenized in the laboratory
before a subsample is taken for analysis. Laboratory homogenization should
be conducted even if samples were homogenized in the field. A subsample
should be analyzed separately for total solids so that BOO can be determined
on a dry-weight basis. The bacterial seed should be from a source having
a salinity similar to that of the environment from which the samples were
taken. The salinity of the dilution water should be adjusted with sea
salt (if necessary) so that its salinity approximates that of the environment
from which the sample was taken. Many synthetic organic components in
sediments are not biodegradable without the seeding procedure because of
either a toxic effect or a deficiency or absence of appropriate microorganisms.
Chlorine residuals must be removed prior to the test because residual chlorine
may be toxic to the microbial population or may oxidi*e organic material.
Because sediments that contain sulfide, sulfite, or ferrous ions create
an immediate demand on the dissolved oxygen, it is necessary to distinguish
187
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Sediment
Biochemical Oxygen Demand
this immediate demand from the true BOD. The depletion of dissolved oxygen
during 15 min in a standard water dilution of the sample was arbitrarily
selected by Plumb (1981) as the initial oxygen demand.
Calibration and Preventive Maintenance—Dissolved oxygen measurements
should be calibrated according to the procedures recommended in the appropriate
sections of this document.
Quality Control Checks--Dup1icate analyses should be conducted on
a minimum of 10 percent of the total number of : mp.es. A dilution water
blank and a g 1 ucose-glut ami c acid standard pro/ide quality control on the
dilution water as. well as on the cleanliness of analytical equipment (e.g.,
incubation bottles) and should each be analyzed in triplicate with each
batch of samples. The most reliable BOD determinations are made when residual
dissolved oxygen is at least 2 mg/L and uptake of dissolved oxygen is at
least 2 mg/L after incubation (Plumb 1981).
Corrective Action—Plumb (1981) and APHA (1985) should be consulted
for methods of correcting for the many kinds of interference that may accompany
BOD analyses.
If the dilution water blanks exceed 0.2 mg/L, cleanliness of containers
and water should be checked. Containers may require 1+1 HC1 rinse after
detergent washing to remove and any residual organic material. Any containers
rinsed with add should be thoroughly rinsed with distilled water to prevent
acid carryover.
If a 2-percent dilution of the glucose-glutamic acid standard check
solution is out sice the range of 218±11 mg/L, BOO determinations made with
the seed and dilution water are suspect. Several methods used to determine
the problem include running a series of dilution water blanks using different
water sources with and without seed, preparing a fresh solution of glucose-
glutamic acid, changing the seed, or preparing fresh reagents for the dilution
188
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Sediment
Biochemical Oxygen Demand
water. The source of the problem should be determined before additional
BOO analyses are performed.
Data Quality and Reporting--BOD should be reported as mg/kg dry weight
of sediment to the nearest 0.1 unit. The laboratory should report the
results of all determinations, including QA replicates, dilution water
blanks, and glucose-glutamic acid standards. Any factors that may have
influenced sample quality should also be reported.
189
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Sediment
Chemical Oxygen Demand
Chemical Oxygen Demand (COD)
Field Procedures--
Col1ection--Samp1es can be collected in glass or plastic containers.
Processing--COD samples should be analyzed immediately after collection.
If a del~.y occurs, samples should be stored at 4° C, and can be held for
up to 7 -jays in that condition. Samples must be kept field moist and free
from air contact during stor -,e to .rsinimize oxidation.
Laboratory Procedures—
Analytical Methods--CQD should be determined according to the method
described in Plumb (1981). This open reflux method is similar to U.S. EPA
Method 410.1 and APHA Method 508A for water and waste water and consists
of oxidizing the organic matter in a reflux apparatus for 2 h.
It is critical that each sample be thoroughly homogenized in the laboratory
before a subsample is taken for analysis. Laboratory homogenization should
be conducted even if samples were homogenized in the field. A subsample
should be analyzed separately for total solids so that COD can be determined
on a dry-weight basis.
Because traces of organic material from external sources may cause
a positive error, care should be taken to avoid contamination of glassware
and the distilled water used to prepare reagents and to reflux the sample.
Because volatile materials may be lost when the sample temperature rises
during the sulfuric acid addition step, the flask should be cooled during
this addition to minimize this loss. Because chlorides are quantitatively
oxidized by dichromate and represent a positive interference, mercuric
sulfate is added to the digestion flask to complex the chlorides.
190
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Sed imen t
Chemical Oxygen Demand
Calibration and Preventive Maintenance--The technique and quality
of reagents should be evaluated by conducting the test on a standard potassium
hydrogen phthalate solution. Laboratory glassware should be kept very
clean to prevent introduction of organic material to the sample.
Quality Control Checks--Duplicate analyses should be conducted on
a minimum of 10 percent of the total number of samples. A method blank
and a potassium hydrogen ohi-halate standard should be analyzed with each
batch of samples.
Corrective Action—If results of the analysis conducted on the potassium
hydrogen phthalate standard indicate that the technique is not performing
properly, check glassware and the distilled water for contamination, ensure
that reagents and sample are thoroughly mixed, ensure that excess potassium
dichromate remains after oxidation is complete, and critically review all
other analytical steps.
Data Quality and Reporting--COD should be reported as mg/kg dry weight
of sediment to the nearest 0.1 unit. The laboratory should report the
results of all determinations, including QA replicates, method blanks,
and potassium hydrogen phthalate standards. Any factors that may have
influenced sample quality should also be reported.
191
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Sediment
Oil and Grease
Oil and Grease
Field Procedures--
Col 1ection--Sediment samples should be collected In wide-mouth glass
jars. Bottles should first be washed with a warm aqueous detergent mixture,
and then, in sequence, thoroughly rinsed with hot tap water, rinsed at
least twice with distilled water, rinsed with l,l,2-trichloro-l,2,2-trifluoro-
ethane (-',e. preon or equivalent), and dried in a clean oven at >_ 105° C
for 30 mir, Bottle lids should be lined with TFE. Aluminum-lined lids
can be used, but contact with seawater corrodes the aluminum. Plastic
containers are not acceptable. Headspace should be left in the sample
container for addition of acid and mixing or expansion during freezing.
Processing—When analysis cannot be made within 24 h, preserve the
sample with approximately 1 ml of concentrated hydrochloric acid per 80 g
(wet weight) of sample. Never preserve with chloroform or sodium benzoate
(APHA 1985). Acid-preserved samples should be stored at 40 C, and can
be held for up to 28 days in that condition. Although U.S. EPA has not
established a recommended maximum holding time for oil and grease in sediments,
28 days is consistent with the recommended holding time for acid-preserved
water samples. Samples can also be preserved by freezing at -20° C, and
can be held under that condition for up to 6 mo. Samples must be kept
field moist during storage because they may lose apparent oil and grease
as a result of drying.
A wet sample should be used for analysis because a dried sample may
yield low results. Concentration on a dry-weight basis can be obtained
by analyzing a separate aliquot for total solids.
Laboratory Procedures—
Analytical Procedures—Methods for oil and grease analysis are described
in Plumb (1981). Soxhlet extraction of the sediment can be inefficient
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Sed imen t
Oil and Grease
due to channeling of solvent through the sediment layer. Therefore, all
sediments should be stirred at least twice during the extraction period.
Because asphaltic materials are insoluole in freon, the recommended method
will give low recoveries for samples containing these materials.
Determination of extracted oil and grease concentrations by the gravimetric
method is reliable for relatively nonvolatile hydrocarbons, vegetable oils,
animal fats, waxes, and soaps. Elemental sulfur is co-extracted with oil
and grease and will interfere wit > gravimetric determinations. Light hydro-
carbons that volatilize at tempe-atures below 700 C (e.g., gasoline through
No. 2 fuel oil) are lost during the solvent removal step.
Determination of extracted oil and grease concentrations by the infrared
method may be more precise at lower concentrations than is the gravimetric
method. This method will give higher results than the gravimetric method
if the sample contains volatile components, and will give lower results
than the gravimetric method if the sample is high in sulfur content. A
limitation of the infrared determination is standardization, which requires
a prepared reference oil. The reference oil may not be comparable to the
type of oil and grease in the samples, thereby resulting in inaccurate
results. When the exact nature of the oil and grease in the samples is
unknown, the reference oil described in U.S. EPA Method 413.2, Section 6.4
is recommended.
Calibration and Preventive Maintenance—For gravimetric oil and grease
analyses, check the accuracy of the analytical balance periodically (minimum
of once per week is recommended) using Class S weights. A service contract
that includes scheduled preventive maintenance at least once per year is
recommended. Scratched, chipped, or cracked boiling flasks should be replaced.
For infrared oil and grease analyses, follow the manufacturer's preventive
maintenance procedures for the infrared spectrophotometer. Cells used
for analysis should be checked for scratches before each use. Scratched
cells should not be used.
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Sediment
Oil and Grease
Quality Control Checks--Repl icate analyses should be performed for
a minimum of 10 percent cf the total number of samples to establish an
estimate of precision. Because spike results may not reflect true extraction
efficiencies, reference sample analysis, if available, is more appropriate.
A method blank should accompany each batch of samples.
Action--If concentrations of oil and grease in procedural
blanks are greater than the detection limit, check the cleanlin'ss of all
glassware. For infrared analyses, severe interferences will result if
the freon solvent contacts any material containing plasticizers (e.g, Tygon
tubing, plastic bottles). For gravimetric analyses, high results will
be obtained if Freon or fumes remain in the flask after distillation.
Poor precision may be caused by channeling of the solvent during ex-
traction. To minimize channeling, ensure that all moisture has been removed
with the addition of the magnesium sulfate and stir the sample more frequently
during the extraction step.
Data Quality and Reporting— The definition of oil and grease is based
on the procedures used. Unless identical procedures are used, oil and
grease determinations are not intercomparable. Thus, the method used for
analysis should always be specified and consistent within a study.
Reported values for precision and accuracy are indicated below:
• Reproducibil ity for gravimetric determinations of oil and
grease in sludge samples was 4.6-percent coefficient of
variation (APHA 1985). Variability in sediment oil and
grease determinations have often been reported as high as
20-30 percent (Disalvo et al . 1977).
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Sediment
Oil and Grease
t Due to the nonspecific nature of the oil and grease analysis,
interpretation of spike analyses is difficult. Analysis
of a reference material (U.S. EPA Municipal Digested Sludge)
has an acceptable recovery range of 50-150 percent of the
reference value.
Measurements should be reported to a minimum of two significant figures
and should clearly state whether concentrations are on a wet- or dry-weight
basis. Detection limits are in the range of 1 mg/kg (wet weight) for infrared
determinations nd 50 mg/kg (wet weight) for gravimetric determinations.
Results of all determinations should be reported, including QA replicates,
blanks, and reference samples. Any factors that may have influenced sample
quality should also be reported.
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Sed iment
Sulfldes (Total and Water Soluble)
Sulfides (Total and Water Soluble)
Field Procedures--
Col lection--Samples can be collected in glass or plastic containers.
Samples should be removed from the grab sampler as soon as possible to
minimize exposure to air.
Processing--Samples should be analyzed for sulfides as soon as possible.
Samples for total sal fides should be preser »d oy adding 2N zinc acetate
solution (approximately 5 ml for 30 g of seduent) and swirling the mixture.
Preserved samples for total sul fides should be stored in the dark at 40
C and should be analyzed within 7 days. Although U.S. EPA has not established
a recommended maximum holding time, 7 days would be consistent with the
holding time recommended for acid-preserved water samples. Samples for
water-soluble sul fides can be preserved by mixing 10-20 g of sediment in
a tared container containing 50 ml of sulfur antioxidant buffer (SAOB).
Preserved samples for water-soluble sulfides should be stored in the dark
at 40 c and should be analyzed within 4 days. All preserved sulfides samples
should be stored in the dark. It is critical that air contact with samples
be minimized and that samples be kept moist to minimize oxidation.
Laboratory Procedures--
Analytical Procedures--Total sulfides should be measured according
to procedures described in Plumb (1981). A distillation under acidified
conditions evolves hydrogen sulfide, which is swept into a zinc acetate
trap and precipitated as zinc sulfide. The sulfide is then solubilized
and measured using a colorimetric procedure.
Water-soluble sulfides should be measured according to the method
described in Green and Schnitker (1974). The buffered sediment must be
maintained in a slurry during titration. Although zinc or lead can be
used as a titrant, cadmium produces the sharpest endpoint. Because the
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Sed imen t
Sulfides (Total and Water Soluble)
SAOB solution will oxidize and darken with time, thereby losing its buffering
capacity, this solution should be as fresh as possible. Because samples
high in sul fides may exceed the buffering capacity of the SAOB solution,
sample size should be adjusted accordingly.
For both sulfides techniques, an aliquot of the original sediment
sample should be analyzed for total solids so that results can be expressed
on a dry-weight basis.
Calibration and Preventive Maintenance—A series f al least three
standard sulfide solutions should be prepare*1. Because si. 1 fide solutions
are very unstable, they should be prepared fresh for each use and used
iimiediately. Stability of these solutions can be increased by using nitrogen-
saturated water for dilution. Because of variation among lots, the methylene
blue solution should be standardized against a standard sulfide solution.
The spectrophotometer used for total sul fides should be calibrated daily
by zeroing the instrument using a method blank and establishing a calibration
curve using the standard sulfide solutions. The curve should span the
expected range of sulfides concentrations. The specific ion probe used
for water soluble sul fides should be calibrated according to the manufacturer's
instructions.
Quality Control Checks—Duplicate analyses should be conducted on
a minimum of 10 percent of the total number of samples.
Corrective Action—The amine-sulfurlc acid solution used in the color-
imetric determination should be pink initially and turn colorless within
3 min when used in the procedure with a sulfide-free sample. If color
appears, prepare a new stock solution with fresh materials.
Data Quality and Reporting—A detection limit of 5-10 mg/kg wet weight
can be expected for both sul fides techniques using a 10-20 g wet-weight
sample. Sul fides concentrations should be reported as mg/kg of sediment
dry weight to the nearest 0.1 unit. Results should be reported for all
197
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Sediment
SuIfides (Total and Water Soluble)
determinations, including QA replicates. Any factors that may have influenced
sample quality should also be reported.
198
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Sediment
Priority Pollutant Metals
Priority Pollutant Metals
Field Procedures--
Co11ection--As with water samples, the best containers for collection
of sediment for trace metal analysis are made of quartz or TFE. Because
these containers are expensive, the preferred containers are made of poly-
propylene or linear polyethylene with a polyethylene cap (APHA 1985).
Borr silicate glass containers can be used and may be preferred if trace
orga. ic compound analyses are to be performed on the same samples. Do
not use soft glass containers or containers with aluminum-lined or cardboard-
lined lids.
Possible problems during sample collection involve contamination from
the sampling device, airborne dust, or cross-contamination from previous
samples. Contamination can be minimized by avoiding the use of metal when
collecting sediment samples. If metal must be used, corrosive resistant
stainless steel is the best material. When using a benthic grab or coring
device, contamination can be minimized by removing only sediment that is
not touching the walls. Prior to use, sample containers should be thoroughly
cleaned with a detergent solution, rinsed with tap water, soaked in acid,
and then rinsed with metal-free water. All glass or plastic parts associated
with the sampling equipment should be cleaned in the same manner. For
quartz, TFE, or glass containers, use 1+1 HN03, 1+1 HC1, or aqua regia
(3 parts concentrated HC1 + 1 part concentrated HNQ/j) for soaking. For
plastic material, use 1+1 HNQ.3 or 1+1 HC1. Reliable soaking conditions
are 24 h at 700 C (APHA 1985). Do not use chromic acid for cleaning any
materials. For metal parts, clean as stated for glass or plastic, except
omit the acid-soak step of the cleaning procedure. Acids used should be
at least reagent grade. If trace organic compound analyses are to be performed
on the same samples, final rinsing with acetone and then high-purity methylene
chloride is acceptable.
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Priority Pollutant Metals
A minimum sample size of 5 g (wet weight) is required for the analysis
of all priority pollutant metals. To allow for duplicates, spikes, and
required reanalyses, a minimum sample size of 50 g (wet weight) is reccrrmended.
To allow for mixing of the sample and ease of collection, a 240-mL (8-oz)
jar is recommended for collection. A 125-mL (4-oz) jar would be adequate
but often difficult to fill.
Processing--S.-.mp1es should be stored in clean containers after collection,
--i.d picked in ice while in the field. Samples should be stored at -20°
C Although freezing is not required for all U.S. EPA procedures, it is
recommended to minimize potential alteration of analytes by microbes.
Care should be taken to prevent container breakage during freezing. Leave
sufficient headspace for water to expand and place the containers at an
angle when freezing.
No recommended holding time for sediments has been established by
U.S. EPA. A maximum holding time of 6 mo (except for mercury samples,
which should be held a maximum of 30 days) is consistent with the maximum
holding time recommended by U.S. EPA for water samples (U.S. EPA 1985).
Laboratory Procedures--
Analytical Procedures — Priority pollutant metals should be analyzed
according to procedures described in Tetra Tech (1986a). Prior to removing
each aliquot for analysis, samples should be mixed thoroughly using nonmetallic
utensils. Mix all water back into the sample. If there is any question
regarding nonrepresentative material (e.g., twigs, leaves, shells, rocks,
and any material larger than 1/4-in), U.S. EPA should be contacted for
guidance. A separate aliquot should be analyzed for a total solids deter-
mination.
Digest sediment samples prior to analysis using the acids specified
in the procedure (Tetra Tech 1986a). The digestate can then be analyzed
by flame Atomic Absorption Spectrophotometry (AAS), graphite furnace AAS,
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Sediment
Priority Pollutant Metals
or Inductively Coupled Plasma (ICP), depending on the sample concentrations
and required detection limit. Mercury digestion and analysis must be performed
on a separate sample aliquot by cold vapor AAS.
ICP can be used to screen samples for elements that are present in
relatively high concentrations. For those that may require more sensitive
analysis, graphite furnace AAS can be used. Analysis by ICP can be subject
to interelement interferences, while graphite furnace AAS can be subject
to matrix problems frcm acid or salt content of the samples. Select the
method with a detection limit that is adequate to determine compliance
with 301(h) program criteria.
Calibration and Preventive Maintenance—In general, all instruments
must be calibrated daily and each time the instrument is set up. For each
analysis, calibration procedures should follow those for the specified
method. Calibration standards must be prepared using the same concentrations
of acids as will result in the samples following sample preparation.
After an instrument has been calibrated, verify the accuracy of the
initial cal ibration by the analysis of certified control solutions at a
frequency of once every 10 samples or every 2 h during an analysis run,
whichever is more frequent, and after the last analytical sample. If a
certified control solution is not available, use a standard that is composed
of the analyte from a different source than that for the initial calibration.
If the deviation of the continuing calibration verification is greater
than the calibration control limits specified 1n the method, the instrument
must be recalibrated, and the preceding 10 samples reanalyzed.
All equipment should have scheduled routine preventive maintenance,
and a record of all maintenance performed should be noted in a logbook.
Critical spare parts should be kept on hand.
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Priority Pollutant Metals
Quality Control Checks—Analyze standard reference materials (SRM)
[e.g., the National Bureau of Standards (NBS) Estuarine Sediment or the
National Research Council of Canada (NRC) Marine Sediments] to provide
a check on digestion efficiency and overall accuracy of the analysis.
A minimum of one SRM should be analyzed for each survey or 2 percent of
the total number of samples (i.e., 1 per 50 samples), whichever is more
frequent.
To estimate r*ecision, 5 percent of the total number of samples should
be analyzed in duplicate or one duplicate for each survey, whichever is
more frequent. When more than 20 samples are to be analyzed for one survey,
the project manager may choose to implement a program of triplicate analyses.
The overall percentage of replicates should be at least 5 percent. To
estimate recovery, analyze samples spiked before digestion at the same
frequency as duplicates. Add spike concentration approximately equal to
the concentration found in the unspiked sample. An acceptable range in
spike concentrations is 0.5 to 5 times the sample concentrations.
Carry a method blank through all digestion and analysis steps at a
minimum frequency of once every 20 samples or once for each batch of samples
analyzed, whichever is more frequent. If the concentration of the blank
is less than the required detection limit, no correction of sample results
is performed. If the blank contamination is extensive (>30 percent of
sample value) then the batch of samples associated with the blank should
be reanalyzed. Data should be corrected by the data user for the blank
values between the required detection limit and the control limit.
For ICP analysis, additional QC checks should include an interference
check sample to verify interelement and background correction factors.
For graphite furnace AAS, additional QC checks should include duplicate
injections, with the mean value reported. The relative standard deviation
of the readings should be within control limits. Otherwise, the sample
should be reanalyzed.
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Priority Pollutant Metals
Corrective Action—If the concentration of the field or method blank
is greater than the required detection limit, all steps in the sample handling
snould be reviewed. Many trace metal contamination problems are due to
airborne dust. Keeping containers closed and rinsing all handling equipment
immediately prior to use minimizes dust problems. In the field, mercury-filled
thermometers should be handled carefully or avoided because broken thermometers
are a potential source of severe mercury contamination. In the laboratory,
sanples for mercury analysis should be isolated from items such as polarographs
or COD reagents.
Poor duplication may be caused by inadequate mixing of the sample
before taking aliquots, inconsistent contamination, gross grain size differ-
ences, inconsistent digestion procedures, or instrumentation problems.
Poor performance on the analysis of the Standard Reference Material
(SRM) or poor spike recovery may be caused for the same reasons as poor
duplication. However, if duplicate results are acceptable, poor SRM performance
or poor spike recovery may be caused by loss of analyte during analysis.
To check for analyte loss during digestion and for low recovery due to
interferences during analysis, spike the sample after digestion and compare
the analysis to the predigestion spike. If the results are different,
the digestion technique should be adjusted. If the results are the same,
dilute the sample by at least a factor of 5 and reanalyze. If spike recovery
is still poor, standard additions, matrix modifiers, or another method
is required.
Data Quality and Reporting—Report measurements as mg/kg to a maximum
of three significant figures on a dry-weight basis. Detection limits can
vary widely because of methods and instrumentation. Consult the analytical
method to determine expected detection limits, precision, and accuracy.
Detection limits actually obtained should be reported for each sample.
The laboratory data summary should include duplicate, spike, and blank
results and state clearly if and how any data were blank-corrected. Data
203
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Sediment
Priority Pollutant Metals
to be included in the ODES database should be blank-corrected by the data
user. The laboratory data suimary should also include the following information
to allow independent QA review:
• Digestion procedures
• Quantity of sample digested and final dilution volume
-i Percent soi ids
• Instrument detection limit for each element
0 Method of detection (I.e., graphite furnace, flame, ICP,
hydride, cold vapor)
• Deviation from the prescribed methods
• Blank associated with sample
• Problems associated with analysis.
For a more thorough QA review, additional documentation (e.g., calibration
curves) may be requested.
204
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Sediment
Priority Pollutant Organic Compounds
Priority Pollutant Organic Compounds
Field Procedures--
Col lection--Priority pollutant organic compounds can be separated
into purgeable (volatile) and extractable (acid, base, and neutral, including
pesticides and PCBs) compounds. Container preparation and collection techniques
differ for these two groups. Sediment samples for analysis of extractable
compounds should be collected in 240-mL (8-oz) or larger, wir^e-mouth glass
jars with TFE-lined screw lids. The container, lid, and lii.jr should be
detergent washed, rinsed twice with tap water, orce with distilled water,
once with acetone, and once with high-purity methylene chloride. Firing
of the glass jar at 450o C for 1 h may be substituted for the final solvent
rinse. Collection of a minimum of 200 g (wet weight) of sample should
provide enough material for a full analysis and all required QC analyses.
Headspace should be left to facilitate mixing of the sample.
If a volatile analysis is required, two separate 40-mL glass containers
should be filled leaving no headspace. This container, screw cap, and
septum should be washed with detergent, rinsed once with tap water, rinsed
again with distilled water, and dried at >105° C. Use of a solvent rinse
will interfere with the analysis. To obtain a sample with no headspace,
fill the vial to overflowing so that a convex meniscus forms at the top
if there is adequate water in the sediment. With the liner's TFE side
down, place the cap carefully on the opening of the vial, displacing the
excess water. Once sealed, invert the bottle to verify the seal by demon-
strating the absence of air bubbles. Samples for volatile analyses should
be taken only from single grab samples because many of the volatile compounds
of interest may be lost while compositing.
An empty sample jar carried through all processing and handling will
serve as a field blank. A solvent rinse of the bottle should be analyzed
205
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Sed imen t
Priority Pollutant Organic Compounds
by the lab with each batch of samples received to serve as a check on con-
tamination that may occur during shipping and storage. The solvent should
be the same as that used for sample extraction.
Process ing--Samples should be stored in the dark at 40 C, on ice,
or frozen until extraction. Care should be taken with frozen samples to
prevent container breakage by leaving headspace for the water to expand
and by freezing containers at an angle. U.S. SPA gives no official guidance
on sediment holding times but recommends that water saroles for extractable
organic compounds stored at 4° C be extracted within l«l days of collection.
Because sediments can be frozen at -20° C, longer holding times (e.g.,
up to 6 mo) are appropriate. Extracts should be analyzed within 40 days
of extraction. Effort should be made to analyze the samples as soon as
possible after extraction because some of the more labile analytes may
degrade in solution. Degradation may occur even in the dark under refriger-
ation, possibly as the result of free radical formation. Freezing is not
recommended for volatile samples because no headspace is to be left in
the vials. A maximum holding time of 14 days for analysis of volatile
compounds in sediment samples would be consistent with the holding time
recommended by U.S. EPA for water samples.
Laboratory Procedures--
Analytical Procedures--Priority pollutant organic compounds should
be determined according to procedures described in Tetra Tech (1986a).
Before each aliquot 1s removed for analysis, the sediment and any standing
water should be stirred well to create a homogeneous mixture. If there
is any question regarding possible nonrepresentative material (e.g., twigs,
leaves, shells, rocks, and any material larger than l/4-in), request guidance
from U.S. EPA. A separate aliquot should be analyzed for a total solids
determination.
To obtain meaningful results, an efficient extraction of the compounds
of interest must be performed. The length of time a sample should be extracted
206
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Sed imen t
Priority Pollutant Organic Compounds
varies with method and matrix. When dealing with an unfamiliar method
or matrix, step-by-step testing should be performed to deterrr.lne completeness
of extraction and to identify special problems. For instance, it has been
found when using the Soxhlet method with wet sediments, the sample must
be stirred two to three times during the procedure to prevent channeling.
Removal of elemental sulfur from marine sediments is critical for
improving column resolution in gas chromatography, and for low-level detection
of compounds such as PCBs by gas chromatography/electron capture. Cleanup
of the ample extracts using column chromatography is often necessary to
remove interferences. Calibration of the columns enables the compounds
of interest to be collected in the proper fraction.
Cross-contamination between samples should be avoided in all steps
of analysis beginning with clean glassware. Injection micro-syringes must
be cleaned well between uses. If separate syringes are used for injection
of standard solutions any bias between syringes should be accounted for.
Carryover can occur when high- and low-level samples are analyzed sequentially.
Analysis of an appropriate solvent blank following a high-level sample
may be necessary to check for carryover.
Generally, achievable detection limits for sediments are outlined
by specific methods. Actual detection limits vary depending on sample
size, final volume, co-extractive compounds, and the source and nature
of sediment. Select the method such that detection limits are adequate
to determine compliance to 301 (h) program objectives.
Calibration and Preventive Maintenance—Before beginning analysis
of samples, a calibration curve that brackets the working range must be
established. This calibration should be repeated after each major equipment
disruption. Calibration checks of the GC/MS system should be done at the
beginning and end of the day and at least every 12 h to demonstrate that
the instrument's response is within control limits. Specific tuning criteria
(e.g., OFTPP, BFB) are provided in each method. Calibration checks of
207
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Sediment
Priority Pollutant Organic Compounds
the GC/ECO system should be done at the beginning of the day and verified
at least every 6 h. These are only recommended minimum frequencies and
the nature of the samples may necessitate more frequent verification.
A routine QC check for each lot of the analytical reagents being used
in extraction can prevent undetected contamination problems. Also, each
lot of alumina, silica gel, sodium sulfate, Florisil, resins, or charcoal
used should be monitored as a possible source of contamination and cleaned
as necessary. Each lot of surroga e mixture should be checked for contam-
inants. The efficacy of the mater als listed needs to be evaluated as
there is variation between lots. Proper storage is essential for these
materials.
Equipment should be maintained and serviced routinely according to
manufacturer's instructions and good laboratory practices. Logbook records
should document maintenance for each measurement device. Critical spare
parts should be kept on hand.
Quality Control Checks—Duplicates and matrix spike analyses should
be performed to measure precision and accuracy. A frequency of 5 percent
of each or one each per survey, whichever is more frequent, is the reccmnended
minimum. When more than 20 samples are to be analyzed for one survey,
the project manager may choose to implement a program of triplicate analyses.
The overall percentage of replicates should be at least 5 percent. Method
interferences can be caused by contaminated glassware, reagents, solvents,
or processing hardware. These materials can be monitored for contamination
by processing of 5 percent method blanks, or one per sample set, whichever
is more frequent. Addition of known amounts of surrogate compounds to
each sample will serve to monitor preparation and analysis of samples.
Standard reference material should be analyzed if available, as another
measure of accuracy. Once for each survey or 2 percent of the total number
of samples, whichever is more frequent, is the recommended minimum.
208
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Sediment
Priority Pollutant Organic Compounds
Corrective Action--Uhen results of QC samples fall outside of established
limits, several courses of action are available. Contamina :ion in the
lab reagent blank sample is cause for similar findings of the same compounds
in samples to be suspect. If contamination is extensive, reanalysis of
the whole associated group may be in order. Blank contamination should
be kept to less than 10 percent of sample values and preferably below the
method detection limit. Contamination found in the field blank should
be considered when looking at the associated samp'ie data. Extensive contam-
ination of lab or field Hanks (>30 percent of sample values) should lead
to a detailed review of labi.ratory, sampling, transport, and storage pro-
cedures. Phthalates, methylene chloride, and toluene are common laboratory
contaminants that may be detected in blanks above the method detection
1imit.
Poor duplication may be caused by inadequate mixing of the sample
before removing aliquots, inconsistent contamination, inconsistent extraction
procedures, or instrument problems. The project manager should be contacted
when results exceed the control limits. Further replication of an analysis
may be necessary to determine the reason for the poor results.
Poor spike recovery may be caused for the same reasons as poor duplication
or by matrix effects produced by co-extracted materials. If the spiked
compound is added at a concentration much less than that found in the sample,
recovery may be difficult to determine. This problem is difficult to avoid
as most environmental samples contain unknown concentrations of organic
compounds. To check for analyte loss during processing, a step-by-step
examination of the method using a spiked blank is necessary, with measurements
of the analyte at each step.
Sample results that fall outside the established calibration curve
are suspect until linearity of the instrument response can be shown at
that concentration or until the extract is diluted appropriately and re-
analyzed. Extremely high concentrations of organic compounds may saturate
209
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Sediment
Priority Pollutant Organic Compounds
the extraction capabilities of the method and may necessitate re-extraction
of a smaller sample size or use of a more appropriate method.
Emulsions, colored extracts, or unusual chemical behavior of a sample
should be noted and considered when reviewing results. Modifications of
a method by an experienced chemist may alleviate some problems. All deviations
from specified methods should be documented in logbooks.
If the continuing calibration (single point) falls outside control
limits, no sample' should be analyzed until the calibration is within these
limits. The standard should be reinjected to confirm the problem and to
discount the possibility of operator error. If still outside of control
limits, the instrument should be recalibrated (multi-point), and at least
the previous sample reanalyzed and results compared. This may indicate
that reanalysis of all samples since last calibration is unecessary.
Data Quality and Reporting--A data summary for each sample should
be submitted. All data should be dry-weight corrected, and reported as
ug/kg using two significant figures. Data should not be blank-corrected.
Spike recoveries, relative percent difference between duplicates, and blank
results (ng/sample)should also be submitted. The following information
is also needed for each sample to allow independent QA review:
t Sample weight extracted
• Percent solids
• Final volume of extract
• Amount of extract injected
• Instrument detection limits
• Method detection limit for each compound
210
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Sed iment
Priority Pollutant Organic Compounds
• Blank associated with sample
• Deviations from the prescribed method
• Problems associated with analysis.
For a more thorough QA review, additional documentation (e.g., chrcmatograms,
computer listings) may be requested.
211
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Sed Imen t
Infauna
Infauna
Field Procedures--
Construction of Sieve Boxes—Sieve boxes should be of sturdy construction,
with high sides to minimize the possibility of material washing out of
the box. They should also be large enough to receive the benthic sample
and wash water without clogging. Swartz (1978) recommends boxes 40 cm
x 40 cm. The boxes should also be constructed to permit nesting of -he
sieves, especially if more than one mesh size will be used. A typical
sieve box might be constructed as in Figure 8. Note the application of
silicone sealant at the mesh wood interface. This sealant will prevent
organisms from crawling into the space where the mesh enters the box frame.
All wood pieces used in construction of the sieve boxes should be treated
with fiberglass or epoxy resin (of the types used in boat building), sanded,
and painted.
It is imperative that the mesh used in the sieve boxes meet specifications
outlined in ASTM E-ll, USA Standard Z23.1, AASHO M92, and Fed. Spec. RR-S-366b.
Such mesh is available from scientific supply houses. Inferior mesh will
not have uniform openings and will not be durable.
Before each cruise, the sieves should be examined for damage and wear.
Look for rips in the mesh, irregular mesh spacing, and sand grains caught
in the mesh. Use water pressure to dislodge the sand. Do not use sharp
objects, as the mesh may be damaged or the mesh spacing may be altered.
Fixative Preparation—The fixative most commonly used for benthic
infauna samples is formalin, an aqueous solution of formaldehyde gas.
Under no circumstances should ethyl or isopropyl alcohol (preservatives)
be used in place of the formalin. Penetration of the alcohol into body
tissues is too slow to prevent decomposition of the specimens.
212
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rXftHitt*tt*Mi%i%*f
f^ilr^fii^afM^m^^m^mlfm-fmft^tflf
•SCREEN LAPS OVER BOTTOM SIOEP1ECE
CONSTRUCTION
1. Construct upper and lowwr box frames (A.B).
2. Nail or stapt* mMh ovw lower tram*.
a Mount upper frame on lower frame.
4. Nail and glue side pieces (C) on upper and
lower frames (A.B). offsetting to permit
sieves to be nested.
& Use waterproof glue (reaortinol. epoxy) and
nails throughout construction.
Figure 8. Construction of a sieve box.
213
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Sed iment
Infauna
Formalin solutions of 5-20 percent strength are recommended for fixing
marine organisms (Gosner 1971; Birkett and Mclntyre 1971; Smith and Carlton
1975; Swartz 1978). Solutions of 10-15 percent are used most commonly.
It is recommended that at least 2 L of diluted formalin solution be on
hand for each replicate sample to be collected, unless experience has shown
otherwise.
The formalin solution should always be buffered to reduce acidity.
Failure to buffer may result in decalcification of mr '.-^scs and echinoderms.
Ideally, pH should be at least 8.2, as calcium carbciate dissolves in more
acidic solutions. Borax (sodium borate, Na£B407) should be used as a buffer
because other buffering agents may hinder identification by leaving a precip-
itate on body tissues and setae.
To prepare a 10-percent buffered formalin solution, add 4 oz of borax
to each gallon of concentrated formalin (i.e., a 40-percent solution of
formaldehyde in water). This amount will be in excess, so use the clear
supernatant when making seawater dilutions. Dilute the concentrate to
a ratio of one part concentrated formalin to nine parts seawater. Seawater
will further buffer the solution. Seawater also makes the fixative isotonic
with the tissues of the animals, thereby decreasing the potential for animal
tissues to swell and break apart, as often happens with freshwater dilutions
of formalin.
It is desirable to prepare fresh fixative prior to each sampling excursion,
as formalin will eventually consume all the buffering capacity of the borax.
This occurs because the formaldehyde gas is in polymer form when in solution.
As the borax neutralizes the excess acidity, the polymer chains become
unstable and break apart, exposing new acid sites. Do not expose formalin
solution of any strength to cold temperatures because the formaldehyde
polymers will degrade into paraformaldehyde and the solution will have
to be discarded.
214
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Sediment
Infauna
Rose Bengal Preparat'on--Rose bengal may be added to the fixed samples
either as a powder or a solution. Both are effective. However, it is
easier, and perhaps less expensive, to use a solution. Simply mix a concen-
trated solution of rose bengal in tap water and put it in a dropper bottle.
The relative quantity of stain being used is much easier to estimate when
dispensed from a dropper than when added in powder form.
Relaxant Preparation—Several relaxants may be used on benthic organisms
prior to -ixation. However, a solution of magnesium chloride in tap water
*s effective on a wide variety of taxa (Gosner 1971), and is easily prepared
and used. The MgCl2 solution should be isotonic with seawater. To prepare,
dissolve 73-g MgCl2*6H20 per liter of tap water. Anhydrous MgCl? can be
purchased (at considerably more cost) and used to prepare the relaxant
solution. However, accurate determinations of mass are very difficult
because of the propensity of the crystals to absorb atmospheric moisture.
Hence, use of the hydra ted form is recommended.
Sample Containers—It is recommended that glass or plastic jars be
used for sample fixation and storage. Plastic lids are preferable to metal
lids because formalin corrodes metal. One- or two-quart containers are
usually adequate for a 0.1-m2 sample after washing. However, more or larger
(i.e., gallon) containers may be required if large quantities of gravel,
peat, wood chips, or other large particles occur in the sample. All sample
containers should be labeled internally and externally using waterproof
materials.
Screen Mesh Selection—It is critical that an appropriate screen mesh
size be chosen for separating the infauna from the sediments. The most
frequently used sieve mesh sizes have been 0.5 mm and 1.0 mm. These sizes
roughly correspond to a trough in the size spectrum for benthic organisms.
This trough occurs at 0.5-1.0 mm and separates meiofauna from macrofauna
(Schwinghaner 1981). Therefore, major deviations from these (or intermediate)
screen sizes will greatly affect the suite of organisms collected.
215
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Sediment
Infauna
Mesh size selection depends largely on the objectives of the study.
Macrofaunal recruitment ana settlement studies definitely require the use
of 0.5-mm mesh, while studies of the distributions and abundances of primarily
adult animals usually require only 1.0-mm mesh. The major disadvantage
of using the larger mesh size is that some small species, particularly
polychaetous Annelids, may be lost. Disadvantages associated with use
of the smaller mesh size are primarily related to processing time. The
residual samples (after washing) v >ntain more detritus and sediment, more
juvenile macrofau.ial forms, and more meiofauna. Therefore, they require
considerably more time to sort. The subadult forms are also much more
difficult to identify, which increases the identification time. It is
not unusual for samples washed on 0.5-mm mesh to require 50 percent more
processing time than sampl.es washed on 1.0-mm mesh. One may argue that
the retention of juveniles permits a more complete characterization of
the community. However, because many of the juveniles that recruit into
a community are rapidly removed through competition or predation, their
inclusion in samples may not represent the more stable adult community
that eventually inhabits a site.
A wise practice 1s to use a set of stacked screens with a 1.0-mm mesh
sieve sitting on top of a 0.5-mm mesh sieve. Both subsets of the sample
can be handled separately, and either or both can be examined based upon
study objectives. Monitoring programs conducted on the west coast of the
United States generally rely on the 1.0-mm analysis, while the 0.5-mm sample
is sometimes archived in case something unusual is found. By contrast,
studies on the east coast of the United States tend to sample the 0.5-mm
fraction more frequently.
The same mesh size should be used consistently through time to permit
temporal comparisons of the biota within the study area. If possible,
the chosen mesh size should be consistent with that used for past studies
in the area and for other studies within the biogeographic region. Such
216
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Sed inent
Infauna
consistency w 11 facilitate biological comparisons with historical data
and with data from adjacent areas.
Collection--Collect benthic samples in accordance with the on-board
procedures for grab sampling discussed earlier.
Proc>iSsing--After qualitative characteristics of the sample have been
recorded, sediments should be washed on the designated sieve(s), using
one of several possible me» tods. Sediments may be gently sprayed with
water from above, agitated by hand in a washtub of water, or washed using
a combination of these techniques. For all methods, it is imperative that
the samples be washed gently to minimize specimen damage. A few minutes
extra care in the field can save hours of time for the taxonomist, and
will result in a better data set.
For most surveys, it is probably easiest to wash the samples from
above with a gentle spray, because efficient, easy-to-use gear may be con-
structed to hold the grab and sieve boxes. An example of a grab stand
is shown in Figure 9. The top section is designed to accept the grab sampler.
Wash water and sediment drain through the openings in the bottom of the
top tray and into the lower section of the sieving stand, where the screen
box(es) is (are) located.
All wash water should be filtered or screened through mesh with openings
one-half the size of those used in the survey, so as not to introduce plank tonic
or bentho-pelagic organisms into the samples. Failure to screen in this
^ay can result in increased sorting time. It can also compromise the quality
of the samples, because it is impossible to distinguish bentho-pelagic
organisms caught by the grab from those entrained in the wash water.
Sieving stands should be designed to fit the specific sampling vessel,
and should never be too large for the available space. Moreover, they
should have attachment points (e.g., eyebolts) at appropriate places with
217
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SPOUT
SIEVE TRAY
EYE SOU-
REFERENCE: STRIPLIN AND HAUPIN (1982)
Figure 9. Example of a sieving stand. Screen boxes
(not shown) are placed in sieve tray.
218
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Sediment
Infauna
wh-ch the stand may be Mshed to the deck or rail. As shown in Figure 9,
all waste water should exit the sieve tray via a spout, to which a hose
should be attached. The wash water can then be discharged overboard through
a scupper. This is especially important in cold weather, when wash waters
may otherwise freeze on the deck and safety may be compromised.
After the contents cf nne grab are emptied into the sieving stand,
the sampler should be washed of all sediments adhering to the inside and
prepared for descent. Take care not to wash a y sediment clinging to the
outside of the grab into the sample. Sediments in the top tray should
be broken apart gently using the spray. Breaking up the sediments in the
upper tray and not 1n the screen boxes eliminates one source by which infaunal
organisms become damaged. Sediments retained on the sieve screens should
be gently washed in the lower section of the sieving stand.
Once sieving is completed, the screen box should be held at an angle
and the remaining material gently washed into one corner. The sample may
then be transferred to an enamel pan or jar for relaxation, or to a jar
for fixation, using as little water as possible. Place the permanent internal
sample label in the enamel pan or jar at this time to avoid confusion later.
If more than one screen fraction 1s generated, be sure to keep them separate
throughout all phases of field and laboratory processing. Be sure to check
the screen for organisms trapped in (or wound around) the mesh wires.
If they cannot be dislodged with gentle water pressure, use a pair of jewelers
forceps. Be careful not to damage the wire mesh. After the screen has
been checked for remaining animals and sample removal is complete, backwash
the screen with a high pressure spray to dislodge any sediment grains that
may be caught in the mesh. Repeat these steps after every sample.
Although relaxation of the organisms in the sample is often omitted
from field sampling programs, 1t is highly recommended for two reasons.
First, relaxed organisms are less likely to distort their shape when fixed,
because the muscles are less likely to contract. The more natural the
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Infauna
appearance of the body, the easier it is to identify the animal. Second,
relaxation helps prevent autotomy, the process whereby organisms (especially
polychaetes) fragment themselves in reaction to the fixative. Complete
animals are much easier to identify than fragments, and are necessary for
the identification of species in some taxonomic groups (i.e., maldanid
polychaetes).
Once the sample is in a jar or enamel pan, it should be ii-wersed in
relaxant (e.g., MgCl2) for about 30 min. After immersion, the sample should
be poured through a small sieve with mesh openings hair the size of those
required for washing the sample. Be sure to recapture the relaxant, as
it may be reused several times (Fauchald 1977). After the animals are
screened, gently transfer the sample to a jar for fixation, using as little
water as possible.
In lieu of using Mgd2 as a relaxant, it may be possible to minimize
damage to the organisms by increasing the strength of the formalin solution
(Word, J., 13 February 1985, personal communication). A strong solution
may kill the organisms before they have time to contract or fragment.
Some experimentation may be needed to determine whether this method is
effective in a given region, and if so, at what formalin concentration.
As mentioned earlier, a 10-percent solution of borax-buffered formalin
is usually used to fix benthic organisms. However, samples containing
large amounts of fine-grained sediments, peat, or woody plant material
require higher concentrations. The sample should be covered with fixative
to a mininum depth of approximately one third that of the sample itself.
Thus, the largest sample that can be placed in a jar occupies only three-
quarters of the available volume. The fixative will occupy the remaining
one-quarter of the volume. The formalin solution should be added to the
sample jar until it is completely filled. This will minimize abrasion
during shipping and handling. If the sample volume exceeds three-quarters
of the jar volume, more than one jar should be used.
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Infauna
Rose bengal may now be added to the sample. Add enough to turn the
fixative cherry-red. This vital stain aids the sorting process by making
animals more visible. However, it renders the body of the organism opaque,
and thereby makes some taxonomic manipulations difficult. For this reason,
the addition of rose bengal is considered optional.
After the formalin end rose bengal (if desired) have been added to
che sample jar, close the container and gently mix the contents f. ensu.-e
that adequate fixative reaches all the organisms. The samples should now
be placed in protective containers for storage and transport to the laboratory.
Wooden boxes with handles are good containers for jars because they are
durable and easily constructed. Bags should be placed in durable water-tight
containers such as snap-lid buckets.
On board ship, samples should be stored so as to minimize exposure
to sunlight and temperature extremes. They should also be stored in a
stable part of the ship to minimize agitation. If special racks are not
available to keep the shipping boxes stationary, they should be lashed
down.
Laboratory Procedures—
Equipment and Supplies—The laboratory should be equipped with both
stereo dissection and compound microscopes. One dissection microscope
capable of magnification to 25-power should be available for each sorter,
and one dissection microscope capable of magnification to 50-power should
be available for each taxonomist. Compound microscopes should be capable
of magnifications up to 1,000-power. At a minimum, one compound microscope
for each three taxonomists is needed; ore compound microscope for each
two taxonomists is preferable. The optics of the dissection and compound
microscopes should be of high quality to reduce sorting and identification
times and to increase acccuracy by reducing the chances for misidentifying
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organisms. Other necessary laboratory supplies include jewelers forceps,
fine scissors, small scalpels, fine needles, flat and depression microscope
slides, cover slips, small dissection trays, immersion oil, and glycerol
alcohol (half glycerol and half 70-percent alcohol).
Preservative Preparation—After the specimens are fixed, alcohol should
be used as a long-term preservative. Either 70-percent ethanol in water
or 70-percent isopropanol may be used. Although isopropanol is less expensive
than ethanol, it is more unpleasant to work wit . Specimens preserved
in isopropanol are unsuitaole for histological examination. If future
studies of anatomy or reproductive biology are anticipated, ethanol must
be used.
It is most cost-effective to purchase isopropanol and ethanol in bulk
solutions of 5-percent water and 95-percent alcohol. Purer grades are
available, but more costly. To prepare 1 L of a 70-percent solution of
either alcohol, add 263 ri. of water to 737 ml of 95-percent alcohol solution.
It may be necessary to use distilled water to dilute the alcohol solution
because hard water mixed with alcohol creates a milky precipitate that
makes examination of the samples difficult. The alcohol solution is most
easily stored in a glass jar with a clamped siphon hose. This apparatus
allows workers to accurately dispense small quantities without wastage.
Use of the 70-percent alcohol /30-percent water solution is adequate
for the preservation of most infaunal organisms. For long-term storage
of crustaceans, however, it is recommended that glycerine be substituted
for some of the water. The glycerine helps keep the exoskeletons supple,
thereby facilitating examination and manipulation. This is especially
critical for crustaceans archived in the reference collection (see below).
An appropriate glycerine-alcohol solution would be 5-percent glycerin in
70-percent alcohol.
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Analytical Procedures--Samp1es should remain in the formalin-seawater
solution for a minimum of 24 h to allow proper fixation (Fauchald 1977).
After 24 h, the samples may be washed (i.e., rescreened) on a sieve with
mesh openings half the size of those used in the field. The smaller screen
size ensures that specimens collected in the field will be retained in
the sample regardless of shrinkage or breakage resulting from contact with
the formalin. It is desirable to wash the formalin from the samples as
soon as possible after the initial 24 h because the buffering capacity
of the bort/ in the formalin solution decreases continually.
If the sample consists of multiple jars, locate all jars prior to
rescreening and wash them at the same time. Carefully pour the contents
of each jar into the appropriately sized screen and rinse the jar to remove
adhering organic material, sediment, or organisms. To reduce the possibility
of sample loss through spilling or splashing, do not fill the screen more
than half full.
Caution should be exercised when handling formalin mixtures because
formalin is toxic and carcinogenic (Kitchens et al. 1976). It can cause
irritation to the eyes, nose, and throat at concentrations as low as 1.0 ppm.
Sensitivity in humans varies with the individual, but in general, the detection
limit is approximately 2 ppm. The technician doing the rescreening should
wear protective clothing, rubber gloves, and safety goggles, and should
work under a properly ventilated fume hood. A protective vapor mask should
be worn, even when working near open windows or under a ventilation hood.
There are several acceptable methods for rinsing formalin from a sample.
One method is to gently flush the sample with large quantities of fresh
water from a low-pressure faucet or hose, being careful not to splash any
sample material. The rinse water should be discharged into a sanitary
sewer. A second method is to partly immerse the sieve in a plastic tub
filled with fresh water and wash the sample by moving the sieve in an up-
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Infauna
and-down motion. Care nust be taken not to let the water rise above the
top of the sieve.
Allow the rinse water to completely drain from the sieve and lightly
rinse the sample with a solution of 70-percent ethanol from a squirt bottle.
Carefully wash the sample material into a sample jar filling it no more
than three-quarters full. Avoid scraping the sample across the screen
with the spoon. Rinse the i*st bit of material into the jar using the
squirt bottle of alcohol. F 1 tne jar to the top with the 70-percent
alcohol solution and screw tne lid on tightly. Gently shake and invert
the jar several times to ensure proper mixing.
Each jar should have one internal label and two external labels.
The internal label should be written with indelible ink on waterproof,
100-percent rag paper. Paper with less than 100-percent rag content or
that is not waterproof will fall apart in the 70-percent alcohol mixture,
and the ink will dissolve. The two external labels should be preprinted
and should be labeled with an indelible marking pen. One label should
be attached to the jar and the second should be attached to the lid of
the jar. All three labels must include all information recorded on the
field data tag, plus all other information needed to ensure proper identifi-
cation of the sample (e.g., if more than one jar, label them 1 of 2, 2
of 2, etc).
Keep all jars of a given sample together (if more than one), and all
replicate samples from a given station together. As the samples are shelved
prior to sorting, each should be cross-referenced to the field log sheet.
At this point the sample custodian should date and initial the rescreening
section of the sample-track ing form for each station. ^Store washed samples
in an upright position, at a cool temperature, and away from direct sunlight.
Storage should be in a secure place, where sample containers are not exposed
to breakage or tampering.
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Several techniques can be used to sort organisms from sediment. The
most common technique involves placing a small amount of the sample into
a glass or plastic petri dish and using a pair of jewelers forceps to sort
through the sample in a systematic manner, removing each organism. This
entire process should be done while viewing the sample through a 10-power
dissecting microscope. Care must be taken that enough liquid is present
in the pet i dish to completely cover the sample, otherwise, reflections
from the sediment/liquid interface will cause distortions and the sorter
may miss some organisms. £* h peu i dish of material should be sorted
twice to oe surb that all organ.sms are removed.
A second sorting technique is a flotation method, which is particularly
effective when the sediment residue is primarily coarse sediment grains
containing small amounts of organic matter (e.g., wood fragments, leaf
debris, sewage sludge). The sample is first washed with fresh water in
a large flat tray. The less dense material that becomes suspended in the
fresh water (organic material, arthropods, and most soft-bodied organisms)
is carefully poured into a sieve, and is sorted using the standard technique
described above. The remaining material is covered with liquid and sorted
using a 5-power self-illuminated hand lens. Organisms remaining in this
portion of the sample generally include molluscs and some tube-dwelling
or encrusting organisms that are associated with sand grains. Because
it is difficult to see extremely small organisms with the 5-power hand
lens, the sorter must remove all molluscs and polychaete tube fragments
for closer inspection. All material collected from this portion is placed
into a labeled sample jar and viewed under a 10-power dissecting microscope
to remove organisms from tubes and to ensure that the molluscs were alive
when captured.
Both sorting techniques expose the sorter to alcohol fumes. Because
these fumes can be irritating to some people, the sorting process can be
done using fresh water. However, as each portion of the sample is sorted,
it should be drained and returned to the alcohol solution immediately.
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Each sample should be sorted by only one person. Organisms should
be sorted Into the following major taxonomic groups: Annelida, Arthropoda,
Mollusca, Echinodermata, and miscellaneous phyla (combined). All organisms
should be placed in large vials containing 70-percent alcohol solution.
The exception is Ophiuroidea, which require air-drying for identification.
Removal of the majority of arms from certain Ophiuroidea (e.g., Amphiuridae)
permits easier identification. This procedure may be performed by experienced
sorters torn nmize identification time. (Note: Special handling of Ophiuroi-
dea should be conducted after biomass analyses, if biomass analyses ure
performed.) Each vial containing a major taxonomic group should have an
internal label listing the survey name, station designation, water depth,
date sampled, and field screen size. All vials in the same sample should
be stored in a common container and immersed in the 70-percent alcohol
solution. To reduce evaporation of alcohol, vial and container lids can
be sealed with plastic tape.
When required, biomass estimates for the major taxonomic groups should
be made prior to identifying the organisms to the species level. However,
it is recommended that taxonomists examine the major taxonomic groups before
biomass measurements are made, to ensure that sorters have correctly grouped
all individuals and fragments. Biomass should be estimated to the nearest
0.1 g (wet weight). All specimens of taxa within the following major groups
should be composited for biomass analyses: Annelida (principally polychaete
worms), Mollusca (principally bivalves, gastropods and aplacophorans),
Arthropoda (principally crustaceans), Echinodermata (principally asteroids,
ophiuroids, echinoids, and holothuroids), and miscellaneous taxa (combined).
These five categories generally are adequate to characterize the standing
stocks of the major infaunal groups. They also are sufficiently distinct
from each other to permit proper assignment of fragments to each of the
groups. All fragments should be placed in their respective major taxonomic
groups prior to weighing.
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There are several major problems associated with the collection end
interpretation of biomass information. Some taxa lose weight when immersed
in preservative fluids, while others gain weight (Howniller 1972; Lappalainen
and Kangas 1975; Wiederholm and Eriksson 1977; Mills et al. 1982). For
this reason, the most accurate biomass estimates are performed on live
material. However, it is rarely practical to sort and weigh live specimens.
Accurate measurements of .iom*ss are further compromised by evaporation
from the specimens while they are on the balance.
Several methods of measuring jiomass are possible. One technique
is to estimate the difference in weight of a tared beaker filled with preser-
vative before and after organisms are placed in the beaker. The individual
organisms are not blotted prior to weighing, and as few individuals as
possible are transferred to the weighing container. These procedures minimize
the transfer of fluids held within a pile of individuals. This technique
can be used for preserved or live animals, and appears to introduce the
least amount of variation into the weighing process.
A second technique for biomass determination consists of air-drying
the organisms on absorbent pa'per for a specific length of time (e.g., 5 min).
Because 70-percent ethanol is volatile, small variations in drying time
may increase the errors associated with the weight measurements. A container
open at one end and covered at the other end with a 0.25-mm mesh screen
(maximum mesh opening) can be used to hold the organisms for weighing.
After the tare weight of the container is measured, the animals are carefully
placed into the container. The container with organisms is then placed
on a paper towel and allowed to air dry for exactly 5 min prior to weighing.
The weight of the organisms is obtained by subtracting the weight of the
container with the organisms from the tare weight of the container. Extremely
large organisms (e.g., large molluscs or asteroids) should be weighed indi-
vidually.
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After biomass estimates are completed, idertification and counting
of the organisms may begin. Identifications shojld be to the lowest taxonomic
level possible, usually the species level. For incomplete specimens, identify
all fragments, but enumerate only the anterior ends (i.e., heads). All
identifications require the use of binocular dissecting and compound micro-
scopes. If possible, at least two pieces of literature should be used
for each species identification, one of which should be tne original descrip-
tion. Moreover, each species identification should be checked \gainst
a reference specimen from a verified voucher museum c"'lection.
After completing an identification, all organisms of a single species
should be placed In a vial containing 70-percent alcohol. All vials should
be stored in a common jar with the original internal label (from the field),
and immersed 1n 70-percent alcohol. Each vial should contain an internal
label with the following information: species name, number of specimens,
survey name, station number, replicate number, collection gear, water depth,
date of collection, latitude, longitude, and initials of the taxonomist.
Any specimens removed fron the sample jar and placed in the reference collection
should be so noted (species, number) on the sample identification sheet.
Each taxonomist should record initial identifications and counts in
a bound, hardcover notebook, which should also include notes and comments
on the organisms in each sample. Upon completion of the sample, the data
should be transferred in ink to the sample data sheets and double-checked.
The taxonomist should then sign and date the sample data sheet. All notebooks
should be kept in the laboratory at all times so the laboratory supervisor
can check questionable identifications and follow the progress of each
sample. Erasures from notebooks or data sheets are not permitted. Simply
draw a horizontal line through incorrect data, initial and date it, and
indicate why the data were removed.
Calibration and Preventive Maintenance—The analytical balance used
for biomass determinations should be calibrated weekly, at a minimum.
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Infauna
The balance and all microscopes should be serviced at rec.ular intervals.
Annual service and inspection is adequate in most cases, unless the manufacturer
recommends otherwise.
Taxonomic identifications should be consistent within a given laboratory,
and with the identifications of other workers. To that end, at least three
individuals of each taxon should be sent for v :r ification to recognized
experts in museums and/or academic institutions. The verified soecimens
should then be placed in the permanent reference museum. Continued co1 •ction
of a verified species does not require additional e.-oert .erif icacion.
Participation of the laboratory staff in a regional taxonomic standardization
program is required (if available) to ensure regional consistency and accuracy
of identifications. The reference collection should be started and completed
as early as possible during the project.
All specimens in the reference collection should be in labeled vials
that are segregated by species and sample. For example, there may be three
labeled vials of Gemma gemma, one from each of three samples. More than
one specimen may be in each vial. The labels placed in these vials should
be the same as those used for specimens in the sample jars. It is important
to complete these labels because future workers may not be familiar with
the survey, station locations, and other details of the work in progress.
In addition, the reverse side of the label should contain information about
the confirmation of the identification by experts in museums or other insti-
tutions (if appropriate). Such Information would Include the outside consul-
tant's name, institution, and date of verification. All vials for a given
species should be placed in a single jar filled with alcohol. To redur
evaporation of alcohol, the lids of vials and jars can be sealed with pi a'
tape. The species (or other taxonomic designation) should be clearly wr
on the outside and on a large internal label. Reference specimen*
be archived alphabetically within major taxonomic groups.
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Infauna
A computer listing of each species name, the name and affiliation
of the person who verified the identification, the location of the individual
specimen in the museum, the status of the sample if it has been loaned
to outside experts, and references to pertinent literature should be maintained
by the laboratory performing identifications for the monitoring program.
This listing should be available near the collection.
Reference specimens are invaluable, and should be retained at the
location where the identifications were perfo, ed, in the offices of the
funding agencies, or at a museum with long-term storage capabilities.
In no instance should this portion of the collection be destroyed. One
person should be identified as the curator of the museum collection and
should be responsible for its integrity. Its upkeep will require periodic
checking to ensure that alcoho> levels are adequate. When refilling the
jars, it is advisable to use full-strength alcohol (i.e., 95 percent),
because the alcohol in the 70-percent solution will tend to evaporate more
rapidly than the water.
Quality Control Checks —It is recommended that at least 10 percent
of each sample be re-sorted for QA/QC purposes. Re-sorting is the exami-
nation of a sample or subsample that has been sorted once and is considered
free of organisms. Each sample should be re-sorted by someone other than
the original sorter. The 10-percent aliquot should be taken after the
entire sample has been spread out in an enamel pan. It is critical that
the aliquot be a representative subsample of the total sample.
A partial re-sorting of every sample should ensure that all gross sorting
errors are detected. In addition, it should give added incentive to sorters
to process every sample accurately. An alternate 10-percent QA/QC approach
of re-sorting one entire sample out of every 10 processed does not have these
advantages. Because 9 out of every 10 samples are not re-sorted, the proba-
bility of randan gross errors going undetected increases, and the incentive
of sorters to process every sample accurately decreases.
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Infauna
In addition to efficient sample sorting, consistent identifications
of organisms among individuals and among sampling programs are critical
to the collection of high quality data. Consistent identifications are
achieved by implementing the procedures discussed below and by maintaining
informal, but constant, interaction among the taxonomists working on each
major group. One important procedure is to verify identifications by camp rison
with the reference collection. To ensure that identifications are correct
and consister. 5 percent of all samples identified by one taxonomist should
be re-identicied by another taxonomist who is also qualified to identi'•/
organisms in that major taxonomic group. Differences in opinion should
be recorded and given to the senior taxonomist. It is the duty of the
senior taxonomist to decide upon the proper identifications. The senior
taxonomist may also decide whether the taxonomic level to which a given
organism is identified is appropriate. If it is not, the senior taxonomist
may decide to drop back to a higher taxonomic level, or to further refine
the taxonomy of that group through additional study.
Three kinds of material should be archived during the course of any
sampling program: sediment residue (after sorting), identified organisms,
and reference specimens. First, the sediment residue (i.e., "grunge")
should be characterized, and a representative aliquot should be archived.
The characterization should include a description of the major sediment
component (e.g., coarse to fine sand, wood, shells, or polychaete tube
fragments, organic material, sewage sludge) and the volume of the total
amount of material. An 8-dram vial filled three-quarters and topped off
with 70-percent alcohol is sufficient for archiving these aliquots. Each
vial should be labeled with the same information found on the internal
sample label. All vials should be-tightly stoppered and stored together
in a container filled with 70-percent alcohol. Black electrical tape wrapped
tightly around the lid of the large container will improve the seal and
insure that the alcohol does not evaporate during storage. These archived
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Infauna
ali'qucts of sediment residue provide a qualitative reference sample should
questions concerning sediment characteristics arise later.
When all identification and QA/QC procedures are completed, the jars
containing the vials of identified species should be topped off with 5-percent
glycerine/7n-percent alcohol. The lids should then be sealed tightly with
black electrical tape to prevent evaporation. All sample iars should be
placed in containers filled with 70-percent alcohol for long-term storage.
The containers should be fitt 1 with a tightly sealed lid, and electrical
tape should again be used to seal the joints. Each container should be
labeled clearly with the survey name,' date, and number and type of samples
within it.
Corrective Action—Foil owing QA/QC procedures discussed earlier, each
sample residue should be checked for complete or nearly complete removal
of organisms. Thus, each sample elicits a decision concerning a possible
re-sort. When a sample is found that does not meet the specified removal
criterion, it should be re-sorted. In addition, the previous three samples
processed by that sorter should be re-sorted (regardless of their aliquot
re-sort results) to ensure that sorting efficiency has been adequate.
If one of the three samples is incompletely sorted, all samples processed
by that individual should be re-sorted.
When a taxonoaic error is found, it is necessary to trace all of the
work of the taxonomist responsible for the error, so as to identify those
samples into which the specific error may have been introduced. This process
can be very time-consuming. However, upon completion of all taxonomic
work, few (if any) taxonomic errors should remain in the data set. Avoiding
errors through the constant interchange of information and ideas among
taxonomists is the best way to minimize lost time due to misidentification.
Data Quality and Reporting—Generally, a sample sorting efficiency
of 95 percent is considered acceptable. That is, no more than 5 percent
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Infauna
of the organisms in a given samp'e are missed by the sorter. Similarly,
specimen identifications can reasonably be expected to be at least 95 percent
accurate. All organisms must be identified to the lowest possible taxon,
and to species level whenever possible. In cases where the identity of
a species is uncertain, a species number will suffice (e.g., Macana sp.l,
Macoma sp.2). Numerical designations must be consistent throughout the
study. Data for each replicate sample should be reported as numbers of
individuals/m2 for each species and as biomass (nearest 1.1 g wet weight/m2)
for each major taxonomic group.
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3IOACCUMULATION/TRAWL ANALYSES
QA/QC procedures are presented in this section for the following tnree
tissue/trawl variables:-
• Priority pollutant metals
• Priority pollutant organic compounds
• Demersal fishes and epibenthic macroinvertebrates.
Or; nlsho to be analyzed for these variables generally will be collected
jsing an otter trawl. Operation of an otter trawl is discussed e-rlie'-
in the general methods section.
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Bioaccumulation
Priority Pollutant Metals
Priority Pollutant Metals
Field Procedures--
Col1ection--The major difficulty in trace metal analyses of tissue
samples is controlling contamination of the sample after collection. In
the field, sources of contamination include sampling gear, grease from
winches or cables, engine exhaust, dust, or ice used for cooling. Care
mijst be taken during handling to avoid these and any otier possible sources
of contamination. For example, during sampling the sli > should be positioned
such that the engine exhausts do not fall on deck. To avoid contamination
from melting ice, the samples should be wrapped in aluminum foil and placed
in watertight plastic bags. The outer skin of the fish or shell of the
shellfish acts as protection against metals contamination from the aluminum
foil.
Sample resection and any subsampling of the organisms should be carried
out in a controlled environment (e.g., dust-free room). In most cases,
this requires that the organisms be transported on ice to a laboratory,
rather than being resected on board the sampling vessel. It is recommended
that whole organisms not be frozen prior to resection if analyses will
be conducted only on selected tissues, because freezing may cause internal
organs to rupture and contaminate other tissue. If organisms are eviscerated
on board the survey vessel, the remaining tissue (e.g., muscle)-may be
wrapped as described above and frozen.
Resection is best performed under "clean room" conditions. The "clean
room" should have positive pressure and filtered air. The "clean room"
should also be entirely metal-free and isolated from all samples high in
contaminants (e.g., hazardous waste). At a minimum, care should be taken
to avoid contamination from dust, instruments, and all materials that may
contact the samples. The best equipment to use for trace metal analyses
is made of quartz, TFE, polypropylene, or polyethylene. Stainless steel
that is resistant to corrosion may be used if necessary. Corrosion-resistant
235
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Bioaccumulatlon
Priority Pollutant Metals
stainless steel is not magnetic, and thus can be distinguished from othe^
stainless steels with a magnet. Stainless steel scalpels have been found
not to contaminate mussel samples (Stephenson 1979). However, low concen-
trations of heavy metals in other biological tissues (e.g., fish muscle)
may be contaminated significantly by any exposure to stainless steel.
Quartz utensils are ideal but expensive. To control contamination when
resecting tissue, separate sets of utensils should be used for removing
outer tissue and for removing tissue for analysis. For bench liners and
bottles, borosilicate glass would be preferred over plastic if tr-ce organic
analyses are to be performed on the same sample.
Resection should be conducted by or under the supervision of a competent
biologist. For fish samples, special care must be taken to avoid contaminating
target tissues (especially muscle) with slime and/or adhering sediment
from the fish exterior (skin) during resection. The incision "troughs"
are subject to such contamination. Thus, they should not be included in
the sample. In the case of muscle, a "core" of tissue is taken from within
the area boarded by the incision troughs, without contacting them. Unless
specifically sought as a sample, the dark muscle tissue that may exist
in the vicinity of the lateral line should not be mixed with the light
muscle tissue that consitutes the rest of the muscle tissue mass.
Prior to use, utensils and bottles should be thoroughly cleaned with
a detergent solution, rinsed with tap water, soaked in acid, and then rinsed
with metal-free water. For quartz, TFE, or glass containers, use 1+1 HNQ-3,
1+1 HC1, or aqua regla (3 parts cone HC1 + 1 part cone HNQ-3) for soaking.
For plastic material, use 1+1 HNQ-3 or 1+1 HC1. Reliable soaking conditions
are 24 h at 700 C (APHA 1985). Do not use chromic acid for cleaning any
materials. Acids used should be at least reagent grade. For metal parts,
clean as stated for glass or plastic, except omit the acid soak step.
If trace organic analyses are to be performed on the same samples, final
rinsing with methylene chloride is acceptable.
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Bioaccumulation
Priority Pollutant Metals
Sample size requirements can vary with tissue type (e.g., liver or
muscle) and detection limit requirements. In general, a minimum sample
size of 6 g (wet weight) is required for the analysis of all priority pollutant
metals. To allow for duplicates, spikes, and required reanalysis, a sample
size of 50 g (wet weight) is recommended. Samples can be stored in glass,
TFE, or high-strength polyethylene jars.
Processing—Samples should be frozen after resection and kept at -200 C.
Although specific holding times have not been recommended by U.S. tPA a
maximum holding time of 6 mo (except for mercury samples, which sho Id
be held a maximum of 28 days) would be consistent with that for water samples.
When a sample is thawed, the associated liquid should be maintained
as a part of the sample. This liquid will contain lipid material. To
arfoid loss of moisture from the sample, partially thawed samples should
be homogenized. Homogenizers used to grind the tissue should have tantalum
or titanium parts rather than stainless steel parts. Stainless steel blades
used during homogenization have been found to be a source of nickel and
chromium contamination.
Laboratory Procedures--
Analytical Procedures—Priority pollutant metals should be analyzed
according to the procedures described in Tetra Tech (1986b). Digest the
homogenized sample prior to analysis, using the acids specified in the
procedure. The digestate can then be analyzed by flame Atomic Absorption
Spectrophotometry (AAS), graphite furnace AAS, or Inductively Coupled Plasma
(ICP), depending on the sample concentration and required detection limits.
ICP can be used to screen samples for elements that are present in relatively
high concentrations. Most elements will require more sensitive analysis
by graphite furnace AAS. Mercury analysis must be performed by cold vapor
AAS.
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Bioaccumulation
Priority Pollutant Metals
Calibration and Preventive Maintenance—Because contamination can
be the limiting factor for reliable quantitation of trace metals in tissue,
several field and method blanks should be analyzed to establish contamination
variability. At least three blanks, or a blank every 20 samples, should
be collected at each step of sample preparation to evaluate contamination
variability. The minimum blanks to analyze are:
a Bottle blanks Bottles should be opened and rinsed with
concentrated HN03. The HN03 rin-,e should be digested and
analyzed with each batch of samples
0 Resection blanks - After routinely cleaning the utensils
used for dissecting and before resecting the next sample,
the utensils that have contacted the tissue to be analyzed
should be rinsed with metal-free water. A rinsate should
be collected and the volume recorded. This rinsate blank
should be analyzed at least once and preferably one blank
with each batch of samples. The total micrograms of each
element in the rinsate should be recorded. This blank will
allow evaluation of the cleaning technique and provide correction
values If necessary.
• Digestion (or method) blanks - Blanks containing the same
quantity of acid as each sample should be digested and analyzed
with each batch of samples.
Preliminary blanks and samples should be analyzed prior to resecting and
analyzing the tissue samples to ensure that the blank concentrations and
variability are low enough to establish adequate assessment of tissue concen-
trations. Blanks should continue to be analyzed with each batch of samples
to monitor sporadic contamination.
All analytical instruments must be calibrated daily and each time
the instrument is set up. Calibration procedures should follow those for
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Priority Pollutant Metals
t^e specified method for each analysis. Calibration standards must be
prepared using the same concentrations of acids as those which will result
in the samples following sample preparations.
After an instrument has been calibrated, the accuracy of the initial
calibration should be verified by the analysis of certified control solutions
at a frequency of 5 percent or every 2 h during an analysis run, whichever
is more frequent, and after running the last analytical sample. If a certif .ed
control solution is not available, a standard should oe used. Analyte
for the standard ;hould be from a different source than that used in the
standards for tlie initial calibration. If the deviation of the continuing
calibration verification is greater than the calibration control limits
specified in the method, the instrument must be recalibrated, and the preceding
10 samples reanalyzed.
All equipment should have scheduled routine preventive maintenance,
and a record of all maintenance performed should be noted in a logbook.
Critical spare parts should be kept on hand.
Quality Control Checks—Analyze appropriate standard reference materials
(SRM) if available (e.g., U.S. EPA Trace Metals in Fish Tissue, NBS Oyster
Tissue, or, if analyzing fish liver, NBS Bovine Liver) to provide a check
on contamination, digestion efficiency, and overall accuracy of the analysis.
A minimum of one SRM should be analyzed for each survey or 2 percent of
the total number of samples (i.e., 1 per 50 samples), whichever is more
frequent).
To estimate precision, a minimum of 5 percent of the total number
of samples should be analyzed in duplicate or one for each survey, whichever
is more frequent. When more than 20 samples are to be analyzed for one
survey, the project manager may choose to implement a program of triplicate
analyses. The overall percentage of replicates should be at least 5 percent.
Analyze samples spiked before digestion at the same frequency as duplicates
to estimate recovery. Add spike concentrations approximately equal to
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Bloaccumulation
Priority Pollutant Metals
tne concentration found in the unspiked sample. An acceptable range in
spine concentrations is 0.5 to 5 times the sample concentrations. Limited
sample size may necessitate reduction in the number of QC samples for bioac-
cumulation monitoring.
The blanks listed under Calibration Procedures and Preventive Maintenance
should be carried through all the appropriate steps. Ideally, the concen-
trations of the bottle blanks, dissection blanks, ancj digestion blanks
are less than the detectir • lim-t and no correction of sample results is
performed. If there is extensive contamination of blanks (>30 percent
of sample value) and if there is enough sample, the batch of samples associated
with the blank should be reanalyzed. The data should be corrected by the
data users for the blank values between the required detection limit and
the control limit.
For ICP analysis, additional QC checks should include an interference
check sample to verify interelement and background correction factors.
For graphite furnace AAS, additional QC checks should include duplicate
Injections with the mean value reported. The relative standard deviation
of the readings should be within control limits, or the sample should be
rerun. If the predigestion spike recovery is poor, the digestate should
be spiked and results compared to the predigestion spike results. If the
digestate spike recovery is as poor as the predigestion recovery, the method
of standard additions should be used to determine digestate concentrations.
Corrective Action—If the concentration of the field or method blank
is greater than the required detection limit, then all steps in the sample
handling should be reviewed. Many trace metal contamination problems are
due to airborne dust. Thorough cleaning of all utensils between processing
each sample is important to minimize cross-contamination. High zinc blanks
may be due to airborne dust or galvanized iron, while high chromium and
nickel blanks indicate contamination from stainless steel. In the field,
the use of mercury-filled thermometers should be avoided. Broken thermometers
can be a potential source of severe mercury contamination. In the laboratory,
240
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Bioaccumulation
Priority Pollutant Metals
samples for mercury analysis should be isolated from items such as polarographs
or COO reagents.
Poor replication may be caused by inadequate mixing of the sample
before taking aliquots, inconsistent contamination, inconsistent digestion
procedures, or instrumentation problems.
Poor performance on the analysis of the Standard Reference Material
(SRM) or poor spike recovery may be caused for the same reasons as poor
duplication. However, if duplica. > results are acceptable, then poor SRM
performance or poor spike recovery may be caused by loss of analyte during
digestion or sample matrix interferences during analysis. To check for
analyte loss during digestion or for low recovery due to interferences
during analysis, spike the sample after digestion and compare the analysis
to the predigestion spike. If the results are different, then the digestion
technique should be adjusted. If the results are the same, then dilute
the sample by at least a factor of five and reanalyze. If spike recovery
is still poor, then the method of standard additions, a matrix modifier,
or another method is required.
Sediment on the outside of shellfish may contaminate the tissue to
be analyzed. Proper cleaning of the outside of the animal is critical.
Data Quality and Reporting—Measurements should be reported as ug/kg
to a maximum of three significant figures on a wet-weight basis. Detection
limits can vary widely because of methods and Instrumentation. The analytical
method should be consulted to determine expected detection limits, precision,
and accuracy. Further guidance on analytical detection limits can be found
Tetra Tech (1985b). '
The data report should include duplicate, spike, and blank results.
Data should not be blank-corrected. In addition, the laboratory data summary
should include:
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Bioaccumulation
Priority Pollutant Metals
• Digestion procedures
0 Quantity of sample digested and final dilution volume
• Instrument detection limit for each element
t Blank associated with sample
• Deviation from the prescribed methods
• Problems associated with analysis.
For a more thorough QA review, additional documentation (e.g., calibration
curves) may be requested.
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Bloaccumulation
Priority Pollutant Organic Compounds
Priority Pollutant Organic Compounds
Field Procedures--
Col1ection--In the field, sources of contamination include sampling
gear, grease from winches or cables, engine exhaust, dust, and ice used
for cooling. Efforts should be made to minimize handling and to avoid
sources of contamination. This will usually require resection of tissue
to be performed in a controlled environment (e.g., laboratory). For examp'.c
to avoid contamination from ice, the samplps should be w jpped in aluminimum
foil, placed in watertight plastic bags and immediately iced. Organisms
should not be frozen prior to resection if analyses will be conducted only
on selected tissues because freezing may cause internal organs to rupture
and contaminate other tissue. If organisms are eviscerated on board the
survey vessel, the remaining tissue (e.g., muscle) may be-wrapped as described
above and frozen. Limited sample size can be a problem when using small
organisms, but an attempt should be made to collect at least 30 g of tissue
for each organic priority pollutant analysis. Additional sample (30 g)
is needed for each spike, duplicate, reanalysis, and archive sample.
Processing—To avoid cross-contamination, all equipment used in sample
handling should be thoroughly cleaned before each sample is processed.
All instruments must be of a material that can be easily cleaned (e.g.,
stainless steel, anodized aluminum, or borosilicate glass). Before the
next sample is processed, instruments should be washed with a detergent
solution, rinsed with tap water, soaked in high-purity methylene chloride,
and finally rinsed with reagent water. Work surfaces should be cleaned
with 95-percent ethanol, and allowed to dry completely. If metals are to
be analyzed, instruments may be washed with acid prior to the methylene-
chloride rinse. Each fish, crab, and mollusc should be handled with clean
stainless steel or quartz instruments (except for external surfaces).
The specimens should come into contact with precleaned glass surfaces only.
Polypropylene and polyethylene surfaces are a potential source of contamination
and should not be used. To control contamination when dissecting tissue,
243
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Bioaccumulation
Priority Pollutant Organic Compounds
separate sets of utensils should be used for removing outer tissue and
for dissecting tissue for analysis. For fish samples, special care must
be taken to avoid contaminating target tissues (especially muscle) with
si ire and/or adhering sediment from the fish exterior (skin) during resection.
The incision "troughs" are subject to such contamination. Thus, they should
not be included in the sample. In the case of muscle, a "core" of tissue
is taken from within the area boarded by thp incision troughs, without
contacting them. Unless specifically sought as a sample, the dark muscle
tissue that may exist in the vicinity of the lateral l1050 C may be substituted for
the solvent rinse if only volatile organics are to be analyzed.
The U.S. EPA and other federal agencies (e.g., National Bureau of
Standards) have not jet provided specific guidance regarding holding times
and temperatures for tissue samples to be analyzed for semi-volatile organic
compounds. Until U.S. EPA develops definitive guidance, the following
holding conditions should be observed. Resected tissue samples should
be maintained at -200 C and extracted as soon as possible, but within 10
days of sample receipt. Complete analyses should be performed within 40
days.
When possible, the entire sample should be used for analysis. There
are several acceptable methods of homogenization of a thawed tissue sample.
If a micro-grinder (e.g., a Tissuemizer) is used, care must be taken to
244
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Bioaccumulation
Priority Pollutant Organic Compounds
ensure that it is thoroughly cleaned between each use. This usually entails
disassembly of the unit. A scapel can be used to cut the tissue into small
pieces. The method chosen will depend upon sample size, time, and resources
available. Devices with large surface areas (i.e., blenders, meat grinders)
should be avoided, as they are difficult to clean and a small sample is
difficult to remove after grinding.
Laboratory Procedures--
Analytical Procedures--Priority pollutant organic compounds shouU
be analyzed according to the procedures described in Tetra Tech (1986b).
Preparation of the tissue will depend upon sample size, extraction method,
and sample type (fish muscle, whole organism, or composite). Avoid use
of plastic or rubber equipment that may come in contact with the sample,
as these can be a source of contamination.
A modified U.S. EPA Method 624 (Hiatt 1981) produces adequate spike
recoveries and detection limits for volatile compounds. Analysis of the
semi-volatile organic compounds involves a solvent extraction and one or
more cleanup procedures.
A Soxhlet extraction with at least a gel permeation column cleanup
provides efficient extraction of organic compounds of interest and eliminates
many interfering coextracted materials.
Cross-contamination should be avoided during all steps of analysis.
All glassware should be clean. Injection micro-syringes must be cleaned
well between uses. If separate syringes are used for injection of standard
solutions, bias between syringes should be accounted. Carryovers can occur
when high- and low-level samples are analyzed sequentially. Analysis of
an appropriate solvent blank following a high-level sample may be necessary
to check for carryover.
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Bioaccumulation
Priority Pollutant Organic Compounds
Recommended detection limits are based on a minimum 20-g wet-weight
sample size with an additional 10 g (wet weight) required for a separate
analysis of volatile compounds. For most of the volatile organic compounds,
detection limits between 5 and 10 ug/kg wet weight are recommended. Detection
limits of 10 ug/kg (wet weight) are recommended for aromatic hydrocarbons
and phthalates analyzed by GC/MS. Detection limits of 10-20 ug/kg (wet
weight) are recommended for chlorinated hydrocarbons and halogenated ethers
analyzed by GC/MS. Detection limits of 50 ug/kg (wet weight) are recommended
for pesticides analyzed by GC/MS. A detection limit of 20 ug/kg (wet weight)
is recommended for PCBs analyzed by GC/Et . If pesticides are analyzed
by GC/EC, a detection limit in the range of 0.1-5 ug/kg (wet weight) is
recoimended (Tetra Tech 1985b).
Actual attainable detection limits will vary with sample size, cleanup,
extraction method, final volume, and amount and nature of any co-extractive
compounds.
Calibration and Preventive Maintenance—Before beginning analysis
of samples, a calibration curve bracketing the working range must be estab-
lished. This calibration should be repeated after all major equipment
disruptions. Calibration checks of the GC/MS system at the beginning of
the day and at least every 12 h will establish that the instrument's response
is in control. Specific tuning criteria (e.g., OFTPP, BFB) are given in
each method. Calibration checks of the GC/ECO system should be done at
the beginning of the day and verified at least every 6 h. These are only
recommended minimum frequencies and the nature of the samples may necessitate
more frequent verification.
A routine QC check for each lot of analytical reagents used in extraction
can monitor a potentially serious source of contamination. Also, each
lot of alumina, silica gel, sodium sulfate, or Florasil used in extract
cleanup should be tested and cleaned as necessary. Any resins or charcoals
used should also be monitored. Surrogate mixtures have also been found
to contain impurities and should be verified prior to use. The fatty acid
246
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Bioaccumulatlon
Priority Pollutant Organic Compounds
content of many tissue samples may overload the cleanup columns. The columns
should be calibrated and monitored to enable the compounds of interest
to be consistently collected in the proper fraction.
Equipment should be maintained and serviced routinely according to
manufacturer's instructions and good laboratory practices. Logbook records
should be kept to document maintenance for each measurement device.
Total Extract.^ble Lipids--The amount of tissue extract required to
determine total ext actable lipid content will depend upon the sensitivity
of the balance used and the amount of lipid in the organism. A maximum
of 1/200 of the original extract from 20 g of tissue should be sufficient
for this determination. Ideally, using an electrobalance able to measure
to 0.1 ug, 1/10,000 of a 20-g extract is more than sufficient. A volume
should be used so as to measure approximately 50 times the smallest unit
i
measured by the balance. If a large percentage of the extract is used
for lipid determination (e.g., >1/200), the results of further analyses
on the remaining extract should be adjusted accordingly for this loss of
material.
Quality Control Checks—To monitor precision and accuracy, duplicate
and matrix spike analyses should be performed. A frequency of 5 percent
of each or one each per survey, whichever is more frequent, is the recommended
minimum. When more than 20 samples are to be analyzed for one survey,
the project manager may choose to implement a program of triplicate analyses.
The overall percentage of replicates should be at least 5 percent. Method
interferences can be caused by contaminated glassware, reagents, solvents,
or processing hardware. These materials can be monitored for contamination
by processing of 5-percent method blanks, or one per batch, whichever is
more frequent. Addition of known amounts of surrogate compounds to each
sample before purging or extraction will serve to monitor preparation and
analysis of samples. An empty tissue jar processed, handled, and stored
as a sample should serve as a field blank. As another measure of accuracy,
a standard reference material should be analyzed if available. One for
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Bioaccumulation
Priority Pollutant Organic Compounds
each survey or 2 percent of the total number of samples, whichever is more
frequent is the recommended minimum.
Corrective Action—When results of QC samples fall outside of established
limits, several courses of action are available. Contamination in the
lab reagent blank is cause for all positive findings of the same compound
in associated samples to be suspect. If contamination is extensive, reanalysis
of the whole associated group of samples may be in order. Blank contamination
should be kept to less ' nan 10 percent of the sample values and, preferably,
below th2 detection IT nit. Contamination found in the field blank should
also be considered when looking at associated sample data. Extensive contam-
ination of a lab or field blank (>30 percent of sample value) should lead
to a detailed review of laboratory, sampling, transport, and storage pro-
cedures. Phthalates, methylene chloride, and toluene are common laboratory
contaminants that may be detected in blanks above the method detection
1imit.
Poor duplication may be caused by inadequate mixing of the sample
before removing aliquots, inconsistent contamination, inconsistent extraction
procedures, or instrument problems. Further replication of an analysis
may be necessary to determine the reason for the poor results.
Poor spike recovery may be caused for the same reasons as poor duplication,
or by matrix effects produced by co-extracted materials. If the spiked
compound is added at a concentration much less than that found in the sample,
recovery may be difficult to determine. This problem is difficult to avoid,
as most environmental samples contain unknown concentrations of organic
compounds. To check if the analyte is being lost somewhere during the
processing, a step-by-step examination of the method using a spiked blank
is necessary, with measurements of the analyte at each step.
Emulsions, colored extracts, or unusual chemical behavior of a sample
should be noted and considered when reviewing results. Modification of
a method by an experienced chemist may alleviate some problems (e.g., substi-
248
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Bioaccumulation
Priority Pollutant Organic Compounds
tuting ethyl ether for the usual solvent can sometimes >ielp emulsions with
tissue samples). All deviations from specified methods snould be documented
in logbooks and discussed in the data report.
Results that fall outside the established calibration curve are suspect
until linearity of response can be shown at that concentration or until
the extract is diluted appropriately and reanalyzed. Extreme values may
necessitate re-extraction of a smaller sample size. Extremely high concen-
trations of organic compounds may saturate the extraction capabilities
of the method and may necessitate re-t .traction of a smaller sample size
or use of a more appropriate method. Chromatographic interference by natural
substances present in the tissue (e.g., fatty acids) may require additional
cleanup.
Data Quality and Reporting—A data summary for each sample should
be submitted. All data should be reported as ug/kg wet weight to a minimum
of two significant figures. Data should not be blank-corrected. Spike
and surrogate recoveries, relative percent difference between duplicates,
and blank results (ng/sample) should also be submitted. The following
additional information is needed for each sample to allow independent QA
review:
0 Sample weight extracted
• Final volume of extract
• Amount of extract injected
• Instrument detection limits
• Detection limit for each compound
• Blank associated with the sample
249
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Bioaccumulation
Priority Pollutant Organic Compounds
• Deviations from the prescribed method
t Problems associated with analysis.
For a more thorough QA review, additional documentation (e.g., chromato-
grams, computer listings) may be requested.
250
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Demersal Fishes and Large Macroinvertebrates
Demersal Fishes and Large Macro invertebrates
Field Procedures—
Collection--After the trawl assembly has been brought on board, the
cod-end choker can be released and the catch emptied into some kind of
collection vessel (e.g., tub, tray, bucket, special table). It is reconnended
that a collection vessel be used to prevent organisms from escaping and
to prevent possible contamination of tissue-contaminant specimens by exposure
to the deck surface.
If organisms from the trawl catch are to be subsampled for tissue
contaminant analyses, these individuals should be noted and removed from
the sample before the remainder is processed for ecological information.
Care should be taken to minimize potential contamination of these specimens.
For example, the deck and all collection containers or equipment should
be washed before and after each haul. Smoking should not be allowed during
sampling and all crew members should wear clean gloves when handling the
organisms. It is also helpful if the skipper positions the vessel so that
stack gases move away from the fantall. Additional precautions are given
in the bioaccumulation sections of this document that describe the detailed
collection methods to be used for specific kinds of tissue analyses.
Processing--Once tissue-contaminant specimens are removed, the remainder
of the catch can be processed without concern for contamination. All organisms
should be Identified and counted. Identifications should be to the species
level, if possible. It is advisable to have a collection of taxonomic
keys to local fauna on board the sampling vessel. For fishes, scientific
and cannon nanes should conform to those recommended by Robins et al. (1980).
If the identity of a species is in doubt, several representative specimens
should be frozen or fixed with 10-percent buffered formalin and identified
by an expert after the cruise.
251
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Demersal Fishes and Large Macroinvertebrates
Following Identification, the size of each individual should be measured.
For fishes, it is recommended that total length (TL) be determined to the
nearest millimeter (mm). Total length is the length from the anterior-
most part of the fish to the tip of the longest caudal fin rays. Two kinds
of total length can be measured (Anderson and Gutreuter 1983). Maximum
TL is determined when the lobes of the caudal fin are compressed dorso-
ventrally, whereas natural TL is measured when the caudal fin is in its
natural state. To be consistent with the :onvention used by most fishery
investigations in the U.S., maximum TL should be measured (Anderson and
Gutreuter 1983).
For fishes that occur in relatively large numbers, individuals may
be classified into size classes rather than measured exactly. This procedure
can reduce processing time considerably. If size classes are used, it
is recommended that they differ by no more than 1 cm each. Furthermore,
it is recommended that each size class be measured from one half-cm value
to the next. For example, a 10-cm size would include fish from 9.50 to
10.49 on.
In some cases, erosion of the caudal fin in a substantial segment
of a population may require that a measurement other than total length
be used for affected individuals. If this occurs, it is recommended that
maximum standard length (SL) be used as a substitute. Standard length
is the length from the anterior-most part of the fish to the posterior
end of the hypural bone. Anderson and Gutreuter (1983) state that in practice,
SL may be measured to some external feature such as the last lateral line
scale, the end of the fleshy caudal peduncle, or the midline of a crease
that forms when the tail is bent sharply. Standard length can be related
to total length by developing a regression relationship between these two
measures for a sample that covers the complete lengch range observed in
the population.
As each individual is measured, it should also be scanned carefully
for grossly visible external abnormalities. Some common external abnormalities
252
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Demersal Fishes and Large Macro invertebrates
found in fishes captured in polluted areas are listed in Table 7. Recent
literature regarding this subject is reviewed by Sindermann (1979). At
least one person on board the sampling vessel should be capable of diagnosing
the abnormalities listed in Table 7. If there is any doubt as to the identity
of an abnormality, then one, or preferably several, specimens afflicted
with the disorder should be frozen or fixed with 10-percent buffered formalin
and taken to an expert for accurate diagnosis. Once disorders have been
confirmed, it is helpful to maintaizn reference specimens on the sampling
vessel or in a shore-based laboratory.
Perhaps the most frequent error associated with recognizing external
abnormalities is mistaking net-related damage as integumental lesions.
This problem occurs most frequently when diagnosing ulcers and fin erosion.
Net-related damage can result from surface abrasion as captured fishes
are dragged along the bottom in the cod end or as .fishes swim against the
net in an attempt to escape. Net-related damage can also result from captured
fishes biting each other. Whenever a suspected lesion appears to be newly
formed or is actively bleeding, net-related damage should be suspected.
Laboratory Procedures--
There are no laboratory procedures for demersal fishes and megainverte-
brates, as all processing will occur in the field.
253
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TABLE 7. SOME COMMON EXTERNAL ABNORMALITIES OBSERVED
IN FISHES FROM POLLUTED AREAS
Integumental Lesions
Fin erosion (fin rot)
Lymphocystis
Ulcers
Neoplasms (tumors)
Skeletal Malformations
Scoliosis
Lordosis
Cranial compresc, in (pugheadedness)
thv.rfism
Deformed fin rays (bent fins)
Pigmentation Anomalies
Ambicol oration
Parasites
Copepods
Isopods
Nematodes
Leeches
254
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structural indices, pp. 283-300. In: Fisheries Techniques. L.A. Nielsen
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Bcrdner, R., J. Winter, and P. Scarpino. 1978. Microbiological methods
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Buchanan, J.B., and J.M. Kain. 1971. Measurement of the physical and
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Oisalvo, L., H.E. Guard, and N.D. Hirsch. 1977. Assessment and significance
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Elliott, J.M. 1971. Some methods for the statistical analysis of samples
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Fauchald, K. 1977. The polychaete worms; definitions and keys to the
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Folk, R.L. 1968. Petrology of sedimentary rocks. University of Texas,
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Glbbs, P.J.. and J. Mathews. 1982. Analysis of experimental trawling
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Gosner, K.L. 1971. Guide to identification of marine and estuarine inver-
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Green, E., 0. Schnitker. 1974. The direct titration of water-soluble
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Greenberg, A.E., R. Trusseil, and L. S. Clersceri (eds). 1985. Standard
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Hiatt, M.H. 1981. Analysis of fish and sediment for volatile priority
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Agency, Washington, DC.
Holme, N.A. 1971. Macroinfauna sampling, pp. 80-130. In: Methods for
f.-cudy of Marine Benthos. N.A. Holme and A.D. Mclntyre (eds). IBP Handbook
No. 16. Blackwell Scientific Publications, Oxford, UK.
Howmiller, R.P. 197 . Effects of preservatives on weights of some common
macrc'^nthic invertel-rates. Trans Am. Fish Soc. 4:743-746.
Kitchens, J.F., R.E. Casner, S.S. Edwards, W.E. Harward III, and B.J. Macri.
1976. Investigation of selected potential environmental contaminants:
formaldehyde. EPA-560/2-76-009. U.S. Environmental Protection Agency,
Washington, DC.
Lappalainen, A., and P. Kangas. 1975. Littoral benthos of the northern
Baltic Sea. II. Interrelationships of wet, dry, and ash-free weights of
macroinfauna In the Tvarminne area. Int. Rev. Gesamten. Hydrobiol. 60:297-312.
Mearns, A.J., and J.M. Allen. 1978. Use of small otter trawls in coastal
biological surveys. EPA-600/3-78-083. U.S. Environmental Protection Agency,
Corvallis, OR. 33 pp.
Metcalf 4 Eddy, Inc. 1979. Wastewater engineering: treatment/disposal/reuse,
Second Edition. McGraw H111 Book Company, New York, NY. 920 pp.
Mills, E.L., K. Pittman, and B. Munroe. 1982. Effect of preservation
on the weight of marine benthic invertebrates. Can. J. Fish. Aquat. Sci.
39:221-224. -
Plumb, R.H., Jr. 1981. Procedures for handling and chemical analysis
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Criteria for Dredged and Fill Material, U.S. Army Waterways Experiment
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Robins, C.R., R.M. Bailey, C.E. Bond, J.R. Brooker, E.A. Lachner, R.N. Lea,
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Schwinghamer, P. 1981. Characteristic size distributions of integral
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256
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Shepard, P.P. 1954. Nomenclature based on sand-siU-clay ratios. J. Sedi-
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Shepard, P.P. 1963. Submarine geology. (Second edition). Harper and
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Sindermann, C.J. 1979. Pollution-associated diseases and abnormalities
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257
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258
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259
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GLOSSARY
a*b - Notation for acid and alkali solutions. In additive volumes (a+b),
tne first number, a,.refers to the volume of a concentrated reagent; the
second number, b, refers to the volume of distilled water required for
Dilution.
Accuracy - The closeness of a measured or computed value to its true value.
Aliquot - A divisor that divides a sample into a number of equal parts,
leaving no remainder; a sample resulting from such a divisor.
Ambient - Surrounding, encircling.
Analyte - The specific component measured in a chemical analysis.
APHA - American Public Health Association.
Apparent Particle Size Distribution - Distribution comprised of both the
inorganic and organic particles in a sample (i.e., organic material is
not oxidized).
Archive • A repository of evidence or information.
Asymptote - A line considered a limit to a curve in the sense that the
perpendicular distance from a moving point on the curve to the line approaches
zero as the point moves an infinite distance from the origin.
Auto-Injection - An automated introduction of a sample into an instrument
for analysis. Usually used for extracts analyzed by gas chromatography.
Background Correction Factor - A number that adjusts the analyte signal
for interfering matrix effects in flame and graphite furnace AA analysis
(usually done automatically by the instrument itself through the use of
continuous deuterium lamp).
Bacteriostasis • The arrestment or inhibition of bacterial growth and repro-
duction.
Batch - Usually refers to the number of samples that can be prepared or
analyzed at one time. A typical batch size is 20 for extraction of organic
com pounds.
Benthic - That portion of the marine environment inhabited by organisms
that live permanently in or on the bottom.
Bioaccumulation - The accumulation of chemical substances in the tissues
of organisms.
260
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Biomass - The weight of living material in all or part of an organism,
population, or community. Commonly expressed as weight per unit area,
a biomass density.
Blank-Corrected • The concentration of an analyte adjusted for the concentration
of that analyte in the blank.
Bow Wake - The pressure wave that forms below a solid object as it is lowered
through the water column.
Bridle - The line that links each otter board with the hydrowire.
Calibration - The systematic standardization of the graduations of the
quantitative measuring instrument.
Carryover - Contamination arising from previous analysis or extraction
of a standard or highly contaminated sample.
Chain-of-Custody - The procedures and forms used to trace and record environ-
mental samples through all stages of collection, shipping, analysis, and
final disposition.
Chelation - The addition of organic complex ing agents, such as EDTA (ethylene-
diaminetetraacetic acid) that preferentially bind with metals, thereby
reducing the exposure and possible toxicity to organisms (e.g., bacteria).
Chief Scientist - The person in charge of the sampling team on a research
vessel.
Cleanup - The removal of co-extracted compounds that may cause interference
from a sample extract.
Cod End - The mesh bag at the back of an otter trawl in which the catch
is collected. It is often of smaller mesh size than the remainder of the
net.
Coefficient of Variation - The standard deviation expressed as a percentage
of the mean.
Co-Extractive - Materials other than analytes of interest that are extracted
along with the analytes. These can be sources of interference.
Composite Sample - A sample composed of two or more grab samples or increments.
Contingency Plans - Procedures to be followed when those that were planned
originally cannot be carried out.
Copenhagen Seawater - Seawater of known chlorinity, an international standard
for salinity determinations.
Corrective Action - Measures taken to remove, adjust, remedy, or counteract
a malfunction or error so that a standard or required condition is met.
261
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Ooss-Contamination - Contamination of a sample or sample extract from
ejoosure to another sample or sample extract, usually of higher concentration.
Demersal - Living on or near the bottom of the sea.
Desiccation - T^e thorough drying of a sample by removal of moisture.
Detection Limit - The smallest concentration of some component of interest
tnat can be measured by a single measurement with a stated level of confidence.
Digestion - In preparing samples for analysis of metals, an acidic solution
added to break organometall ic bonds, freeing the metals for analysis by
atomic absorption or atomic emission spectrophotometry.
Distillation - The vuporization of a liquid mixture with subsequent collection
o* components by differential cooling to condensation.
Duplicate Analysis - A second analyi s made on the same (or identical)
sample of material to a_3ist in. the evaluation of measurement variance.
Effluent - Something that flows out. For example, the liquid material
discharged by sewage treatment plants.
Electrolyte - A substance that dissociates into ions In solution or when
fused, thereby becoming electrically conducting.
Emulsion - A suspension of small globules of one liquid in a second liquid
with which the first will not mix.
Epibenthic - Residing primarily on the sediment surface.
Equilibrium - The state of a reaction in which its forward and reverse
reactions occur at equal rates so that the concentration of the reactants
does not change with time.
Extraction - A method of separation in which a solid or solution is contacted
with a liquid solvent to transfer one or more component into the solvent.
False Positive - A positive measurement of an analyte* not attributable
to the sample.
Field Blank - An empty container or uncontaminated representative matrix
carried through the field routine in the same manner as a sample.
Filter Blank - An unused filter extracted and analyzed in the same manner
as filters used to collect samples. The filter should be prepared in the
same manner as sample filters.
Fixation - The process of putting something into a stable or unalterable
form.
Footrope - The line that forms the front edge of the bottom of an otter
trawl. Weight is often added to it to keep it on the bottom when being
towed .
262
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Formalin - A trademark for a 37 percent by weight aqueous solution of formal-
dehyde with some methanol.
Formazin - A polymer used as a reference standard suspension for turbidity
measurements.
Grab Sample - 1) see increment, or 2) a sample of bottom sediment collected
by a grab sampler.
Gravimetric - Of or pertaining to measurement by weight.
Headrope - The line that forms the front edge of the top of an otter trawl .
Floats are attached to it to hold the net open when being towed.
Keadspace - The airspace between a collected sample and the container lid.
Hydr. wire - The cable used to deploy equipment over the side of a vessel.
It usually is attached to a winch at one end and to the piece of equipment
at the other end.
Hygroscopic - Readily absorbing moisture, as from the atmosphere.
Increment - An individual portion of material collected by a single operation
of a sampling device, from parts of a lot separated in time or space.
Increments may be either tested individually or combined'(composited) and
tested as a unit.
In Situ - In something's original place.
Interelement Correction Factor - A number that adjusts the analyte signal
for the interfering effects of other elements present in a sample undergoing
ICP analysis (usually done automatically by the instrument itself).
Interference - A substance present in the sample that impedes the accurate
measurement of an analyte of interest.
Interference Check Sample - A solution contaminating both interfering and
analyte elements of known concentration that can be used to verify background
and interelement correction factors in ICP analysis.
In Vivo - Within the living organism.
Isobath - A contour of constant depth.
Macroinvertebrate - An invertebrate retained by a sieve having a mesh size
of 1 mm.
Matrix Spike Compound - A known amount of an analyte added to a sample,
usually prior to extraction or digestion.
Method Blank - The contamination by the analyte from all sources external
to the sample. The blank value is determined by proceeding through all
phases of extraction and analysis with no addition of sample.
263
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NBS - National Bureau of Standards.
Offshore - In this document, refers to all aspects of the receiving environment
for effluents (i.e., water, sediirent, and organisms).
Otter Board - A flat board that is attached to each side of the front end
of an otter trawl. Its planing action when being towed holds the mouth
of the net open.
Oxidation - An increase in positive valence or a decrease in negative valence
by the loss of electrons.
Penetration Depth - The maximum depth below the sediment surface that -
grab sampler achieves during a single cast.
Performance Samp". • - A sample or solution with known concentrations of
anaiytes of interest, submitted to a laboratory for the purpose of evaluating
the performance of that laboratory.
Phi Value - A measure of particle size commonly used by geologists. A
phi value is equal to the negative logarithm (base 2) of the diameter of
a particle expressed in millimeters.
Population - A generic term denoting any finite or infinite collection
of individual things, objects, or events; in the broadest concept, an aggregate
determined by some property that distinguishes things that do and do not
belong.
Precision - The degree of mutual agreement characteristic of independent
measurements as the result of repeated application of the process under
specified conditions.
Preservation - Maintenance in an unaltered form.
Preventive Maintenance - Procedures conducted routinely to ensure that
equipment continues to operate properly.
Primary Standard -A substance or artifact, the value of which can be accepted
(within specific Units) without question when used to establish the value
of the same or related property of another material.
Priority Pollutant - Those toxic pollutants defined by the U.S. EPA in
1976 that are the primary subject of regulation of the Clean Water Act.
A list of these substances can be found in the Code of Federal Regulations
Vol. 40, Section 401.15.
Purge - The removal of volatile organic compounds from the sample matrix
for analysis.
Quality - An estimation of acceptability or suitability for a given purpose
of an object, item, tangible, or intangible thing.
264
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Quality Assurance - A system of activities to provide to the producer or
user of a product or a service the assurance that it meets defined standards
of quality.
V
Quality Assessment • The overall system of activities wh-ose purpose is
to provide assurance that the quality control activities are being done
effectively. It involves a continuing evaluation of performance of the
production system and the quality of the products produced.
Quality Control - The overall system of activities whose purpose is to
control the quality of a product or service so that it meets the needs
of users. The aim is to provide quality that is satisfactory, adequate,
dependable, and economic.
Quantification - The determination or expression of the number or amount
of something.
Reagent - A solvent or other chemical used during sample p »paration or
analysis.
Recovery - The amount of an analyte detected relative to the amount added
(e.g., spike) or known to be present (e.g., standard reference material).
Usually expressed as a percentage.
Reduction - A decrease in positive valence by the loss of electrons or
an increase in negative valence by the gain of electrons.
Reference Area - A station or group of stations with which potentially
impacted stations are compared to determine the degree of impact. Ideally,
the reference area represents unaltered background conditions.
Reference Collection - A group of preserved organisms of known and verified
taxonomic identity that is used as the standard for comparison for future
taxonomic identifications.
Relative Percent Difference - Difference of two measurements xj and X2,
divided by the mean of the measurements, multiplied by 100.
Relaxation - Reduction of muscular or nervous tension.
Replicate - A counterpart of another, usually referring to an analytical
sample or a measurement. It is the general case for which duplicate is
the special case consisting of two samples or measurements.
Reproducibility - The ability to produce the same results for a measurement.
Often measured by calculation of relative percent difference or coefficient
of variation.
Resection - The surgical removal of part of an organ or structure.
Sample - A portion of a population or lot. It may consist of an individual
or groups of individuals. It may refer to objects, materials, or to measure-
ments, conceivable as part of a larger group that could have been considered.
265
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Sample Integrity - The unaltered composition of a sample.
Sample Matrix • The material in which the analytes of interest are found
(e.g., water, sediment, tissue).
Sample Tracking - Monitoring the course 0" samp1-?* through all phases of
laboratory analysis.
Scope - The length of hydrowire used when towing an otter trawl.
Secondary Standard - A standard whose value is based upon comparison with
some primary standard.
Seed - A population of microbiological organisms added to a sample for
BOD analysis because the sample does not contain a m.irobial population
sufficient for the needs of the analysis.
Sensitivity • Capability of methodology or instrumentation to discriminat
between samples having differing concentrations of an analyt* .
Significant Figure -A figure(s) that remains to a number or decimal after
the ciphers to the right or left are cancelled.
Sort - To separate benthic organisms from the inorganic and plant material
that are collected in sieved grab samples.
Spike • The addition of a known amount of an analyte or internal standard
to a sample.
Split - A replicate portion or sub-sample of a total sample obtained in
such a manner that it is not believed to differ significantly from other
portions of the sane sample.
Standard - A substance or material, the properties of which are believed
to be known with sufficient accuracy to permit its use to evaluate the
same property of another. In chemical measurements, it often describes
a solution of substance, commonly prepared by the analyst, to establish
a calibration curve or the analytical response function of an instrument.
Standard Reference Material - A material or substance one or more properties
of which are sufficiently well established to be used for the assessment
of a method or the calibration of an apparatus.
Sterilization - Reaoval of all bacteria and other microorganisms from an
object.
Subsample - A portion taken from a sample. A laboratory sample may be
a subsample of a field sample. Similarly, a test portion may be a subsample
of a laboratory sample.
Surrogate Spike Compound - A known amount of a compound, with characteristics
similar to that of an analyte, added to a sample prior to extraction. This
compound can be used to estimate recovery of analytes of interest. Also
called "recovery internal standard."
266
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Tare - The waight of a container or wrapper that is deducted from the gross
weight to obtain net weight.
Taxon - A group of organisms constituting one of the categories or formal
units in taxonomic clarification, such as phylum, class, order, family,
genus, or species.
Taxonomy - The theory, principles, and process of classifying organisms
in established categories.
TFE - A plastic tetrafluoroethylene polymer composed of very long chains
of CFg units. Commonly known by the trade name Teflon.
Titration - The process or method of determining the concentration of a
substance in solution by adding to it a standard reagent of known concentration
in carefully measured amounts until a reaction of definite and known proportion
is completed, as shown by a color change c by electrical measurement,
and then calculating the unknown concentration
Trace - Very small quantity of analyte in the sample.
True Particle Size Distribution - Distribution comprised only of inorganic
particles after organic material is oxidized completely.
Volatile Organic Compounds - Organic compounds with high vapor pressures.
In this document it refers to the 29 U.S. EPA priority pollutants considered
as volatiles (e.g., benzene).
Winnow - To separate different constituents of a substance by means of
a current of air or water.
ZID - Zone of initial dilution for an effluent discharged into the environment.
267
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APPENDIX A
-------
APPENDIX A
RECOMMENDFD METHODS FOR METALS IN EFFLUENT
-------
TABLE A-l. LIST OF TEST PROCEDURES APPROVED BY U.S. EPA
FOR INORGANIC COMPOUNDS IN EFFLUENT
Note: This table is an exact reproduction of Table IB (U.S. EPA 1984),
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A-4
-------
TABLE A-l. (Continued)
To» _______
19 *•>*»•» 01 "weon* SuMoncoo « v»«e ^d too* Soa¥nan>v- US OoBd>tmoiii n no mm US Cmou* SIM Qaon-'
gi ino>ganc SuMiancn « *•<• ond F^nvi 5ii*m»m.- NW Skeugno. at *» US Gaooocoi Sirwy T«nnaun o> v»our Aciowm
i»>*d|i9B01
•*• wiormmato* 31 101*1 «•«*« «• uneM » not wor*o Mtert grec tiara A ugoiuon orecoowo « reaped to lotaon* luoowaoa nwanoi *na 10 doiva* onto* arganc « n n i moon ottano. o* •*• IB oaiommamn* tar e*ru« aumom aucfi a* arwnc. M non* itni«ia marewy. tmjn. ana nonun I*OM* a "aonrni
**•*•*• •"• '" •• e§»" •* """^ """• «•""* M «""«**'t' «0«a*t muvenono and/or ejutena.
HfgBi n my tfg***on prooodw* tor dvla0 aapvoton or yaffln kfnaoo awnc ofioorpoon •noanvB i^dbdod 01 on* ol M ooior aoproMd rotorancoa • otftarom man 01* *DOV* *n* E0*
•« pat weugn • 0 «J moan »"»i«im« *av ^aaoMig N»iMn at in* Mno* «• r«M>«nc*a proeMu>* >v tout m«jit
or ngnnnn at u« orgr« iwnpK moan "or aw moun mor ao omnoo >or AA iar«i UOMBOX v gjiom* iurnfc.ii
Seocvomtinc M«noa tar Tree* EMrmm AnvyM o« w«i«r ana vnoun » an*« «i Aeovai C 31 im P«I
• 1ignj*t omuunon a not raggHl^ UiiioanDtn flau on iicnHnun* «mu«m aamoM* ar* on a» N*. • :•.
' (C80CX) mm not Do OOMUMO •"» m* noononoi BOO> law • It rnoaonaa tow BOO Tho oooMn ol in* nmtcaDon MWHQT .a mi
ra rocwii Oni ««» • aomoiBjr > panM iprarctfy turn CBOOk « '•»*•• em DM p*mnoo i*eon au» oourao uovio, w*
an NMml StMofdOK Mwoa^ PmwngEHkim. At* L I97S AmoM kom ANSI. ««» aVotaoof NOB van NT IOC
Bo. 2990 Coxoao StakalWl 77»40
iw p«fl*r tnduon » *• and Sworn mo
10019
. . .
»«*cidtS01 MKf. H»I*»OO» at Wtlor Anory**. 1979. MOON OwncM Comotn* . »0 Boi MS. U»«Ml»x* "»»mcidi n EPA MamoBi 33SOJ iCyanaal or «» 2 (cnonaiM or* amoXiod 6» eomKimg n* ro-umoH in* arocny 10 n*
IMB IWOT M MOMd OS B» Mm « 2 moKd Do 'ootiead •*< BH DufUr 7 1 tound a UOBIOO 33» 2
tid&eMdt MMtod. mauow umod Nurwar J7»-7SWA, Omoor 197*. Ttemcan AMO-AMTfiar H TKMMOH mauuiw Sniora
1009. 1990. MOO» CKomeal Company. PO te 399 LOMUne. CoMroao 90537
H9a Hoen MandDoo* ol woaiovaior AnaMav t—
e*ny. PO Bm 199 Lovoiana Couraao 90S37
Mama. Modiud 9030 Hocn MandDoo* ol woaiovaior AnaMav 1979 oogoa 2-n3 and 2-n7 Mien Cnomcai Comam* Lo.aiane Cooaoo 9M17
-- - — .. - - . ao (0537
Cnrornnogriony jownol ol CnromiiogriBny voi 47 NO 1 00 42<- on aovooua Ouflor ol kooMn moauitaia ana uoum "rffoioa 10 a a»
« i J nw*»oV* tar *<•• ol MM ODOM 1 nw/L »ink ol UR«M tfouM DO OM*d 10 100 im. or Movtg 40 m. aocn ol 2 M Na.SA and 2M NoOn Sianaaroa ineud DO araovoo « M
MM mavvvr Por^vooi ot a*»*r oocw i >nyL W iaconvnanoa0 "* I""*T *> infantry
i""*——* HH F—, jr om limntfi r •Woar TomuBiMo innygnail Ticion. rigid Momtonmn ana Oiu P>»o«miBDn. U S Gadoacal Sun»» T*ennojya 01 *aioi «•
' i i Ooptor 01 1971
zJw kMOaoMMM 9009 HO* Hondboo* ol W*» Anaom. 1979 eogo* 2-231 and 2-333 MOCA OwrmcM Comoony toMUna Ceioraao 90S37
. «--— , ^^ Q^, ^ „„ yn^d Juit, tiMimMBonji Proionon Agoner.' tKpiaitiani H n* faaanai Eanon ol SiMfO*n minoea *r «, f.
a« NMor an* WKMOBBUV 1491 Eoajon. Tha uaumninc '**ciari • tonnxiad n a an 01 '00 : 02 >o
110A lor a^anaon. Uootod SlOi tor 91* rnonu* uonimiainc groeoov* or uwnao HOC tor in*
97-70 1977.
A-5
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APPENDIX B
RECOMMENDED METHODS FOR ORGANIC COMPOUNDS IN EFFLUENT
-------
TABLE B-l. LIST OF TEST PROCEDURES APPROVED BY U.S. EPA
FOR NON-PESTICIDE ORGANIC COMPOUNDS IN EFFLUENT
Note: This table is an exact reproduction of Table 1C (U.S. EPA 1984),
IP* i
QC
00 US
OBUT
14,
il
it
17
11
IfL
a
t*
XL «
31 «
Ml.
•01.
M
M.
•14. 104
•14. 1U4
•14. I«M
U4.1U4
B-l
-------
TABLE 3-1. (Continued)
•M ««f M
807 M ••*. or
-------
TABLE B-2. LIST OF TEST PROCEDURES APPROVED BY U.S. EPA
FOR PESTICIDES IN EFFLUENT
Note: This table is an exact reproduction of Table ID (U.S. EPA 1984)
ASTM
(BOM I Now 3. 0 7. NOW 4. 0. X
NOWl. aU.NoWt0.SM.
NOWl 0.M NOWC.0 SU
NOW! 0 U.NOW* 0 SM
NOW! 0 13. NOW* 0 SM
Now 1. 0 IS. Now • p SSI
NOW 3. 0 10V NOW • 0 SM
NOWl 0 7
M NOWl 0 SM
X NOW • 0 S7J
7
104 NOW • 0 5*4
M» NOW 4 0 3S
7 NOW 4 0 30
7 NOW 4 0 X
7
IS NOW 6 0 SSI
« NOW I 0 SSI
M NOW 4 0 X NOW t 3
us
X. NOW • 0 S71
7
7 NOW* 0 X
» NOW • P S71
NOW* P SSI
104 NOW • 0 SM
7
7 NOW 4 p X
Now«0X:Naw«.0S73
NOW3.0 104, NOW 10 SM
NOW 1. 0 10* NOW •. p SM
NOW 3 0. 7 NOW 4 p X
NOW 1. 0. 7 NOW 4 0. X NOW 6 P
sn.
B-3
-------
TABLE B-2. (Continued)
»«yu
ASTM
u
MM
MOW 4 e X NOW • 0 SM
Mow 3 0 104 NOW « o W
MOW3P2SNoW4BXNeUll
NOW 3 0 04 NOM • V SM
NOW I. e i NOW 4 » x
'NOW! 0 §4 NOW* 0 SM
'MOW3 0 7
i MOW 3. 0 104 Now • 0 SS4
i NOW 3 0 104 NOW I 0 S44
I NOW 3 0 104 NOW 6 0 S*4
INOW3 0.M NOW 4 e X
' Noli 3
U. PtMnim
M. r i» i •iar
• «• •kBMMBIMVkBIO]
Si. P nciim ,
U ftuowrp __— _^^____
• 1 fin «••!.
M. tim»w
fl»*HP
M, H.S.T
£ f^Urr^11
fiC
oc
ee
TIC
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Tie
ae
ae
ne
ae
ae
ae
ae.
ae
M
NOB
— H
MM
MM
i
j Tjua*
- -
OHM
NOW 10 t
NOW 3 J U NOWI 0 SM
NOW 3. 0. U. Now • 0 SM
Now 1 0 U Now •. 0 SM
MM 1 0 104. NOW •. 0 SM
NOW 1 0 M. New • 0 SM
MOW10U. NOW(»1M
M0H 1 0. 104 NOW C 0 M4
MOM Ipo. Mowt. p SM
New 3.0 r
NOM 1 0 104 MOW • 0 St4
Maw 1 0. MI Mew 4. p Jt
MOW 10. Ill
Mew 10 O- MOW* p SM
MOW 10. t M0M4 p 30
MOM 3. p. r
•PMadM on KM « Hi MM oteaim
•Th» M w«t o» iiiirum eot «na to or*
nwr bf taw*
TIBW c.
! ot BW reiaor laamioi POM _
> I* AOOYM el Oojnc POM***." o( *•> *M 131. Tho i
ill 'pponiti •. "Oitinui 0na ProoMvoier w» Oounvwaan et *w
Uxnaa DOWCBBI utw 91
H'l
M n««tod oiioi.«un tni IUOU *ar two* wo)
Seek S. OMBWI A) iiiru
an • rw er«M»M nwa«a
ana tmum 10% • «§ wnowo vwrtM «w
B-4
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