United States
Environmental Protection
Agency
Off ice of Water
(4305)
EPA-823-B-01-002
October 2001
Methods for Collection, Storage and
Manipulation of Sediments for Chemical
and Toxicological Analyses:
Technical Manual
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EPA-823-B-01-002
October 2001
Methods for Collection, Storage and
Manipulation of Sediments for Chemical and
Toxicological Analyses: Technical Manual
Office of Science & Technology
Office of Water
U.S. Environmental Protection Agency
Washington, DC 20460
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Disclaimer
This technical manual provides a compilation of current information and recommendations for collecting,
handling and manipulating sediment samples for physicochemical characterization and biological testing
that are most likely to yield accurate, representative sediment quality data based on the experience of
many monitoring programs and researchers. This manual has no immediate or direct regulatory
consequence. It does not impose legally binding requirements on EPA, States, Tribes, other regulatory
authorities, or the regulated community, and may not apply to a particular situation based upon the
circumstances. EPA, State, Tribal, and other decision makers retain the discretion to adopt approaches
on a case-by-case basis that differ from those in this manual where appropriate. EPA may update this
manual in the future as better information becomes available.
This document has been approved for publication by the Office of Science and Technology, Office of
Water, U.S. Environmental Protection Agency. Mention of trade names, products, or services does not
convey and should not be interpreted as conveying, official USEPA approval, endorsement, or
recommendation for use.
The suggested citation for this document is:
U.S. EPA. 2001. Methods for Collection, Storage and Manipulation of Sediments for Chemical and
Toxicological Analyses: Technical Manual. EPA 823-B-01-002. U.S. Environmental Protection
Agency, Office of Water, Washington, DC.
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Technical Manual
Acknowledgments
This document is a general purpose manual intended to provide the user with sediment collection,
storage, and manipulation methods that are most likely to yield accurate, representative sediment
quality data for toxicity and chemical anlayses based on the experience of many monitoring programs
and researchers. The approaches described in this manual represents a compilation of information
presented in many publications, including Puget Sound Estuary Program (PSEP, 1997), Washington
State Department of Ecology (1995), Environment Canada (1994), US Environmental Protection
Agency - US Army Corps of Engineers (USEPA-USACE, 1998), American Society for Testing and
Materials (ASTM, 2000), and USEPA (2000).
The principal authors of this manual are Kathy Zirbser, Richard Healy, Leanne Stahl, Bill Tate
(USEPA, Office of Science and Technology), Jerry Diamond (Tetra Tech, Inc.), Allen Burton
(Wright State University), Michael Johns (Windward Environmental LLC), and John Scott (SAIC).
Review comments from the following individuals led to substantial improvements in the manual for
which we are grateful:
Tom Armitage
Justine Barton
Brett Belts
Kathryn Bragdon-Cook
James Brannon
Robert Burgess
Scott Can-
Scott Cieniawski
Philip Crocker
Alan Crockett
Kathleen Dadey
Robert Engler
Ken Finkelstein
Maria Gomez-Taylor
Tom Gries
Erika Hoffman
Chris Ingersoll
Laura Johnson
Ash Jain
Peter Landrum
Sharon Lin
Ed Long
Gui Lotufo
Don MacDonald
John Malek
USEPA- OST
USEPA - Region 10
WA Dept of Ecology: Sediment Management Unit
Toxics Cleanup Program: Sediment Management Unit
Army Corps of Engineers - WES
USEPA - NHEERL Atlantic Ecology Division
USGS - Marine Ecotoxicology Research Station
USEPA - Region 5
USEPA - Region 6
Consultant
USEPA - Region 9
Army Corps of Engineers - WES
USEPA - Region 1
USEPA - BAD
WA Dept of Ecology
USEPA - Region 10 Sediment Management Program
USGS - Columbia Environmental Research Center
USEPA - OCPD
EPRI
NOAA - Great Lakes Env. Research Laboratory
USEPA - OCPD/WD
NOAA
Army Corps of Engineers - WES
NOAA - Office of Response and Restoration
USEPA - Region 10 Sediment Management Program
Acknowledgments
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Cornell Rosiu USEPA - Region 1
Brian Ross USEPA - Region 9 Dredging and Sed. Management
Timothy Sherman Army Corps of Engineers - Portland District
Robert Shippen USEPA- OST
Mark Siipola Army Corps of Engineers - Portland District
Jerry Smrchek USEPA - OPPT/OPPTS
Mark Sprenger USEPA - OERR-ERTC
Marc Tuchman USEPA - Region 5
Ernest Waterman USEPA - Region 1
KathyZirbser USEPA- OST
We are very grateful to Sherwin Beck (Tetra Tech, Inc.) as well as contributions from Carmela
Biddle, Marcus Bowersox, Brenda Fowler, Abby Markowitz, Patricia McCreesh, Kristen Pavlik
(Tetra Tech, Inc.), and Corinne Marino (VJB Associates) to this manual.
Front cover photographs provided by Allen Burton.
iv US Environmental Protection Agency
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Technical Manual
Acronym List
ACOE
ARCS
ASTM
AVS
BMPs
BOD
CEC
COD
CV
DOC
DQO
EDMI
EMAP
ERM
GC/MS
GC/FID
GC/ECD
GLNPO
GPC
GPS
HPLC
ICP-AES
ICP-MS
IR
LORAN
NAWQA
Army Corps of Engineers
Assessment and Remediation of Contaminated Sediments
American Society for Testing and Materials
Acid Volatile Sulfides
Best Management Practices
Biochemical Oxygen Demand
Cation Exchange Capacity
Chemical Oxygen Demand
Coefficient of Variation
Dissolved Organic Carbon
Data Quality Objectives
Electronic Distance Measurement Instrument
Environmental Monitoring & Assessment Program
Effect Range Medium
Gas Chromatography/Mass Spectrophotometry
Gas Chromatography/Flame lonization Detection
Gas Chromatography/Electron Capture Detection
Great Lakes National Program Office
Gel Permeation Chromatography
Global Positioning System
High Performance Liquid Chromatography
Inductively Coupled Plasma Atomic Emission Spectoscopy
Inductively Coupled Plasma Mass Spectrometry
Infrared Spectrophotometer
LOng RAnge Navigation
National Water Quality Assessment
Acronym List
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
NDIR
NEP
NOAA
NSI
NST
ORP
OSHA
PCE
POC
PSEP
QA
QAPP
QC
RADAR
ROV
RPD
SATNAV
SCV
SEM
SIM
SOC
SOD
SOPs
SPMD
SRM
TIC
TMDLs
TOC
TPH
TVS
Non-Dispersive Infrared Detector
National Estuary Program
National Oceanic and Atmospheric Administration
National Sediment Inventory
National Status & Trends
Oxidation Reduction Potential
Occupational Safety & Health Administration
Power Cost Efficiency
Particulate Organic Carbon
Puget Sound Estuary Program
Quality Assurance
Quality Assurance Project Plan
Quality Control
RAdio Detecting and Ranging
Remotely Operated Vehicle
Relative Percent Difference
SATellite NAVigation
Secondary Chronic Value
Simultaneously Extracted Metals
Selected Ion Monitoring
Suspended Organic Carbon
Sediment Oxygen Demand
Standard Operating Procedures
Semi-Permeable Membrane Device
Standard Reference Materials
Total Inorganic Carbon
Total Maximum Daily Loads
Total Organic Carbon
Total Petroleum Hydrocarbons
Total Volatile Solids
VI
US Environmental Protection Agency
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Technical Manual
USEPA United States Environmental Protection Agency
USGS United States Geologic Survey
XRF X-Ray Fluorescence
Acronym List vii
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viii US Environmental Protection Agency
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Technical Manual
Foreword
Sediments provide essential habitat for many freshwater, estuarine, and marine organisms. In aquatic
systems, most anthropogenic chemicals and waste materials, particularly persistent organic and
inorganic chemicals, may accumulate in sediments. These sediments become repositories for many
of the more toxic chemicals that are introduced into surface waters. United States Environmental
Protection Agency's National Sediment Inventory (NSI) (USEPA 1998), a biennial report to
Congress on sediment quality in the United States, demonstrates that sediment contamination exists
in every state of the country. Contaminated sediments represent a hazard to aquatic life through
direct toxicity as well as to aquatic life, wildlife and human health through bioaccumulation in the
food chain. Assessments of sediment quality commonly include analyses of anthropogenic
contaminants, benthic community structure, physicochemical characteristics, and direct measures of
whole sediment and pore water toxicity. Accurate assessment of environmental hazards posed by
sediment contamination depends in large part on the accuracy and representativeness of these
analyses.
The methods described in this Manual are intended to provide the user with sediment collection,
storage, and manipulation methods that are most likely to yield accurate, representative sediment
quality data (e.g., toxicity, chemical) based on the experience of many monitoring programs and
researchers.
This Manual represents a compilation of information presented in many publications, including:
• American Society for Testing and Materials (ASTM) 2000 document: Standard Guide for
Storage, Characterization, and Manipulation of Sediments for Toxicological Testing, E-
1391-94.
• Environment Canada 1994 manual: Guidance Document on Collection and Preparation of
Sediments for Physicochemical Characterization and Biological Testing, EPS l/RM/29.
• U.S. Environmental Protection Agency. 2000 manual: Methods for Measuring the Toxicity
and Bioaccumulation of Sediment-Associated Contaminants with Freshwater Invertebrates.
Second Edition. EPA/600/R-99/064.
• U.S. Environmental Protection Agency/Army Corps of Engineers. 1998. Inland Testing
Manual: Evaluation of Dredged Material Proposed for Discharge in Waters of the U.S. -
Testing Manual. EPA-823-B-98-004.
• U.S. Environmental Protection Agency / Army Corps of Engineers. 1991. Ocean Testing
Manual: Evaluation of Dredged Material Proposed for Ocean Disposal: Testing Manual.
EPA-503/8-91/001.
In addition to many recent peer-reviewed technical journal papers, other publications that were relied
on extensively include:
Foreword ix
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
• Puget Sound Estuary Program (PSEP) 1997 manual: Recommended Guidelines for Sampling
Marine Sediment, Water Column, and Tissue in Puget Sound
• Washington Department of Ecology 1995 Document: Guidance on the Development of
Sediment Sampling and Analysis Plans Meeting the Requirements of the Sediment
Management Standards
• Great Lakes National Program Office (GLNPO) 1994 manual: Assessment and Remediation
of Contaminated sediments (ARCS) Program - Assessment Guidance EPA-905-B94-002.
• U.S. Environmental Protection Agency. 2000 document: Estuarine and Near Coastal Marine
Waters: Bioassessment and Biocriteria Technical Guidance. EPA-822-B-00-004.
This Manual addresses several needs identified in EPA's Contaminated Sediment Strategy
(USEPA 1998) including: (1) an organized discussion of activities involved in sediment sampling
and sample processing; (2) important issues that need to be considered within each activity; and
(3) recommendations on how to best address issues such as sampling design, proper sampling
procedures, and sample manipulations. Throughout this Manual, different considerations pertaining
to sampling and sample processing are presented depending on the program need (e.g., dredge
remediation versus status and trends monitoring).
EPA along with other agencies, assesses aquatic sediment quality under a variety of legislative
requirements including:
• National Environmental Policy Act (NEPA)
• Clean Air Act; the Coastal Zone Management Act (CZMA)
• Federal Insecticide, Fungicide, and Rodenticide Act (FIFRA)
• Marine Protection, Research, and Sanctuaries Act (MPRSA)
• Resource Conservation and Recovery Act (RCRA)
• Toxic Substance Control Act (TSCA)
Clean Water Act (CWA)
• Comprehensive, Environmental and Liability Act (CERCLA)
• Great Lakes Critical Programs Act of 1990.
In addition, many EPA offices coordinate sediment monitoring studies in specific geographic areas,
such as through the Chesapeake Bay Program, the Great Lakes National Program, the Gulf of Mexico
Program, the Washington State Sediment Management Standards Program, and in the States of
Washington, Florida, California, New York, New Jersey, South Carolina, Texas, Massachusetts, and
Wisconsin. To address its responsibilities within the above legislative acts, EPA has several ongoing
programs that may involve sediment quality evaluation as summarized below.
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Dredged Material Management
The U.S. Army Corps of Engineers (USAGE), the Federal agency designated to maintain navigable
waters, conducts a majority of the dredging projects and disposal under its Congressionally-
authorized civil works program. The balance of dredging and disposal is conducted by a number of
local public and private entities. In either case, the disposal is subjected to a regulatory program
administered jointly by the USAGE and EPA under Section 103 of the Marine Protection, Research,
and Sanctuaries Act (MPRSA) for ocean disposal, and Section 404 of the Clean Water Act (CWA)
for discharge at open water sites, confined disposal facilities with return flow to waters of the U.S.,
or for beneficial uses. EPA shares the responsibility of managing dredged material, principally in the
development of the environmental criteria and guidelines by which proposed discharges are
evaluated and disposal sites are selected, and in the exercise of its environmental oversight authority.
Joint EPA/USACE guidance manuals detailing the testing and analysis protocols for dredged
material disposal are well established.
National Estuary Program
EPA administers the National Estuary Program, established under the Clean Water Act to identify,
restore, and protect nationally significant estuaries in the United States. Within the existing 28
programs, environmental monitoring is a key element of watershed protection strategies developed to
maintain the chemical, physical, and biological properties of the estuarine ecosystems. The Puget
Sound Estuary Program (PSEP), in particular, has been actively monitoring ecological health,
including sediment quality, in Puget Sound, Washington for many years. PSEP, which includes
EPA, the Puget Sound Water Quality Authority, and the Washington Department of Ecology, has
developed sediment sampling and analysis procedures in collaboration with local governments and
stakeholder groups (PSEP, 1997). The protocols are cited in and support the Washington
Department of Ecology's (WDE) sediment management standards regulation, and have served as the
foundation for many other guidance documents such as those produced by Environment Canada
(1994) and American Society of Testing Materials (ASTM, 2000). This manual frequently refers to
PSEP and WDE guidance.
Resource Conservation and Recovery Act (RCRA)
Under RCRA, EPA assesses whether releases from a hazardous waste treatment, storage, or disposal
facility have contaminated sediments and requires corrective action, including possible remediation,
if contamination is discovered. In many cases, sediment sampling and analyses, as discussed in this
manual, are needed in RCRA facility assessments and RCRA facility investigations.
Office of Water
The Office of Water has been expanding provisions for sediment monitoring under the Clean Water
Act, in the national monitoring framework developed by the Intergovernmental Task Force on
Monitoring Water Quality (ITEM, 1995). Through this framework, agreements have been reached
with other Federal, State, and local agencies concerning incorporation of sediment monitoring
protocols, sediment monitoring QA/QC procedures, and appropriate information system linkages into
monitoring programs. The Office of Water and the Office of Information Resources Management are
also ensuring that the capability to store and use sediment data is enhanced as part of the ongoing
modernization of the Agency's water quality data systems (STORET), and in coordination with the
water quality data elements procedures being recommended by the National Methods and Data
Comparability Board under the National Water Quality Monitoring Council. These data elements
include information describing how samples were collected, stored, and processed prior to analysis.
Foreword xi
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Regional Environmental Monitoring and Assessment Program (REMAP)
REMAP, within the Office of Research and Development, gathers chemical and biological data
describing sediment quality at many EMAP sampling stations. Data collected under REMAP are
entered into the National Sediment Inventory (NSI). These data are used to assess status and trends
on a regional scale, particularly for aquatic systems that may have water quality and/or sediment
quality impairment.
Comprehensive, Environmental and Liability Act (CERCLA)
Under CERCLA, EPA carries out a detailed analysis at a site, evaluating the risks posed by
contaminants to human health and the environment, and the feasibility of various response action
alternatives to reduce risk. The Risk Assessment Guidance for Superfund (USEPA, 1997) provides a
framework for the assessment of human health and environmental impacts. The CERCLA Program
is using the EPA-wide sediment testing methods of the Tiered Testing Framework in the Remedial
Investigation/Feasibility Study (CRI/FS) stage of analysis to help determine options for remedial
actions. Much of the guidance presented in this manual supports the Tiered Testing Framework
applicable to CERCLA sites.
Great Lakes Critical Programs Act of 1990
Annex 14 of the Great Lakes Water Quality Agreement between the United States and Canada (as
amended by the 1987 Protocol) stipulates that the cooperating parties will identify the nature and
extent of sediment contamination in the Great Lakes, develop methods to assess impacts, and
evaluate the technological capability of programs to remedy such contamination. The 1987
amendments to the Clean Water Act authorized the Great Lakes National Program Office (GLNPO)
to coordinate and conduct studies and demonstration projects relating to the appropriate treatment of
toxic contaminants in bottom sediments. To fulfill the requirements of the Act, GLNPO initiated the
Assessment and Remediation of Contaminated Sediments (ARCS) Program to help address
contaminated sediment concerns in the development of Remedial Action Plans (RAPs) for all 43
Great Lakes Areas of Concern (AOCs, as identified by the United States and Canadian governments),
as well as similar concerns in the development of Lakewide Management Plans. This manual
frequently relies on information documented by the GLNPO and the ARCS program.
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TABLE OF CONTENTS
Page
Acknowledgments iii
Acronym List v
Foreword ix
Table of Contents xiii
List of Tables xvii
List of Figures xvii
List of Recommendations xix
Technical and Grammatical Terms xxi
Using the Manual xxiv
1. INTRODUCTION 1-1
1.1 Background 1-1
1.2 Significance and Use of this Manual 1-1
1.3 Applicability and Scope of this Manual 1-2
2. SEDIMENT MONITORING AND ASSESSMENT STUDY PLANS 2-1
2.1 Data Quality Objectives Process 2-1
2.2 Study Plan Considerations 2-5
2.2.1 Definition of the Study Area and Study Site 2-5
2.2.2 Controlling Sources of Variability 2-5
2.2.3 Sampling Using an Index Period 2-6
2.3 Sampling Designs 2-7
2.3.1 Probabilistic and Random Sampling 2-7
2.3.2 Targeted Sampling Designs 2-10
2.4 Measurement Quality Objectives 2-11
2.4.1 Sample Volume 2-12
2.4.2 Number of Samples 2-15
2.4.3 Replicate and Composite Samples 2-15
2.5 Site-Specific Selection Considerations for Selecting Sediment
Sampling Stations 2-17
2.5.1 Review Available Data 2-19
2.5.2 Site Inspection 2-19
2.5.3 Identify Sediment Deposition and Erosional Zones 2-19
2.6 Positioning Methods for Locating Sampling Stations 2-20
2.7 Preparations for Field Sampling 2-21
2.8 Health and Safety 2-23
3. COLLECTION OF WHOLE SEDIMENTS 3-1
3.1 General Procedures 3-1
3.2 Types of Sediment Samplers 3-5
3.2.1 Grab Samplers 3-5
3.2.2 Core Samplers 3-9
3.3 Sample Acceptability 3-13
3.4 Equipment Decontamination 3-14
3.5 Field Measurements and Observations 3-15
3.6 Documentation of Sample Collection 3-17
Table of Contents xiii
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TABLE OF CONTENTS (CONTINUED)
Page
4. FIELD SAMPLE PROCESSING, TRANSPORT, AND STORAGE OF
SEDIMENTS 4-1
4.1 Sample Containers 4-1
4.1.1 Container Material 4-3
4.1.2 Container Preparation 4-3
4.2 Subsampling and Compositing Samples 4-5
4.2.1 General Procedures 4-5
4.2.2 Grab Samples 4-6
4.2.3 Core Samples 4-7
4.3 Homogenization 4-11
4.3.1 General Procedures 4-11
4.4 Sample Transport and Storage 4-13
4.4.1 General Procedures 4-13
4.5 Sample Holding Times 4-15
5. SEDIMENT MANIPULATIONS 5-1
5.1 Sieving 5-1
5.1.1 Sieving Methods 5-3
5.1.2 Alternatives to Sieving 5-5
5.2 Formulated Sediment and Organic Carbon Modification 5-6
5.2.1 General Considerations 5-6
5.2.2 Sediment Sources 5-7
5.2.3 Organic Carbon Modification 5-7
5.3 Spiking 5-8
5.3.1 Preparation for Spiking 5-9
5.3.2 Methods for Spiking 5-10
5.3.3 Equilibration Times 5-12
5.3.4 Use of Organic Solvents 5-13
5.4 Preparation of Sediment Dilutions 5-13
5.5 Preparation of Sediment Elutriates 5-14
6. COLLECTION OF INTERSTITIAL WATER 6-1
6.1 General Procedures 6-1
6.2 In-situ Collection 6-2
6.2.1 Peeper Methods 6-5
6.2.2 Suction Methods 6-7
6.2.3 Processing of Field-Collected Interstitial Water Sample 6-8
6.3 Ex-situ Extraction of Interstitial Water 6-8
6.3.1 General Procedures 6-8
6.3.2 Centrifugation 6-10
6.3.3 Sediment Squeezing 6-12
6.3.4 Pressurized and Vacuum Devices 6-12
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TABLE OF CONTENTS (CONTINUED)
Page
7. QUALITY ASSURANCE AND QUALITY CONTROL 7-1
7.1 General Procedures 7-1
7.2 QA/QC Procedures for Sediment Collection and Manipulation 7-2
7.3 The Quality Assurance Project Plan (QAPP) 7-2
7.4 Standard Operating Procedures 7-3
7.5 Sediment Sample Documentation 7-3
7.6 Sample Tracking Documentation 7-4
7.7 Record Keeping 7-5
7.8 QA Audits 7-5
7.9 Corrective Action (Management of Non-conformance Events) 7-5
7.10 Data Reporting 7-6
8. REFERENCES 8-1
APPENDICES
A EXAMPLES OF SEDIMENT QUALITY SAMPLING DESIGNS USING THE DATA
QUALITY OBJECTIVES (DQO) PROCESS
B EXAMPLES OF MEASUREMENT QUALITY OBJECTIVES USED
IN SEDIMENT QUALITY MONITORING STUDIES
C STATISTICAL CONSIDERATIONS IN DETERMINING THE APPROPRIATE
NUMBER OF REPLICATE SAMPLES NEEDED AT EACH SAMPLING STATION
D ADVANTAGES AND DISADVANTAGES OF DIFFERENT STATION
POSITIONING TECHNIQUES
E ADVANTAGES, DISADVANTAGES AND ILLUSTRATIONS OF GRAB AND CORE
SAMPLING DEVICES USED IN SEDIMENT MONITORING STUDIES
F EXAMPLES OF FIELD FORMS USED TO DOCUMENT STATION AND SAMPLE
CHARACTERISTICS AND SAMPLE TRACKING
G PHYSICO-CHEMICAL SEDIMENT CHARACTERIZATION
Table of Contents xv
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LIST OF TABLES
Table Page
2-1 Suggestions for selecting an appropriate sampling design (from USEPA 2000b) 2-9
2-2 Conventional sediment variables and their use in sediment investigations (Adapted
from WDE, 1995) 2-13
2-3 Typical sediment volume requirements for various analyses per sample 2-14
2-4 Practical considerations for site-specific selection of sampling stations in developing
a sampling plan 2-18
4-1 Recommended sampling containers, holding times, and storage conditions for
common types of sediment analyses 4-4
6-1 In Situ interstitial water collection methods 6-5
LIST OF FIGURES
Figure Page
1-1 Flow chart summarizing activities for collection, storage, and manipulation of
sediments and interstitial water 1-3
2-1 Flow chart summarizing the process that should be implemented in designing
and performing a monitoring study (modified from MacDonald et al. (1991) 2-2
2-2 Flow chart summarizing the Data Quality Objectives Process (after USEPA
2000a) 2-3
2-3 Description of various sampling methods (adapted from USEPA 2000c) 2-8
3-1 General types of considerations or objectives that are appropriate for grab or core
sampling devices 3-1
3-2 Flowchart for selecting appropriate grab samplers based on site-specific or design
factors 3-2
3-3 Flowchart for selecting appropriate core samplers based on site-specific factors 3-3
3-4 Illustrations of acceptable and unacceptable grab samples 3-15
4-1 Flowchart of suggested sediment processing procedures 4-2
4-2 Alternatives for subsampling and compositing sediment grab samples 4-8
4-3 Alternatives for subsampling and compositing sediment core samples 4-9
Table of Contents xvii
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LIST OF FIGURES (CONTINUED)
Figure Page
5-1 Flowchart depicting relationships between common sediment manipulations
including important considerations 5-2
6-1 Considerations for selecting the appropriate type of interstitial water sampling
method 6-3
6-2 Front view and components of peeper sampling devices (top: plate device; bottom:
cylindrical probe) 6-6
6-3 Summary of recommended procedures and considerations for laboratory isolation
of interstitial water 6-9
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LIST OF RECOMMENDATION BOXES
Chapter 2 Page
Box #1 What type of sampling strategy should be used 2-10
Box #2 How many samples and how much sample volume should be collected 2-14
Box #3 How should station positioning be performed 2-20
Box #4 What health and safety precautions should be followed 2-24
Chapter 3
Box #1 What are appropriate sampling devices given different study
objectives 3-9
Box #2 How should sampling devices be used 3-13
Box #3 What information should be documented for each sample collected 3-16
Chapter 4
Box #1 Sample containers 4-1
Box #2 How should sediment samples be subsampled and composited 4-5
Box #3 How should samples be homogenized 4-11
Box #4 Sample transport and storage 4-14
Box #5 How long should samples be stored before analysis 4-15
Chapter 5
Box #1 Should sediment be sieved prior to analyses 5-3
Box #2 What type of sieve should be used 5-5
Box #3 How should sediments be spiked with a chemical or other test material 5-9
Box #4 How should sediment elutriates be performed 5-15
Chapter 6
Box #1 In-situ interstitial water collection 6-2
Box #2 Extraction of interstitial water 6-10
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Technical Terms
The following definitions were derived primarily from ASTM, USEPA, ACOE, and Environment
Canada sources.
Acid Volatile Sulfide. The sulfides removed from sediment by cold acid extraction, consisting
mainly of iron sulfide. AVS is the principal binding phase in sediment for divalent metals.
Artifact. An undesirable, detectable feature (e.g., chemical or physical change) in a sample, that has
resulted from sampling, sample handling or storage, or from manipulations of the sample.
Benthic. Associated with the bottom of a waterbody.
Bioaccumulation. The net accumulation of a substance by an organism as a result of uptake from all
environmental sources.
Bioavailability. The degree to which a chemical is taken up by aquatic organisms.
Chain-of-custody. The documentation that establishes the control of a sample between the time it is
collected and the time it is analyzed. It usually applies to legal samples to demonstrate that there was
no tampering with, or contamination of, the sample during this time.
Clean. Denotes a sediment or water test sample determined to not contain concentrations of
contaminants which cause apparent and unacceptable harm (or effects) to the test organisms.
Composite sample. A sample that is formed by combining material from more than one sample or
subsample.
Concentration. The ratio of weight or volume of test material(s) to the weight or volume of
sediment or water.
Contaminated sediment. Sediment containing chemical substances at concentrations that pose a
known or suspected threat to environmental or human health.
Control sediment. A sediment that is essentially free of contaminants and is used routinely to assess
the acceptability of a test. Any contaminants in control sediment may originate from the global
spread of pollutants and do not reflect any substantial input from local or non-point sources.
Comparing test sediments to control sediments is a measure of the toxicity of a test sediment beyond
inevitable background contamination.
Core sample. A sediment sample collected to obtain a vertical profile using a variety of instruments.
Data Quality Objectives (DQOs). Qualitative and quantitative statements that clarify the purpose
of the monitoring study, define the most appropriate type of data to collect, and determine the most
appropriate methods and conditions under which to collect them.
Decontamination. A process of washing or rinsing that removes chemicals adhering to equipment
and supplies.
Terms xxi
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Ecotox Thresholds (ET). Benchmark values in ecological risk assessments defined as media-
specific contaminant concentrations above which there is sufficient concern regarding adverse
ecological effects to warrant further site investigation.
Elutriate. An aqueous solution obtained after adding water to a solid substance or loose material
(e.g., sediment, tailings, drilling mud, dredge spoil), shaking the mixture, then centrifuging or
filtering it or decanting the supernatant.
Equilibration. The condition in which a material or contaminant is at steady state between the solid
or particulate sediment and the interstitial water.
Formulated Sediment. Mixtures of materials used to mimic a natural sediment.
Global Positioning system (GPS). A navigation system that relies on satellite information. It can
give continuous position reports(i.e., latitude and longitude) that vary in accuracy depending on the
sophistication of the receiving unit.
Grab. Any device designed to "bite" or "scoop" into the bottom sediment of a lake, stream, estuary,
ocean, and similar habitats to sample the benthos. Grabs are samplers with jaws that are forced shut
by weights, lever arms, springs or cables. Scoops are grab samplers that scoop sediment with a
rotating container.
Head Space. The space in the storage container between the top of the sample and the lid of the
container.
Holding time. The period of time during which a sediment or water sample can be stored after
collection, and before analysis or use in a biological test. Changes that occur in sediments or water
should be minimal during this period and the integrity of the sample should not be compromised to
any substantial degree with respect to its physical, chemical, or biological characteristics.
Homogenization. The complete mixing of sediment, either by hand or mechanical means, until
physical, chemical, and /or biological homogeneity of the sample is achieved.
Index Period. Specific time period in which sampling or in-situ analyses are conducted. Generally
pertains to an ecologically important season and/or desired environmental conditions under which
sampling is performed.
In Situ. Refers to the original (field) location from which test samples are collected, or at which
organisms are exposed to undisturbed water or sediments for extended periods.
Interferences. Characteristics of sediments or sediment test systems that can potentially affect
analytical results or test organism response aside from responses related to sediment contamination.
Types of interferences include: non-contaminant characteristics (e.g., sediment texture or grain size,
lighting); changes in chemical bioavailability due to sample handling or storage (e.g., ammonia
generation); and the presence of indigenous organisms. Also referred to as confounding factors.
Interstitial water. Water occupying space between sediment or soil particles.
Measurement Quality Objectives (MQOs). Statements that describe the amount, type, and quality
of data needed to address the overall project objectives.
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Overlying water. The water placed over sediment in a test chamber during a test.
Peepers. Devices that collect interstitial water by diffusion through membranes attached to
collection chambers. The chambers are typically placed in the sediment for extended periods of time
to allow for equilibration between the internal water environment of the peeper and the surrounding
ambient sediment/interstitial water matrix.
Pore water. See interstitial water.
Quality Assurance Project Plan. Project-specific document that specifies the data quality and
quantity requirements needed for the study as well as all procedures that will be used to collect,
analyze, and report those data.
Reference sediment. A whole sediment, collected near an area of concern, that is used as a point of
comparison to assess sediment conditions exclusive of the material(s) or activities of interest. The
reference sediment may be used as an indicator of localized sediment conditions exclusive of the
specific pollutant input of concern. Such sediment would be collected near the site of concern and
would represent the background conditions resulting from any localized pollutant inputs as well as
global pollutant input. Program-specific guidance documents should be consulted, as some EPA
programs have specific definitions and requirements for reference sediment.
Sampling Platform A working space, such as the deck of a boat, from which all sample collection
activities are conducted.
Sediment. Particulate material that usually lies below water, or formulated particulate material that
is intended to lie below water in a test.
Sediment Quality Triad. A weight-of-evidence sediment quality assessment approach which
integrates data from sediment toxicity tests, chemical analyses, and benthic community assessments.
Sieving. Selectively removing certain size fractions of the sediment sample by processing sediment
through selected mesh sizes.
Site. A study area that can be comprised of multiple sampling stations.
Spiking. Addition of a known amount of test material to a sediment often used as a quality control
check for bias due to interference or matrix effects.
Station. A sampling location within a study area or site, where physical, chemical, or biological
sampling and/or testing occurs.
Supernatant. The water separated from a sediment/water mixture following centrifugation or other
separation techniques.
Toxicity. The property of a chemical, or combination of chemicals, to adversely affect organisms,
tissues or cells.
Whole sediment. Sediment and associated interstitial water which have had minimal manipulation.
Also referred to as bulk sediment.
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Grammatical Terms
Consistent with guidance formulated by the American Society for Testing and Materials (ASTM),
the following grammatical phrases, used in this manual, are defined as follows:
The words "must", "should", "may", "can", and "might" have specific meanings in this manual.
"Must" is used to express an absolute requirement, that is, to produce accurate results, a sample
ought to be handled or manipulated in a specified manner, unless the purpose of the study requires a
different procedure.
"Should" is used to state that the specified condition or procedure is recommended and ought to be
met if possible. Although violation of one "should" is rarely a serious matter, violations of several
will often render the results questionable.
"Desirable" is used in connection with less important factors.
"May" is used to mean "is allowed to." "Can" is used to mean "is able to." "Might" is used to mean
"could possibly." Thus, the classic distinction between "may" and "can" is preserved, and "might" is
not used as a synonym for either "may" or "can."
Using the Manual
Throughout this Manual, there are three categories of information that are organized into text boxes
as part of the effort to make this methods document more useful and accessible to users. Each box
always appears with the same icon throughout the Manual:
Recommendations for procedures and equipment.
Consideration, or issues, that should be addressed
Checklists of information
The full list of Recommendation Boxes are identified on page xix as part of the Table of Contents.
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Introduction
1.1 Background
Protecting sediment quality is an important part of restoring and maintaining the biological integrity
of our Nation's waters as well as protecting aquatic life, wildlife and human health. Sediment is an
integral component of aquatic ecosystems, providing habitat, feeding, spawning, and rearing areas for
many aquatic organisms. Sediment also serves as a reservoir for pollutants and therefore a potential
source of pollutants to the water column, organisms, and ultimately human consumers of those
organisms. These pollutants can arise from a number of sources, including municipal and industrial
discharges, urban and agricultural runoff, atmospheric deposition, and port operations.
Contaminated sediment can cause lethal and sublethal effects in benthic (sediment-dwelling) and
other sediment-associated organisms. In addition, natural and human disturbances can release
pollutants to the overlying water, where pelagic (water column) organisms can be exposed. Sediment
pollutants can reduce or eliminate species of recreational, commercial, or ecological importance,
either through direct effects or by affecting the food supply that sustainable populations require.
Furthermore, some sediment pollutants can bioaccumulate through the food chain and pose health
risks to wildlife and human consumers even when sediment-dwelling organisms are not themselves
impacted.
The extent and severity of sediment contamination in the U.S. has been documented in the National
Sediment Inventory (NSI)1 and through other historical information. The NSI screening evaluation
of sediment contamination data indicates that associated adverse effects are probable in thousands of
locations throughout the country. The results emphasize the widespread need to address sediment
contamination in the U.S.
1.2 Significance and Use of this Manual
Sediment quality assessment is an important component of water quality protection programs.
Sediment assessments commonly include physicochemical characterization, toxicity tests, and/or
bioaccumulation tests, as well as benthic community analyses. USEPA's NSI, for example, collates
this information to develop a biennial report to Congress on sediment quality in the United States,
required under the Water Resources Development Act of 1992. The use of consistent sediment
collection, manipulation, and storage methods will help provide high quality samples with which
accurate data can be obtained for the national inventory and for other programs to prevent, remediate,
and manage contaminated sediment.
It is now widely known that the methods used in sample collection, transport, handling, storage, and
manipulation of sediments and interstitial waters can influence the physicochemical properties and
'The National Sediment Inventory, or NSI, is the database of sediment quality information used to
develop EPA's 1997 Report to Congress, The Incidence and Severity of Sediment Contamination in Surface
Waters of the United States, Volume 1: National Sediment Quality Survey (U.S. EPA, 1997a). The database is
updated periodically with new available information on sediment quality at sites throughout the U.S.
http://www.epa.gov/OST/cs/report.html
Chapter 1: Introduction 1 -1
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
the results of chemical, toxicity, and bioaccumulation analyses. Addressing these variables in an
appropriate and systematic manner will help assure more accurate sediment quality data and facilitate
comparisons among sediment studies.
This Technical Manual provides current information and recommendations for collecting and
handling sediments for physicochemical characterization and biological testing, using procedures that
are most likely to maintain in situ conditions, most accurately represent the sediment in question, or
satisfy particular program needs, to help ensure consistent, high quality data collection.
1.3 Applicability and Scope of this Manual
This manual is intended to provide technical support to those who design or perform sediment quality
studies under a variety of regulatory and non-regulatory programs. Information is provided
concerning general sampling design considerations, field and laboratory facilities needed, safety,
sampling equipment, sample storage and transport procedures, and sample manipulation issues
common to chemical or lexicological analyses. Information contained in this manual reflects the
knowledge and experience of several internationally-known sources including American Society for
Testing and Materials (ASTM), Puget Sound Estuary Program (PSEP), Washington State
Department of Ecology (WDE), United States Environmental Protection Agency (USEPA), US Army
Corps of Engineers (ACOE), National Oceanic and Atmospheric Administration (NOAA), and
Environment Canada. This manual attempts to present a coherent set of recommendations on field
sampling techniques and sediment/interstitial water sample processing based on the above sources, as
well as extensive information in the current peer-reviewed literature.
As the scope of this manual is broad, it is impossible to adequately present detailed information on
every aspect of sediment sampling and processing for all situations or all programs. Nor is such
detailed guidance warranted because much of this information (e.g., how to operate a particular
sampling device or how to use a Geographical Positioning System (GPS) device) already exists in
other published materials referenced in this manual. Furthermore, many programs have specific
sampling and sample processing procedures. While an attempt is made to give examples from
different programs, the manual repeatedly instructs the reader to check their own specific program
requirements.
Given the above constraints, this manual: (1) presents an organized discussion of activities involved
in sediment sampling and sample processing; (2) alerts the user to important issues that need to be
considered within each activity; and (3) gives recommendations on how to best address the issues
raised such that appropriate samples are collected and analyzed. An attempt is made to alert the user
to different considerations pertaining to sampling and sample processing depending on the program
need (e.g., dredge remediation versus status and trends monitoring).
Figure 1-1 presents a flow chart of the general activities discussed in this manual. The organization
of these activities reflects the desire to give field personnel and managers a useful tool for choosing
appropriate sampling locations, characterize those locations, collect and store samples, and
manipulate those samples for analyses. Chapters are written so that the reader could obtain
information on only one activity or set of activities (e.g., subsampling or sample processing), if
desired, without necessarily reading the entire manual. Many sections are cross-referenced so that
the reader is alerted to relevant issues that might be covered elsewhere in the manual. This is
particularly important for certain chemical or toxicological applications in which appropriate sample
processing or laboratory procedures are associated with specific field sampling procedures.
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Sediment Monitoring & Assessment Study Plans
(Chapter 2)
' Data Quality Objectives Process
Study Plan Considerations
Sampling Designs
Measurement Quality Objectives
Site-Specific Selection Considerations
for Sediment Sampling Stations
Positioning Methods for Locating
Sampling Stations
Preparations for Field Sampling
. Health & Safety
Collection of Whole Sediments
(Chapter 3)
Types of Sediment Samples
Sample Acceptability
Equipment Decontamination S>-
Field Measurements and Observations
Documentation of Sample Collection I
Collection of
Interstitial Water
(Chapter 6)
In Situ Collection
Ex-Situ Extraction of Interstitial
Water
Field Sample Processing, Transport, and Storage of
Sediments (Chapter 4)
[Sample Containers
Subsampling & Compositing Samples
_^ Homogenizatlon
I Sample Transport and Storage
Sample Holding Times
^
Sediment Manipulations
(Chapter 5)
| Sieving
Formulated Sediment and Organic
I Carbon Modification
~""S Spiking
Preparation of Sediment Dilutions
I Preparation of Sediment Elutriates
Quality Assurance and Quality Control
(Chapter 7)
Procedures for Sediment Collection
and Manipulation
The Quality Assurance Project Plan (QAPP)
Standard Operating Procedures (SOPs)
Sediment Sample Documentation
Sample Tracking Documentation
Record Keeping
QA Audits
Corrective Action (Management of Non-
conformance Events)
Data Reporting
Figure 1-1. Flow chart summarizing activities for collection, storage, and manipulation of
sediments and interstitial water.
Chapter 1: Introduction
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The methods contained in this manual are widely applicable to any entity wishing to collect
consistent, high quality sediment data. This manual does not provide guidance on how to implement
any specific regulatory requirement, or design a particular sediment quality assessment, but rather it
is a compilation of technical methods on how to best collect environmental samples that most
appropriately address common sampling objectives.
Although the data from these samples might be used in environmental decision-making at a variety of
levels, this manual does not address how data are to be used. The Foreword section summarizes a
variety of EPA programs that assess sediment quality and may benefit from the methods described in
this manual. Other Agencies and programs are also encouraged to consider these methods in order to
generate consistent and high quality sediment data.
The information presented in this manual should not be viewed as the final statement on all the
recommended procedures. Some of the areas covered in this document (e.g., sediment holding time,
formulated sediment composition, interstitial water collection and processing) are being actively
researched and debated. As data from sediment monitoring and research becomes more available in
the future, EPA may update this manual as necessary.
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Sediment Monitoring and
Assessment Study Plans
Every study site and project are unique; therefore, sediment monitoring and assessment study plans
should be carefully prepared to best meet the project objectives (MacDonald et al., 1991; see
Figure 2-1).
Considerations
The initial issues that
need to be considered
prior to preparing study
plans are...
define the potential problem or general
project objective
determine what resources (e.g., time,
money, personnel) are available for the
project
review existing information and identify
specific objectives of the study
determine what data are likely to be
needed to answer project objectives,
including the role of site-specific
conditions and/or issues that might
influence the process of data collection
and analyses
Before collecting any environmental data, it is
important to determine the type, quantity, and
quality of data needed to meet the project
objectives (e.g., specific parameters to be
measured) and support a decision based on the
results of data collection and observation. Not
doing so creates the risk of expending too much
effort on data collection (i.e., more data are
collected than necessary), not expending enough
effort on data collection (i.e., more data are
necessary than were collected), or expending the
wrong effort (i.e., the wrong data are collected).
2.1 Data Quality Objectives
Process
The Data Quality Objectives (DQO) Process
developed by EPA (GLNPO, 1994; USEPA,
2000a) is a flexible planning tool that
systematically addresses the above issues in a
coherent manner. The purpose of this process is
to improve the effectiveness, efficiency, and defensibility of decisions made based on the data
collected, and to do so in an effective manner (USEPA, 2000a). The information compiled in the
DQO process is used to develop a project-specific Quality Assurance Project Plan (QAPP) (see
Chapter 7 and USEPA, 2000a) which should be used to plan the majority of sediment quality
monitoring or assessment studies. In some instances, a programmatic QAPP may be prepared, as
necessary, on a project-by-project basis.
The Data Quality Objectives (DQO) process addresses the uses of the data (most importantly, the
decision(s) to be made) and other factors that will influence the type and amount of data to be
collected (e.g., the problem being addressed, existing information, information needed before a
decision can be made, and available resources). From these factors the qualitative and quantitative
data needs are determined (see Figure 2-2). DQOs are qualitative and quantitative statements that
clarify the purpose of the monitoring study, define the most appropriate type of data to collect, and
determine the most appropriate methods and conditions under which to collect them. The products
of the DQO process are criteria for data quality and a data collection design that ensures that data
will meet the criteria.
Chapter 2: Sediment Monitoring and Assessment Study Plans
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Propose general
project goals
Define personnel and budgetary constraints
Review existing data
Data Quality Objectives
Process
Identify specific study objectives and
measurement quality objectives (MQOs)
Define monitoring parameters, sampling
frequency, sampling location, and
analytic procedures
Evaluate hypothetical or, if
available, real data
Will the data meet the proposed
MQOs?
Yes
No
Is the proposed monitoring
program compatible with
available resources?
Yes
No
Modify the
study
procedures
to meet
MQOs
Initiate monitoring activities on a pilot basis
Analyze and evaluate data
QA Implementation
&
Assessment
Does the pilot project meet
the study objectives?
Yes
No
Revise
sampling
and
analysis
plan as
needed
Continue monitoring, data analysis,
and ongoing QA/QC
Reports and recommendations
Figure 2-1. Flow chart summarizing the process that should be implemented in designing and
performing a monitoring study (modified from MacDonald et al. (1991)).
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Stepl.
State the Problem
Define the problem, identify the planning team,
examine budget, schedule.
i
Step 2.
r
Identify the Decision
State decisions, identify study questions, define
alternative actions.
i
Step 3.
r
Identify the Inputs to the Decision
Identify information needed for the decision (information
sources, basis for Action Level, sampling/analysis
methods).
^
Step 4.
r
Define the Boundaries of the Study
Specify sample characteristics, define spatial/temporal
limits, units of decision making.
l
Step 5.
r
Develop a Decision Rule
Define statistical parameter (mean, median), specify
Action Level; develop logic for action.
i
Step 6.
r
Specify Tolerable Limits on Decision Errors
Set acceptable limits for decision errors relative to
consequences (health effects, costs).
i
Step 7.
r
Optimize the Design for Obtaining Data
Select resource-effective sampling and analysis plan
that meets the performance criteria.
Figure 2-2. Flow chart summarizing the Data Quality Objectives
Process (after USEPA, 2000a).
Chapter 2: Sediment Monitoring and Assessment Study Plans
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Checklist
In the DQO process, the following steps should be addressed:
/ Clearly state the problem: purpose and objectives, available resources, members
of the project team: e.g., The purpose might be to evaluate current sediment quality
conditions, historical conditions, evaluate remediation effects, or validate a sediment
model. It is important to review and evaluate available historical data relevant to the
study at this point in the process.
/ Identify the decision; the questions(s) the study attempts to address: e.g., Is site A
more toxic than site B?; Are sediments in Lake Y less toxic now than they used to be?;
Does the sediment at site D need to be remediated? What point or nonpoint sources are
contributing to sediment contamination?
/ Identify inputs to the decision: information and measurements that need to be
obtained; e.g., analyses of specific contaminants, toxicity test results, biological
assessments, bioaccumulation data, habitat assessments, hydrology, and water quality
characterization.
/ Define the study boundaries (spatial and temporal). Identify potential sources of
contamination; determine the location of sediment deposition zones; determine the
frequency of sampling and need for a seasonal sampling and/or sampling during a
specific index period; consider areas of previous dredged or fill material
discharges/disposal. Consideration of hydraulic patterns, flow event frequency, and/or
sedimentation rates could be critical for determining sampling frequency and locations.
/ Develop a decision rule: define parameters of interest and determine the value of a
parameter that would cause follow-up action of some kind; e.g., exceedance of
Sediment Quality Guideline value, NOAA Effect Range Median (ERM) value, or toxicity
effect (e.g., 50% mortality), results in some action (Long et al., 1995). For example, in
the Great Lakes Assessment and Remediation of Contaminated Sediments (ARCS)
Program, one decision rule was: if total PCB concentration exceeds a particular action
level, then the sediments will be classified as toxic and considered for remediation
(GLNPO, 1994). Specifying decision rules or criteria is especially critical in sediment
remediation programs and any study in which the results could be subject to legal
scrutiny (e.g., superfund).
/ Specify limits on decision errors: establish the measurement quality objectives
(MQOs) which include determining the level of confidence required from the data;
precision, bids, representativeness, and completeness of data; the sample size (weight
or volume) required to satisfy the analytical methods and QA/QC program for all
analytical tests; the number of samples required, to be within limits on decision errors,
and compositing needed, if any.
/ Optimize the design: choose appropriate sampling and processing methods; select
appropriate method for determining the location of sampling stations; select an
appropriate positioning method for the site and study. Consult historical data and a
statistician before the study begins regarding the sampling design (i.e., the frequency,
number, and location of field-collected samples) that will best satisfy study objectives.
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For most programs, a Sampling and Analysis Plan (SAP) is developed prior to sampling which
should describe the study objectives, sampling design and procedures, and other aspects of the DQO
process outlined above (see Appendix B for an example of SAP requirements recommended by
Washington State Department of Ecology). The following sections provide guidance on many of the
primary issues that should be addressed in the study plan.
2.2 Study Plan Considerations
Monitoring and assessment studies are performed for a variety of reasons (ITFM, 1995) and sediment
assessment studies can serve many different purposes. Developing an appropriate sampling plan
is one of the most critical steps in monitoring and assessment studies. The sampling plan, including
definition of the site and sampling design, will be a product of the general study objectives
(Figure 2-1). Station location, selection, and sampling methods will necessarily follow from the
study design. Ultimately, the study plan should control extraneous sources of variability or error to
the extent possible so that data are appropriately representative of the sediment and fulfill the study
objectives.
2.2.1 Definition of the Study Area and Study Site
The study area refers to the body of water that contains
the study site(s) to be monitored and/or assessed, as well
as adjacent areas (land or water) that might affect or
influence the conditions of the study site. The study site
refers to the body of water and associated sediments to
be monitored and/or assessed. EMAP, for example,
often defines a site as an area of concern (AOC) which
might extend several miles in length, or may encompass
large geographical or coastal areas. CERCLA defines a
site in terms of a specific source of contamination such
as a waste disposal area.
The size of the study area will greatly influence the type
of sampling design (see Section 2.3) and site positioning
methods that are appropriate (see Section 2.6). The
boundaries of the study area need to be clearly defined at
the outset and should be outlined on a hydrographic chart
or topographic map.
2.2.2 Controlling Sources of Variability
A key factor in effectively designing a sediment quality study is controlling those sources of
variability in which one is not interested (USEPA 2000a,b). There are two major sources of
variability that, with proper planning, can be minimized, or at least accounted for, in the design
process, thereby ensuring a successful study. In statistical terms, the two sources of variability are
sampling error and measurement error (USEPA 2000b; Solomon et al., 1997).
Sampling error is the error attributable to selecting a certain sampling station that might not be
representative of the site or population of sample units (e.g., an estuary or a CERCLA site).
Sampling error is controlled by either: (1) using unbiased methods to select stations if one is
performing general monitoring of a given site (USEPA, 2000b); or (2) several stations along a spatial
gradient if a specific location is being targeted (see Section 2.3).
Common purposes of sediment
quality studies:
• Status and trends
• Evaluating program or BMP (best
management practice)
effectiveness
• Validating sediment quality models
• Designing regulatory programs
• Identifying whether significant
contamination exists and extent of
contamination
• Identifying sources of
contamination
• Ranking existing and identifying
emerging problems
• Establishing goals for sediment
remediation
• Evaluating dredged or fill material
discharges/disposal
Chapter 2: Sediment Monitoring and Assessment Study Plans
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Measurement error is the degree to which the investigator accurately characterizes the sampling unit
or station. Thus, measurement error includes components of natural spatial and temporal variability
within the sample unit as well as actual errors of omission or commission by the investigator.
Measurement error is controlled by using standardized and comparable methods: standardized
methods include proper training of personnel and quality assurance procedures. To help minimize
measurement error, each station should be sampled in the same way, within a site or study, using a
standardized set of procedures and in the same time frame to minimize confounding sources of
variability (see Section 2.2.3). In analytical laboratory or toxicity procedures, measurement error is
estimated by duplicate determinations on some subset of samples (but not necessarily all). Similarly,
in field investigations, some subset of sample units (e.g., 10% of the sites) should be measured more
than once to estimate measurement error (see Replicate and Composite Samples, Section 2.4.3).
Checklist
To minimize measurement
error:
/ Sample all stations similarly within a study
/ Use standardized procedures
/ Sample during the same time period
/ Collect and analyze multiple samples at a
station
/ Collect and analyze composited samples
Measurement error can be reduced by
analyzing multiple observations at each
station (e.g., multiple grab samples at
each sampling station, multiple
observations during a season), or by
collecting depth-integrated, or spatially
integrated (composite) samples (see
Section 2.4.3).
Optimizing sampling design requires
consideration of tradeoffs among the
measures used, the effect that is
considered meaningful, desired power,
desired confidence, and resources
available for the sampling program.
Statistical power is the ability of a given
sampling design to detect an effect that
actually exists, and will be a product of the collection methods, analytical procedures, and quality
control processes used. Power is typically expressed as the probability of correctly finding a
difference among sites or between reference and test sites (e.g., toxicity or biological impairment)
when one exists. For a fixed confidence level (e.g., 90%), power can be increased by increasing the
sample size or the number of replicates (see Section 2.4.3 for more information). Most programs do
not estimate power of their sampling design because this generally requires prior information such as
pilot sampling, which entails further resources. One study (Gilfillan et al., 1995) reported power
estimates for a shoreline monitoring program following the Valdez oil spill in Prince William Sound,
Alaska. However, these estimates were computed after the sampling took place. It is desirable to
estimate power before sampling is performed to ensure credibility of non-significant results (see
Appendix C).
2.2.3 Sampling Using an Index Period
Most monitoring programs do not have the resources to characterize variability or to assess sediment
quality for all seasons. Sampling can be restricted to an index period when biological and/or
lexicological measures are expected to show the greatest response to pollution stress and within-
season variability is small (Holland, 1985; Barbour et al., 1999). This type of sampling might be
especially advantageous for characterizing sediment toxicity, sediment chemistry, and benthic
macroinvertebrate and other biological assemblages (USEPA, 2000c). In addition, this approach is
useful if sediment contamination is related to, or being separated from, high flow events. By
sampling overlying waters during both low and high flow conditions, the relative contribution of
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each to pollutant loads or sediment contamination can be better assessed, thereby better directing
remedial activities, or other watershed improvements.
Those programs that sample the same site over multiple years (e.g., many EMAP and superfund
studies), are interested in obtaining comparable data with which they can assess changes over time,
or following remediation (GLNPO, 1994). In these cases, index period sampling is especially useful
because hydrological regime (and therefore biological processes) is likely to be more similar between
similar seasons than among different seasons.
2.3 Sampling Designs
Sampling Design refers to the array, or network,
of sampling sites selected for a monitoring
program; usually taking one of two forms:
• Probabilistic Design — Network that includes
sampling sites selected randomly in order to
provide an unbiased assessment of the
condition of the waterbody at a scale above
the individual site or stream; can address
questions at multiple scales.
• Targeted Design — Network that includes
sampling sites selected based on known
existing problems, knowledge of upcoming
events in the watershed, or a surrounding
area that will adversely affect the waterbody
such as development or deforestation; or
installation of BMPs or habitat restoration that
are intended to improve waterbody quality;
provides assessments of individual sites or
reaches.
As mentioned in earlier sections of this
chapter, the type of sampling design used is a
function of the study Data Quality Objectives
and more specifically, the types of questions
to be answered by the study. A summary of
various sampling designs is presented in
Figure 2-3 along with recommendations
concerning the conditions under which a
given design is appropriate. Generally,
sampling designs fall into two major
categories: random or probabilistic, and
targeted (USEPA, 2000b). USEPA (2000b;c)
present a thorough discussion of sampling
design issues and detailed information on
different sampling designs. Some program-
specific guidance documents (e.g.,
USEPA/ACOE 1991, 1998 for dredged
material disposal issues) also discuss relevant
sampling designs. Table 2-1 presents
suggested sampling designs given different
overall objectives and constraints. Appendix A presents hypothetical examples of sediment quality
monitoring designs given different objectives or regulatory applications.
2.3.1 Probabilistic and Random Sampling
Probability-based or random sampling designs avoid bias in the results of sampling by randomly
assigning and selecting sampling locations. A probability design requires that all sampling units
have a known probability of being selected. Both EPA's Environmental Monitoring Assessment
Program and NOAA's National Status and Trends Program use a probabilistic sampling design to
infer regional and national patterns with respect to contamination or biological effects.
Sites can be selected on the basis of a truly random scheme or in a systematic way (e.g., sample every
10 meters along a randomly chosen transect). In simple random sampling, all sampling units have
an equal probability of selection. This design is appropriate for estimating means and totals of
environmental variables if the population is homogeneous. To apply simple random sampling, it is
necessary to identify all potential sampling times or locations, then randomly select individual times
or locations for sampling.
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In grid or systematic sampling, the first sampling location is chosen randomly and all subsequent
stations are placed at regular intervals (e.g. 50m apart) throughout the study area. Clearly, the
number of sampling locations could be large if the study area is large and one desires "fine-grained''
contaminant or toxicological information. Thus, depending on the types of analyses desired, such
sampling might become expensive unless the study area is relatively small and/or the density of
stations (that is how closely spaced are the stations) is relatively low. Grid sampling might be
effective for detecting previously unknown "hot spots" in a limited study area.
Sampling Methods
Simple Random: Samples are independently located
at random
Systematic:
Samples are located at regular
intervals
Stratified:
Multistage:
The study area is divided into
nonoverlapping strata and samples
are obtained from each
Large primary units are selected
which are then subsampled
Figure 2-3. Description of various sampling methods. Adapted from USEPA, 2000c.
In stratified designs, the selection probabilities might differ among strata. Stratified random
sampling consists of dividing the target population into non-overlapping parts or subregions (e.g.,
ecoregions, watersheds, or specific dredging or remediation sites) termed strata to obtain a better
estimate of the mean or total for the entire population. The information required to delineate the
strata and estimate sampling frequency must either be known prior to sampling using historic data,
available information and knowledge of ecological function, or obtained in a pilot study. Sampling
locations are randomly selected from within each of the strata. Stratified random sampling is often
used in sediment quality monitoring because certain environmental variables can vary by time of day,
season, hydrodynamics, or other factors. Major environmental monitoring programs that incorporate
a stratified random design include EPA's Mid-Atlantic Integrated Assessment (MAIA). One
disadvantage of using random designs is the possibility of encountering unsampleable sites that were
randomly selected by the computer. Such problems result in the need to reposition the vessel to an
alternate location. Furthermore, if one is sampling to determine the percent spatial extent of
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degradation, it might be important to sample beyond the boundaries of the study area to better
evaluate the limits of the impacted area.
A related design is multistage sampling in which large subareas within the study area are first
selected (usually on the basis of professional knowledge or previously collected information).
Stations are then randomly located within each subarea to yield average or pooled estimates of the
variables of interest (e.g., concentration of a particular contaminant or acute toxicity to Hyalelld) for
each subarea. This type of sampling is especially useful for statistically comparing variables among
specific parts of a study area.
Table 2-1. Suggestions for selecting an appropriate sampling design (from USEPA 2000b).
If you are...
performing a screening phase of
an investigation and with an
understanding of a relatively
small-scale problem
developing an understanding of
when contamination is present
developing an understanding of
where contamination is present
estimating a population mean
estimating a population mean or
proportion
delineating the boundaries of an
area of contamination
estimating the prevalence of a
rare trait
assessing whether a population
contains a rare trait
and you have...
a limited budget and/or a
limited schedule
adequate budget for the
number of samples needed
adequate budget for the
number of samples needed
adequate budget
budget constraints and
analytical costs that are high
compared to sampling costs
budget constraints and
professional knowledge or
inexpensive screening
measurement that can assess
the relative amounts of the
contaminant at specific field
sample locations
spatial or temporal
information on contaminant
patterns
a field screening method
analytical costs that are high
compared to sampling costs
the ability to physically mix
aliquots from the samples
and then retest additional
aliquots
consider
using...
judgmental or
targeted
sampling
systematic
sampling
grid sampling
systematic or
grid sampling
compositing
ranked set
sampling
stratified
sampling
stratified
sampling
random and
composite
sampling
composite
sampling and
retesting
in order to...
assess whether further investigation is
warranted that should include a
statistical probabilistic sampling
design.
have coverage of the time periods of
interest.
have coverage of the area of concern
and have a given level of confidence
that you would have detected a hot
spot of a given size.
also produce information on spatial or
temporal patterns.
produce an equally precise or a more
precise estimate of the mean with
fewer analyses and lower cost.
reduce the number of analyses needed
for a given level of precision.
increase the precision of the estimate
with the same number of samples, or
achieve the same precision with fewer
samples and lower cost.
simultaneously uses all observations
in estimating the mean.
produce an equally precise or more
precise estimate of the prevalence with
fewer analyses and lower cost.
classify all samples at reduced cost by
not analyzing every sample.
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Recommendation Box #1
What type of sampling strategy should be used?
Historical data, if available, should be considered when selecting sampling stations.
Location of sediment depositional zones can be important in defining subareas for
sampling or for stratifying sampling in some programs.
If the objective of the survey is to identify areas of toxic and/or contaminated sediments on
a quantitative spatial and/or temporal basis (e.g., superfund site), a systematic or regular
grid-sampling strategy might be most appropriate (USEPA, 2000b).
If the monitoring objective is to determine sediment contamination originating from a
specific source or tributary, a targeted site location design might be most appropriate.
Factors affecting dispersion of substances or materials from the point source (e.g.,
currents) should be considered.
Stratified random sampling should be used where historical, sediment-mapping data are
available and there are well-defined zones of different sediment types or adjacent land
uses (Burton, 1991). This design is commonly used in NOAA National Status and Trends
(NS&T) monitoring of sediment quality to ensure that the data can be attributed to the
strata in which they were collected (Long et al., 1996).
For dredge management programs, multi-stage, stratified-random, or even targeted
sampling is often appropriate, since the need is to represent specific areas to be dredged
and disposed.
For watershed or regional assessment programs, a probabilistic sampling design might
be most appropriate.
Small-scale, targeted study designs might require many samples within a small area if
fine spatial resolution is needed (e.g., Superfund).
Use of random sampling designs might also miss relationships among variables, especially if there is
a relationship between an explanatory and a response variable. As an example, estimation of benthic
response or contaminant concentration, in relation to a discharge or landfill leachate stream, requires
sampling targeted around the potential contaminant source, including stations presumably unaffected
by the source (e.g., Warwick and Clarke, 1991). A simple random selection of stations is not likely
to capture the entire range needed because most stations would likely be relatively removed from the
location of interest.
2.3.2 Targeted Sampling Designs
In targeted (also referred to as judgmental, or model-based) designs, stations are selected based on
prior knowledge of other factors, such as contaminant loading, depth, salinity, and substrate type.
The sediment studies conducted in the Clark Fork River (Pascoe and DalSoglio, 1994; Brumbaugh et
al., 1994), in which contaminated areas were a focus, used a targeted sampling design.
Targeted designs are useful if the objective of the investigation is to screen an area(s) for the
presence or absence of contamination at levels of concern, such as risk-based screening levels or
toxicity, or to compare specific sediments against reference conditions or biological guidelines. In
general, targeted sampling is appropriate for situations in which any of the following apply (USEPA,
2000b):
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• The site boundaries are well defined or the site physically distinct (e.g., superfund or CERCLA
site, proposed dredging unit).
• Small numbers of samples will be selected for analysis/characterization.
• Information is desired for a particular condition (e.g., "worst case") or location.
• There is reliable historical and physical knowledge about the feature or condition under
investigation.
• The objective of the investigation is to screen an area(s) for the presence or absence of
contamination at levels of concern, such as risk-based screening levels. (Note that if such
contamination is found, follow-up sampling is likely to involve one or more statistical designs to
compare specific sediments against reference conditions, chemical or biological guidelines, or
applicable sediment quality values).
• Schedule or budget limitations preclude the possibility of implementing a statistical design.
• Experimental testing of a known pollution gradient to develop or verify testing methods or
models (i.e., as in evaluations of toxicity tests, Long et al., 1990).
Because targeted sampling designs often can be quickly implemented at a relatively low cost, this
type of sampling can often meet schedule and budgetary constraints that cannot be met by
implementing a statistical design. In many situations, targeted sampling offers an additional
important benefit of providing an appropriate level-of-effort for meeting investigation objectives
without excessive consumption of project resources.
Targeted sampling, however, limits the inferences made to the stations actually sampled and
analyzed. Extrapolation from those stations to the overall population from which the stations were
sampled is subject to unknown selection bias. This bias might be unimportant for those regulatory
programs in which information is needed for a particular condition or location (e.g., Dredged
Management Materials Program or Superfund).
2.4 Measurement Quality Objectives
As noted in Section 2.1, a key aspect of the DQO process is specifying measurement quality
objectives (MQOs): statements that describe the amount, type, and quality of data needed to address
the overall project objectives.
Appendix B presents examples of MQOs and sampling designs that have been used in several
different programs. Also included in Appendix B is excerpted information from Washington
Department of Ecology's Sampling and Analysis Plan Guidance (WDE, 1995). Similar to Quality
Assurance Project Plans (QAPP) mentioned earlier in Section 2.1, a Sampling and Analysis Plan
includes, among other things, many of the elements of the Data Quality Objectives Process, including
MQOs.
A key factor determining the types of MQOs needed in a given project or study is the types of
analyses required because these will determine the amount of sample required (see Section 2.4.1) and
how samples are processed (see Chapter 4). The case examples presented in Appendix B illustrate a
variety of chemical, biological, and toxicological analyses that are often included in sediment quality
monitoring projects. Metals, organic chemicals (including pesticides, PAHs, and PCBs), whole
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sediment toxicity, and organism bioaccumulation of specific target chemicals, are frequently
analyzed in many sediment monitoring programs.
A number of other, more "conventional" parameters, are also often analyzed as well to help interpret
chemical, biological, and toxicological data collected in a project. Table 2-2 summarizes many of the
commonly measured conventional parameters and their uses in sediment quality studies (WDE,
1995). It is important that conventional parameters receive as much careful attention, in terms of
sampling and sample processing procedures, as do the contaminants or parameters of direct interest.
The guidance presented in Chapters 3 and 4 provides information on proper sampling and sample
processing procedures, respectively, to ensure that one has appropriate samples for these analyses.
This section concentrates on three aspects of MQO development that are generally applicable to all
sediment quality studies, regardless of the particular program or objectives: sample volume, number
of samples, and replication vs. composite sampling.
Checklist
MQOs are defined in terms of the following attributes:
/ Detection Limit - The lowest concentration of an analyte that a specified analytical
procedure can reliably detect.
/ Bias - The difference between an observed value and the "true" value (or known
concentration) of the parameter being measured; bias is the first component of accuracy,
which is the ability to obtain precisely a nonbiased (true) value.
/ Precision - The level of agreement among multiple measurements of the same
characteristic; precision is the second component of accuracy.
/ Representativeness - The degree to which the data collected accurately represent the
population of interest (e.g., contaminant concentrations).
/ Comparability - The similarity of data from different sources included within individual or
multiple data sets; the similarity of analytical methods and data from related projects
across areas of concern.
/ Completeness - The quantity of data that is successfully collected with respect to the
amount intended in the experimental design.
2.4.1 Sample Volume
Before commencing a sampling program, the type and number of analyses and tests should be
determined, and the required volume of sediment per sample calculated. Each physicochemical and
biological test requires a specific amount of sediment which, for chemical analyses, depends on the
detection limits attainable and extraction efficiency by the procedure and, for biological testing,
depends on the test organisms and test method. Typical sediment volume requirements for each end
use are summarized in Table 2-3. Specific program guidance should be consulted regarding sample
volumes that might be required.
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Table 2-2. Conventional sediment variables and their use in sediment investigations (Adapted from
WDE, 1995).
Conventional Sediment Variable
Total organic carbon (TOC)
Acid Volatile Sulfide (AVS)
Sediment grain size
Total solids
Ammonia
Total sulfides
Use
• Normalization of the concentrations of
nonionizable organic compounds
• Identification of appropriate reference sediments
for biological tests
• Normalization of the concentrations of divalent
metals in anoxic sediments
• Identification of appropriate reference sediments
for biological tests
• Interpretation of sediment toxicity test data and
benthic macroinvertebrate abundance data
• Evaluation of sediment transport and deposition
• Evaluation of remedial alternatives
• Expression of chemical concentrations on a dry-
weight basis
• Interpretation of sediment toxicity test data
• Interpretation of sediment toxicity test data
When determining the sample volumes necessary, one must know what is required for all of the
sample analyses (considering adequate replication) and it is also helpful to know the general
characteristics of the sediments being sampled. For example, if interstitial water analyses or elutriate
tests are to be conducted, the percent water (or percent dry weight) of the sediment will greatly affect
the amount of water extracted. Many non-compacted, depositional sediments have interstitial water
contents ranging from 30 to 70%. However, interstitial waters are very difficult to remove from
sandy or gravel-rich sediments.
For benthic macroinvertebrate bioassessment analyses, sampling a prescribed area of benthic
substrate is at least as important as sampling a given volume of sediment. In many programs,
macroinvertebrates are sampled using multiple grab samples within a given station location, typically
to a standard sediment depth (e.g., per 10-20 cm of sediment; Klemm et al., 1990; GLNPO, 1994;
Long et al., 1996; USEPA 2000c ). More than 6 liters of sediment from each station might be
necessary in order to have adequate numbers of organisms for analyses, especially in many lakes,
estuaries, and large rivers (Barbour et al., 1999). However, this is very site specific and should be
determined by the field sampling crew. This only applies to whole sediment sampling methods and
not to surficial stream methods using methods such as kick-nets and Surber samplers. If the sediment
quality triad approach is used (i.e., biological, lexicological, and physicochemical analyses
performed on samples from the same sites), more than 10 liters of sediment from each site might be
required depending on the specific analyses conducted. NOAA routinely collects 7-8 liters of
sediment at each station for multiple toxicity tests and chemical analyses (Long et al., 1996).
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Table 2-3. Typical sediment volume requirements for various analyses per sample
Sediment Analysis
Inorganic chemicals
Non-petroleum organic chemicals
Other chemical parameters
(e.g., total organic carbon, moisture content)
Particle size
Petroleum hydrocarbons1
Acute and chronic whole sediment toxicity tests2
Bioaccumulation tests3
Benthic macroinvertebrate assessments
Pore water extraction
Elutriate preparation
Minimum Sample Volume
90 mL
230 mL
300 mL
230 mL
250-1000 mL
1-2 L
15 L
8-16 L
2L
1L
1 The maximum volume (1000 mL) is required only for oil and grease analysis; otherwise, 250
mL is sufficient.
2 Amount needed per whole sediment test (i.e., one species) assuming 8 replicates per sample
and test volumes specified in USEPA, 2000d
3 Based on an average of 3 L of sediment per test chamber and 5 replicates (USEPA, 2000d).
Recommendation Box #2
How many samples and how much sample volume should be
collected?
The testing laboratory should be consulted to confirm the amount of sediment required
for all desired analyses.
The amount of sediment needed from a given site will depend on the number and types
of analyses to be performed. If biological, lexicological, and chemical analyses are
required (sediment triad approach), then at least 10 liters of sediment might be required
from each station.
Since sampling events might be expensive and/or difficult to replicate, it is useful to
collect extra samples if possible, in the event of problems encountered by the analytical
laboratories, failure of performance criteria in assays, or need to verify/validate results.
Consider compositing samples from a given station or across similar station types to
reduce the number of samples needed.
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Considerations
The appropriate number
of samples is usually
determined by...
size of the study site
type and distribution of the
contaminants being measured
characteristics and homogeneity of the
sediment
concentrations of contaminants likely to
be found in the sediments
sample volume requirements
desired level of statistical resolution or
precision
2.4.2 Number of Samples
The number of samples collected directly affects
the representativeness and completeness of the
data for purposes of addressing project goals. As
a general rule, a greater number of samples will
yield better definition of the areal extent of
contamination or toxicity. Many programs
specify a certain number of samples per location
(e.g., CERCLA site or dredging unit).
Accordingly, sample requirements should be
determined on a case-by-case basis. The number
of samples to be collected will ultimately be an
outcome of the questions asked. For example, if
one is interested in characterizing effects of a
point source or a gradient (e.g., effects of certain
tributaries or land uses on a lake or estuary), then
many samples in a relatively small area might
need to be collected and analyzed. If, however,
one is interested in screening "hot spots" or locations of high contamination within a watershed or
water body, relatively few samples at regularly-spaced locations might be appropriate. In most
monitoring and assessment studies, the number of samples to be collected usually results from a
compromise between the ideal and the practical. The major practical constraints are the costs of
analyses and logistics of sample collection.
The major costs associated with the collection of sediment samples are those for travel to the site and
for sample analysis. The costs of actual on-site sampling are minimal by comparison. Consequently,
it is good practice to collect an excess number of samples, and a subset equal to the minimum
number required is selected for analysis. The archived replicate samples can be used to replace lost
samples, for data verification, to rerun analyses yielding questionable results, or for the independent
testing of a posteriori hypotheses that might arise from screening the initial data. However, storage
of sediments might result in changes in bioavailability of chemical contaminants (see Section 4.5).
Therefore, follow-up testing of archived samples should be done cautiously.
2.4.3 Replicate and Composite Samples
Replicate Samples
As mentioned in the previous section, the number of samples collected and analyzed will always be a
compromise between the desire of obtaining high quality data that fully addresses the overall project
objectives (MQOs) and the constraints imposed by analytical costs, sampling effort, and study
logistics. Therefore, every sampling program needs to find a balance between obtaining information
to satisfy the stated DQOs or study goals in a cost-effective manner, and yet have enough confidence
in the data to make appropriate decisions (e.g., remediation, dredging; Step 3 in the DQO process,
Figure 2-2). Two different concepts are used to satisfy this challenge: replication and sample
compositing.
Replication is used to assess precision of a particular measure and can take many forms depending on
the type of precision desired. For most programs, analytical replicates are the most frequently used
form of replication because most MQOs are concerned with analytical data quality (see examples in
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Appendix B). The extent of analytical replication (duplicates) varies with the program or study
DQOs. Performing duplicate analyses on at least 10% of the samples collected is considered
satisfactory for most programs (GLNPO, 1994; USEPA/ACOE, 1991; PSEP, 1997a; USEPA/ACOE,
1998). An MQO of < 20 - 30% relative percent difference (RPD) is commonly used for analytical
replicates depending on the analyte.
Checklist
Replication can take several
forms and satisfy different
purposes:
Collect field replicate samples at a station if
there is a need to statistically compare
results among stations within a site.
Analytical replicates: separate laboratory
analyses on subsamples from the same
field sample.
Field replicates: separate samples collected
at a station each of which is analyzed
individually.
Field-split replicates: a single field sample is
split into subsamples, each of which is then
analyzed individually.
Compositing samples is one way to reduce
the number of replicates needed for
analysis.
Field replicates can provide useful
information on the spatial distribution of
contaminants at a station and the
heterogeneity of sediment quality within
a site. Furthermore, field replicates
provide true replication at a station
(analytical replicates and split samples at
a station provide a measure of precision
for a given sample, not the station) and
therefore can be used to statistically
compare analyses (e.g., toxicity, tissue
concentration, whole sediment
concentration) across stations.
Results of field replicate analysis yield
the overall variability or precision of both
the field and laboratory operations (as
well as the variability between the
replicate samples themselves, apart from
any procedural error). Because field
replicate analyses integrate a number of
different sources of variability, they
might be difficult to interpret. As a
result, failure to meet a precision MQO
for field replicates might or might not be a cause of concern in terms of the overall study objectives
but would suggest some uncertainty in the data. Many monitoring programs perform field replicates
at 10% of the stations sampled in the study as a quality control procedure. An MQO of < 30 - 50%
relative percent difference (RPD) is typically used for field replicates depending on the analyte (see
examples in Appendix B). Many regulatory programs (e.g., Dredged Disposal Management within
the Puget Sound Estuary Program) routinely use 3-5 field replicates per station. Appendix C
summarizes statistical considerations in determining the appropriate number of replicate samples
given different sampling objectives.
Split sample replication is less commonly performed in the field because many programs find it more
useful to quantify data precision through the use of analytical and field replicates described above.
However, split sample replication is frequently used in the laboratory in toxicity and bioaccumulation
analyses (USEPA, 2000d) and to verify homogeneity of test material in spiked sediment tests (see
Section 5.3). In the field, samples are commonly split for different types of analyses (e.g., toxicity,
chemistry, benthos) rather than to replicate a given sample. This type of sample splitting or
subsampling is further discussed in Section 4.2.
Composite Samples
A composite sample is one that is formed by combining material from more than one sample or
subsample. Because a composite sample is a combination of individual aliquots, it represents an
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"average" of the characteristics making up the
sample. Compositing, therefore, results in a less
detailed description of the variability within the
site as compared to taking field replicates at each
station. However, for characterizing a single
station, compositing is generally considered an
excellent way to provide quality data with
relatively low uncertainty. Furthermore, many
programs find it useful to average the naturally
heterogeneous physicochemical conditions that
often exist within a station (or dredging unit, for
example), even within a relatively small area
(GLNPO, 1994; PSEP, 1997a; ASTM, 2000a).
Many programs find it useful to composite 3-5
samples from a given location or depth strata
(PSEP, 1997a; GLNPO, 1994).
Considerations
Composite samples are
collected because
they...
Yield a single "average estimate for a
given station with less cost than using
replicates.
Can obtain useful information over
many stations at reduced analytical
costs.
Are an efficient way to provide sufficient
sample volume for multiple types of
analyses, particularly biological/toxicity
analyses.
Compositing is also a practical way to control analytical costs while providing information from a
large number of stations. For example, with relatively little more sampling effort, five analyses can
be performed to characterize a project segment or site by collecting 15 samples and combining sets of
three into five composite samples. The increased coverage afforded by taking composite samples
might justify the increased time and cost of collecting the extra 10 samples in this case
(USEPA/ACOE, 1998). Compositing is also an important way to provide the large sample volumes
required for some biological tests (see Table 2-1) and for multiple types of analyses (e.g., physical,
chemical, toxicity, and benthos). However, compositing is not recommended where combining
samples could serve to "dilute" a highly toxic but localized sediment "hot spot" (WDE, 1995;
USEPA/ACOE, 1998). Also, samples from stations with very different grain size characteristics or
different stratigraphic layers of core samples should not be composited (see Section 4.3).
Checklist
Before sampling:
/ Review available information about the site including physical conditions and potential
contaminant sources.
/ Inspect the site to confirm that the sampling design and procedure chosen are feasible.
/ Perform a pilot or screening sampling, if possible, to ensure that sampling equipment and
procedures are adequate for the types of stations selected.
2.5 Site-Specific Considerations for Selecting Sediment Sampling
Stations
Several site-specific factors might ultimately influence the appropriate location of sampling stations,
both for large-scale monitoring studies, in which general sediment quality status is desired, and for
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smaller, targeted studies, in determining the need for sediment remediation. If a targeted or stratified
random sampling design is chosen, it might be important to locate sediment depositional and
erosional areas to properly identify contaminant regimes. Table 2-4 presents a summary of site-
specific factors that should be considered when developing a sampling plan. A comprehensive
review of such considerations is provided by Mudroch and MacKnight (1994).
Table 2-4. Practical considerations for site-specific selection of sampling stations in developing a
sampling plan.
Activity
Consideration
Determination of areas where
sediment contamination might occur
Hydrologic information
• quality and quantity of runoff
• potential depositional inputs of total suspended solids
• up-wellings
• seepage patterns
Determination of depositional and
erosional areas
Bathymetric maps and hydrographic charts
• water depth
• zones of erosion, transport, and deposition
• bathymetry
• distribution, thickness, and type of sediment
• velocity and direction of currents
• sedimentation rates
Climatic conditions
• prevailing winds
• seasonal changes in temperature, precipitation, solar radiation,
etc.
• tides, seiches
• seasonal changes in anthropogenic and natural loadings
Determination of potential sources
of contamination
Anthropogenic considerations
location of urban centers
• historical changes in land use
• types, densities, and size of industries
• location of waste disposal sites
• location of sewage treatment facilities
• location of stormwater outfalls and combined sewer overflows
• location, quantity, and quality of effluents
• previous monitoring and assessment or geochemical surveys
• location of dredging and open-water dredged material disposal
sites
• location of historical waste spills
Factors affecting contaminant
bioavailability
Geochemical considerations
• type of bedrock and soil/sediment chemistry
• physical and chemical properties of overlying water
Determination of representativeness
of samples
area to be characterized
volume to be characterized
depth to be characterized
possible stratification of the deposit to be characterized
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2.5.1 Review Available Data
Review of available historical and physical data is critical in the sample selection process and
subsequent data interpretation. Local experts should be consulted to obtain information on site
conditions and the origin, nature, and degree of contamination. Other potential sources of
information include government agency records, municipal archives, harbor commission records,
past geochemical analyses, hydrographic surveys, bathymetric maps, and dredging/disposal history.
Potential sources of contamination should be identified and their locations noted on a map or chart of
the proposed study area. It is important that recent hydrographic or bathymetric data be used in
identifying representative sampling locations, especially for dredging or other sediment removal
projects. The map or chart should also note adjacent land and water uses (e.g., fuel docks, storm
drains, etc.). The quality and age of the available data should be critically weighed.
2.5.2 Site Inspection
A physical inspection of the site is strongly recommended when developing a study plan, in order to
assess the completeness and validity of the collected historical data, and to identify any significant
changes that might have occurred at the site or study area (Mudroch and MacKnight, 1994). A site
inspection of the immediate drainage area and upstream watershed might also identify potential
stressors (such as erosion), and help determine appropriate sampling gear (such as corer vs. grab
samplers and boat type) and sampling logistics.
If resources allow, it is useful to perform some screening or pilot sampling and analyses at this stage
to further refine the actual sampling design needed. Pilot sampling is particularly helpful in defining
appropriate station locations for targeted sampling or to identify appropriate strata or subareas in
stratified or multistage sampling, respectively.
2.5.3 Identify Sediment Deposition and Erosional Zones
When study DQOs direct sampling to the highest contamination levels or specific subareas of a site,
it might be important to consider sediment deposition and sediment erosional zones, since grain size
and related physicochemical characteristics (including conventional parameters such as total organic
carbon and acid volatile sulfide, as well as contaminants), are likely to vary between these two types
of zones. Depositional zones typically contain fine-grained sediment deposits which are targeted in
some sampling programs because fine-grained sediments tend to have higher organic carbon content
(and are therefore a more likely repository for pollutants) relative to larger sediment particle size
fractions (e.g., sand and gravel) (ASTM, 2000a; Environment Canada, 1994). However, for some
programs such as remediation dredging evaluations or superfund, eroding sediment beds and non-
depositional zones might be of most concern as these could be a major source of pollutants in the
water column and in organisms (USEPA/ACOE, 1991,1998).
Various non-disruptive technologies are available to assist in the location of fine-grained sediments
ranging from simplistic to more advanced. For example, use of a steel rod or PVC pipe can be used
in many shallow areas to quickly and easily probe the sediment surface to find coarse (sand, gravel)
vs. fine sediments (silt, clay). This technique can not, however, determine sediment grain size at
depth. Other more advance methods, including acoustic survey techniques (e.g., low frequency echo
sounding, seismic reflections, etc.) and side-scan sonar used with a sub-bottom profiler (Wright et al.,
1987), can provide useful information on surficial as well as deeper sediment profiles. However,
these techniques are often limited in their accuracy and have high equipment costs (Guigne et al.,
1991).
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Aerial reconnaissance, with or without satellite imagery, might assist in visually identifying
depositional zones where clear water conditions exist. These methods are not reliable, however, if
waters are turbid. Other methods that can be used to locate sediment deposition zones include grab
sampling, inspection by divers, or photography using an underwater television camera or remotely
operated vehicle (Burton, 1992; ASTM, 2000a).
2.6 Positioning Methods for Locating Sampling Stations
The most important function of positioning technology is to determine the location of the sampling
station (e.g., latitude and longitude), so that the user can later re-sample to the same position
(USEPA, 1987). Knowing the precise location of sampling stations is also important so that
regulators can determine if the area(s) of interest have been sampled. There are a variety of
navigation and/or position-fixing systems available, including optical or line-of-site techniques,
electronic positioning systems, and satellite positioning systems. Global Positioning System (GPS)
is generally regarded as the positioning technique of choice as it is accurate, readily available, and
often less expensive than many other comparably sophisticated systems. Given the removal of
selective availability of satellite data by the U.S. military, GPS is now capable of high accuracy
positioning (1-10 m). The characteristics, advantages, and disadvantages of a variety of positioning
systems are summarized in Appendix D.
Recommendation Box #3
How should station positioning be performed?
Depending on level of accuracy needed, regular calibration of the positioning system by at
least two methods might be required to ensure accuracy.
For monitoring and assessment studies of large areas (e.g., large lakes or offshore
marine environments), where an accuracy of ± 100 m typically is sufficient, either the
Long Range Navigation (LORAN) or Global Positioning System (GPS) system is
recommended.
For near-shore areas, or areas where the sampling stations are numerous or located
relatively close together, GPS or a microwave system should be used if the required
position accuracy is less than 10 m. Where visible or suitable and permanent targets are
available, RADAR can be used if the required position accuracy is between 10 and 100
m.
For small water bodies and urban waterfronts, GPS is often capable of giving precise
location information. Alternatively, visual angular measurements (e.g., sextant) by an
experienced operator, a distance line, or taut wire could also provide accurate and
precise positioning data.
Regardless of the type of system selected, calibration of the system is recommended by using at least
two of these methods to ensure accuracy particularly for stations that will be reoccupied. At each
sampling station, a fathometer or meter wheel can be used to determine the sampling depth. This
will ensure that the water is the desired depth and the bottom is sufficiently horizontal for proper
operation of sampling equipment. Ideally, it is best to print out a copy of the ship's location from the
GPS monitor navigation chart, as well as the latitude / longitude, so the sampling station can be
placed in a spatial context.
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2.7 Preparations for Field
Sampling
Proper preparation for any field sampling
study is an essential part of Quality
Assurance that ensures a successful project
outcome and adherence to the objectives
specified in the Quality Assurance Project
Plan (QAPP). Chapter 7 further discusses
related Quality Assurance/Quality Control
procedures that should be used in sediment
quality studies.
Prior to performing field work,
characteristics of the site and accessibility of
the individual sampling stations should be
ascertained. Pictures of sampling stations
both before as well as during sampling are
often useful to ensure that the correct stations
were sampled and to document weather and
water conditions during sampling. Adequate
reconnaissance of stations prior to sampling
is also valuable for preparing against
potential sampling hazards or unforeseen
difficulties. Such a reconnaissance can also
help determine the necessary time needed to
perform the desired sampling (i.e., time to get
from one station to the next).
The appropriate vessel or sampling platform is one of the most important considerations in preparing
for field sampling. The vessel must be appropriate for the water body type, and should provide
sufficient space and facilities to allow collection, any on-board manipulation, and storage of samples.
Ice chests or refrigeration might be required for sample storage, depending on the time course of the
operation. The vessel should provide space for storage of decontamination materials, as well as
clean sampling gear and containers to avoid contamination risks associated with normal vessel
operations. Space for personal safety equipment is also required.
Additionally, the vessel should be equipped with sufficient winch power and cable strength to handle
the weight of the sampling equipment, taking into account the additional suction pressure associated
with extraction of the sediments. Large sampling devices typically weigh between 50 and 400 kg
empty, and when filled with wet sediment might weigh from 125 to over 500 kg.
Care should be taken in operating the vessel to minimize disturbances of the sediment to be sampled
as well as sampling equipment. This would include physical disturbance through propellar action
and chemical contamination from engines or stack emissions. For example, Page et al. (1995)
reported that they positioned the ships' stern into the wind to prevent stack gases from blowing onto
sampling equipment during deployment, recovery, and subsampling of sediments in Prince William
Sound, Alaska.
Checklist
Logistical
Considerations:
site description
study site accessibility
adequate sampling platform
qualified personnel
specific responsibilities of field crew
locating and maintaining stations
adequate time for sampling
adequate space and equipment
communication system
access to temporary field storage
health, safety, and waste management
emergency plans and equipment
number of samples to be collected
sample holding times
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The sampling plan and projected time schedule should be posted for view by all personnel. The
names, addresses, and telephone numbers of all participants involved with the preparation and
execution of the sampling program should be available to all participants, and the duties and
responsibilities of each participant clearly documented. The study supervisor should ensure that the
appropriate personnel clearly understand their role and are capable of carrying out their assigned
responsibilities and duties. Contingency planning should address the need for backup personnel in
the event of accident or illness.
A variety of sampling and sample handling equipment and supplies are often needed in sediment
monitoring studies. Besides the actual samplers themselves (e.g., grab or core device to be used),
equipment is needed to remove and process the samples such as spatulas, scoops, pans or buckets,
and gloves. If it is important to maintain anoxic conditions of the sample, a glovebox and inert gas
source (e.g., nitrogen) is needed. Sample storage and transport equipment and supplies need to be
available as well. These include refrigeration, ice chests, dry ice or ice, insulation material to
stabilize samples in transport, custody seals, and shipping airbills.
The reagents for cleaning, operating or calibrating equipment, and/or for collecting, preserving or
processing samples should be handled by appropriately qualified personnel and the appropriate data
for health and safety (e.g., Material Safety Data Sheets) should be available. Written approved
protocols and standard operating procedures (including QA/QC requirements) should be readily
accessible at all times, to ensure proper and safe operation of equipment. Data forms and log books
should be prepared in advance so that field notes and data can be quickly and efficiently recorded.
Extra forms should be available in the event of a mishap or loss. These forms and books should be
waterproof and tear resistant. Under certain circumstances audio or audio/video recordings might
prove valuable.
All equipment used to collect and handle samples must be cleaned and all parts examined to ensure
proper functioning before going into the field. A repair kit should accompany each major piece of
equipment in case of equipment failure or loss of removable parts. Backup equipment and sampling
gear should be available.
Checklist
Equipment and/or reagents needed:
/ sampling equipment and spare parts
/ sample handling equipment
/ special sample handling equipment (e.g., glovebox or shielded compartment).
/ decontamination and cleaning equipment
/ field measurement equipment and supplies
/ sample storage supplies/equipment
/ sample transport supplies
/ personnel supplies
/ maps, navigation, and communication equipment
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Storage, transport, and sample containers, including extra containers should be available in the event
of loss or breakage (see Section 4.2 for more information on appropriate containers). These
containers should be pre-cleaned and labeled appropriately (i.e., with a waterproof adhesive label to
which the appropriate data can be added, using an indelible ink pen capable of writing on wet
surfaces). The containers must have lids that are fastened securely, and if the samples are collected
for legal purposes, they should be transported to and from the field in a locked container with
custody seals secured on the lids. Samples to be frozen before analyses must not be filled to the lids.
Leave a 10% headspace to accommodate expansion during freezing. Whether for legal purposes or
not, all samples should be accompanied by a chain-of-custody form that documents field samples to
be submitted for analyses (see Chapter 7). Transport supplies also include shipping airbills and
addresses.
A sample-inventory log and a sample-tracking log should be prepared in advance of sampling. A
single person should be responsible for these logs who will track the samples from the time they are
collected until they are analyzed and disposed of or archived.
2.8 Health and Safety
Collection and processing of sediments for analyses and testing might involve substantial risks to
personal safety and health; particularly in situations involving potentially hazardous materials or
challenging sampling conditions. If a Quality Assurance Project Plan (QAPP) or a Sampling and
Analysis Plan (SAP) is prepared prior to sampling, it should include or reference health and safety
procedures. A health and safety field officer should be appointed to ensure that personnel use safety
precautions and equipment applicable to the operation of the vessel, the sampling equipment, and
sample handling. Personnel collecting or handling sediment samples should not work alone, and they
should take all safety precautions necessary for the prevention of bodily injury and illness which
might result from sampling activities (e.g., boat safety), ingestion or invasion of infectious agents,
inhalation or absorption of corrosive or toxic substances through skin contact, or asphyxiation.
Because sediment collection often occurs without complete knowledge of the source or degree of
hazard, contact with sediment should to be minimized by: (1) using gloves, laboratory coats, safety
glasses, face shields and respirators, as appropriate, and (2) manipulating sediments in open air,
under a ventilated hood, or in an enclosed glove box. USEPA (1986a), Walters and Jameson (1984),
and the Occupational Health and Safety Administration (OSHA) standards provide guidance on safe
sediment handling. Program specific guidance should be consulted first when available (e.g.,
Washington Department of Ecology's Sampling and Analysis Plan Guidance [WDE, 1995] or Puget
Sound Estuaries Program [PSEP, 1997a]). Other references (e.g., ASTM, 2000b; Waters, 1980)
should also be consulted concerning special safety procedures for sampling and handling samples
from hazardous waste sites. The NOAA Diving Manual (NOAA, 1991) or the EPA Diving Safety
Manual (USEPA, 1997b) should be consulted for information regarding diving safety plans and
protocols.
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Recommendation Box #4
What health and safety precautions should be followed?
Follow Coast Guard approved safety procedures, including use of life vests.
All samples must be handled in a manner that satisfies the Quality Assurance Project
Plan, Standard Operating Procedures, and DQOs.
Skin contact with sediment should be minimized to avoid potential contact with
hazardous substances. Protective clothing and equipment (e.g., gloves, boots, lab coats
or aprons, safety glasses, and respirator) are recommended during sampling, sample
handling, and preparation of test substances or sediments.
Handling of samples should be performed in a well-ventilated area (e.g., outside, in a
fume hood, or in an enclosed glove box) to minimize the inhalation of sediment gases
such as hydrogen sulfide if present.
A spill control protocol should be in place in the sampling vessel and laboratory.
Disposal of all hazardous waste should be in accordance with applicable laws,
guidelines, and regulations.
Provide procedures regarding hazard assessment (chemical and physical hazards).
Provide procedures regarding decontamination.
Meet the training and medical monitoring requirements.
Provide emergency planning and emergency contacts.
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Collection of Whole Sediments
Most sediment collection devices are designed to isolate and consistently retrieve a specified volume
and surface area of sediment, from a required depth below the sediment surface, with minimal
disruption of the integrity of the sample and no contamination of the sample. Maintaining the
integrity of the collected sediment, for the purposes of the measurements intended, is a primary
concern in most studies because disruption of the sediment's structure could change its
physicochemical and biological characteristics, thereby influencing the bioavailability of
contaminants and the potential toxicity of the sediment. This chapter discusses the factors to be
considered in selecting a sediment collection device. A variety of samplers are described (and
pictured in Appendix E), and recommendations are made regarding their use in different situations.
The flowchart in Figure 3-1 shows recommended sampling gear based on monitoring objective or
site-specific issues of concern. Figures 3-2 and 3-3 provide recommended grab and core samplers,
respectively, based on site factors
(such as depth and particle size),
and sampling requirements (such
as sample depth and volume of
sample needed).
3.1 General
Procedures
• Characterization of
contamination in deeper
sediments is important
• Comparison of recent
surficial vs. historical
deeper sediments
• Reduced sediment gradient
disruption needed
• Reduced oxygen exposure
needed
• Sediments soft and fine
grained
1 Large sediment volumes
needed
1 Larger grained sediments
are common
1 Larger surface area of
surficial sediment needed
See Kgure 3-2
See Figure 3-3
Figure 3-1. General types of considerations or objectives that
are appropriate for grab or core sampling devices.
The planned mode of access to
the sampling area (e.g., by water,
over land or ice, or from the air)
plays an important role in the
selection of sampling gear. If the
sampling gear needs to be
transported to a remote area or
shipped by air, its weight and
volume might need to be taken
into account. It is often the case
that a specific vessel, having a
fixed lifting capacity based on the configuration of its winch, crane, boom, A-frame, or other support
equipment, is the only one available for use. This will affect the type of sampling equipment that can
be safely operated from that vessel.
Many samplers are capable of recovering a relatively undisturbed sample in soft, fine-grained
sediments, but fewer are suitable for sampling harder sediments containing significant quantities of
sand, gravel, firm clay, or till (Mudroch and Azcue, 1995). One of the most important factors in
determining the appropriate sampling device for the study are Data Quality Objectives. Many
monitoring programs, such as EPA's Environmental Monitoring and Assessment Program (EMAP)
and the NOAA National Status and Trends program, are primarily interested in characterizing recent
environmental impacts in lakes, estuaries and coastal waters and therefore sample surface sediments
Chapter 3: Collection of Whole Sediments
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
I Factor: Water Depth
Ponar, Ponar, Van Veen,
Petersen mini-Shipek
< 4m or very low current
conditions; smooth water
> 4m or mild-moderate
current
( Factor: Particle Size J
Birge-Ekman (and mini),
vJPonar (and mini), VanVeen
Shipek
Compacted Sediments
( Factor: Able to subsample directly from sampler? )
Ponar, Smith-Maclntyre,)
Petersen
Birge-Ekman (and mini),
Ponar (and mini),
Van Veen, Shipek
Factor: Sample depth (surficial
sediment only vs. a broader
biologically active layer)
Petersen, Smith-
Maclntyre, mini-Shipek
Birge-Ekman (and mini),
vPonar (andmini),Shipek (andmini),
Mini Birge-Ekman,
petite Ponar, Shipek, mini-Shipek
Factor: Sample volume (affects
how many samples needed per
site for all analyses)
Smith-Maclntyre,
Van Veen,Petersen,
I
Birge-Ekman, Ponar, Petersen
Smith-Maclntyre, Van Veen
Figure 3-2. Flowchart for selecting appropriate grab samplers based on site-specific or design factors
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(Factor: Water Depth)
Phleger, Kajak-BrinkhurstT
Alpine, gravity, Box Corer,
Vibracorer
Soft sediment only
Factor: Particle Size J
( — ^ak-B
\_gravity,
BoxCor
Semi-consolidated and
Soft sediment
Piston, Boomerang^
Vibratory
'Tube, Hand Corer/
Phleger, Alpine
Corer, Tube, Box Corer;
Phleger, Kajak-Brinkhurst,
Boomerang
Factor: Sample depth: how deep a profile can
be obtained
Gravity, Alpine,
BMH-53,Pisto
Piston, Boomerang,
Vibratory
:Factor: Sample volume: indication of how
many cores may be required per site
Hand Corer, Tube,
Phleger
Box, Gravity, Piston,
Boomerang, Vibratory
Kajak-Brinkhurst,
Alpine, BMH-53, Piston
Factor: Lifting capacity needed: indication
of boat and which equipment needed
Box, Gravity, Alpine,
Piston, Vibrai
Phleger, Kajak-Brinkhurst,
Boomerang
Figure 3-3. Flowchart for selecting appropriate core samplers based on site-specific factors
Chapter 3: Collection of Whole Sediments
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Considerations
Although there is no
one sampler that
satisfies all possible
considerations, the
ideal sampler...
avoids a pressure wave
penetrates cleanly to minimize
disturbance
closes tightly
allows for subsampling
can accommodate weighting
collects sufficient sediment volume
retrieves sediment from a wide range
of water and sediment depths
does not contaminate the sample
is easy and safe to operate
is easily transported/assembled at the
site
(e.g., Long et al., 1996). Other programs (e.g.,
dredged material characterization studies
conducted for EPA and the US Army Corps of
Engineers), are concerned with the vertical
distribution of contaminants in sediment to be
dredged and therefore seek to characterize a
sediment column (USEPA/ACOE, 1991,1998).
Each program would employ different sampling
devices.
Related to study objectives, another important
factor in selecting a sampler is desired depth of
sediment penetration. For monitoring and
assessment studies where historical
contamination is not the focus, the upper 10 to 15
cm is typically the horizon of interest. Generally,
the most recently deposited sediments and most
epifaunal and infaunal organisms are found in
this horizon. To ensure minimum disturbance of
the upper layer during sampling, a minimum
penetration depth of 6 to 8 cm is recommended,
with a penetration depth of 10 to 15 cm being
preferred. However, if sediment contamination is
being related to organism exposures (e.g., benthic
macroinvertebrates and/or fish) then more precise
sampling of sediment depths might be needed,
such as with a core sampler. The life history and
feeding habits of the organisms (receptors) of concern should be considered. For example, some
organisms (e.g., shrimp, rotifers) might be epibenthic and are only exposed to surficial sediments
(e.g., 0 to 1 cm) while others (e.g., amphipods, polychaetes) that are infaunal irrigators might receive
their primary exposure from sediments that are several centimeters in depth. Relating contaminant
levels that occur in sediment layers other than where resident organisms are exposed, might produce
incorrect conclusions.
Sampling of the surface layer provides information on the horizontal distribution of parameters or
properties of interest for the most recently deposited material. Information obtained from analysis of
surface sediments can be used, for example, to map the distribution of a chemical contaminant in
sediments across a specific body of water (e.g., lake, embayment, estuary). A sediment column,
including both the surface sediment layer and the sediment underneath this layer, is collected to study
historical changes in parameters of interest (as revealed through changes in their vertical distribution)
and to characterize sediment quality with depth.
Once study objectives and the general type of sampler have been identified, a specific sampler is
selected based on knowledge of the bathymetry and areal distribution of physically different
sediment types at the sampling site. Therefore, it is strongly recommended that this information be
gathered during the initial planning stage of all sample collection efforts (see Section 2.5.1).
The quantity of sediment to be collected at each sampling site may also be an important
consideration in the selection of a sampling device (see also Section 2.4.1). The required quantity of
sediment typically depends on the number and type of physicochemical and biological tests to be
carried out (See Table 2-3 for typical sediment volumes needed for different analyses).
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Regardless of the type of sampler used, it is important to follow the standard operating procedures
specific to each device. Before retrieving the sample, the outside of the sampling device should be
carefully rinsed with water from the sampling station. Between each sampling event, the sampling
device should be cleaned, inside and out, by dipping the sampler into and out of the water rapidly or
by washing with water from the location being sampled. More rigorous between-sample cleaning of
the sampler (e.g., chemical decontamination or washing with soap) might be required, depending on
the nature of the investigation and specific program guidance (see Section 3.5).
To minimize cross-contamination of samples and to reduce the amount of equipment
decontamination required, it might be prudent to sample reference sites (i.e., relatively clean sites)
first, followed by test stations. If certain stations are known to be heavily contaminated, it might be
prudent to sample those stations last when sampling many locations at one time.
3.2 Types of Sediment Samplers
There are three main types of sediment sampling devices: grab samplers, core samplers, and
dredge samplers. Grab samplers (see Appendix E) are typically used to collect surficial sediments
for the assessment of the horizontal distribution of sediment characteristics. Core samplers (see
Appendix E) are typically used to sample thick sediment deposits, or to collect sediment profiles for
the determination of the vertical distribution of sediment characteristics or to characterize the entire
sediment column. Dredge samplers are used primarily to collect benthos. Dredges cause disruption
of sediment and pore water integrity, as well as loss of fine-grained sediments. For these reasons,
only grab and core samplers are recommended for sediment physicochemistry or toxicity evaluations.
Since many grab samplers are appropriate for collecting benthos as well (Klemm et al., 1990; ASTM,
2000c), grab samplers are likely to be more useful than dredges in sediment quality assessments.
Therefore, dredges are not considered further in this document.
Advantages and disadvantages of various grab and core samplers are summarized in Appendix
Tables E-l and E-2, respectively, and are discussed briefly in the following sections. Figure 3-1
provides recommendations regarding the type of sampler that would be appropriate given different
study objectives. For many study objectives either cores or grab samplers can be used, however, in
practice, one will often be preferred over the other depending on other constraints such as amount of
sample required for analyses and equipment availability.
3.2.1 Grab Samplers
Grab samplers consist either of a set of jaws that shut when lowered into the surface of the bottom
sediment or a bucket that rotates into the sediment when it reaches the bottom (see Appendix E).
Grab samplers have the advantages of being relatively easy to handle and operate, readily available,
moderately priced, and versatile in terms of the range of substrate types they can effectively sample.
Of the grab samplers, the Van Veen, Ponar (see photograph on page 3-6), and Petersen are the most
commonly used. These samplers are effective in most types of surface sediments and in a variety of
environments (e.g., lakes, rivers, estuaries, and marine waters). In shallow, quiescent water, the
Birge-Ekman sampler also provides acceptable samples and allows for relatively nondisruptive
sampling. However, this sampler is typically limited to soft sediments. The Van Veen sampler, or
the modified Van-Veen (Ted Young), is used in several national and regional estuarine monitoring
programs, including the NOAA National Status and Trends Program, the EPA Environmental
Monitoring and Assessment Program (EMAP), and the EPA National Estuary Program because it can
sample most types of sediment, is less subject to blockage and loss of sample than the Peterson or
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Examples of grab samplers
Photo courtesy of Allen Burton
w**~* ttes£™~ ^" '* £ »—<»~" *
"**— i, ,-',"•":•: •:> : -',
Photo courtesy of Allen Burton
Ponar
Eckman grab
Photo courtesy of Ed Long
Photo courtesy of Scott Carr
Double VanVeen grab
Birge-Eckman grab
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Photos on this page, courtesy ot Ed Long
Taking subsamples from a Ted Young
modified VanVeen sampler
Sampling using a Ted-Young modified
VanVeen. Large grab samplers such as
these require winches and sufficient boat
size for efficient operation.
Ted-Young VanVeen sampler in supporting
frame. Illustrating movable cover flap to enable
direct sampling from the grab sampler. Note the
overlying water in the sampler and adequate
volume, indicating an acceptable grab sample.
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Ponar samplers, is less susceptible to forming a bow wave during descent, and provides generally
high sample integrity (Klemm et al., 1990). The support frame further enhances the versatility of the
VanVeen sampler by allowing the addition of either weights (to increase penetration in compact
sediments) or pads (to provide added bearing support in extremely soft sediments). However, this
sampler is relatively heavy and requires a power winch to operate safely (GLNPO, 1994).
As shown in Appendix Table E-l, grab sampler capacities range from approximately 0.5 L to 75 L.
If a sampler does not have sufficient capacity to meet the study plan requirements, additional samples
can be collected and composited to obtain the requisite sample size (see Section 5.3). Grab samplers
penetrate to different depths depending on their size, weight, and the bottom substrate. Heavy, large
volume samplers such as the Smith-Mclntyre, large Birge-Ekman, Van Veen, and Petersen devices
can effectively sample to a depth of 30 cm. These samplers might actually sample sediments that are
too deep for certain study objectives (i.e., not reflective of recently deposited sediments). Smaller
samplers such as the small Birge-Ekman, standard and petite Ponar, and standard Shipek devices can
effectively collect sediments to a maximum depth of 10 cm. The mini-Shipek can sample to a depth
of 3 cm.
Another consideration in choosing a grab sampler is how well it protects the sample from disturbance
and washout. Grab samples are prone to washout which results in the loss of surficial, fine grained
sediments that are often important from a biological and contaminant standpoint. The Ponar, Ted-
Young modified grab, and Van Veen samplers are equipped with mesh screens and rubber flaps to
cover the jaws. This design allows water to pass through the samplers during descent, reducing
disturbance from bow waves at the sediment-water interface. The rubber flaps also serve to protect
the sediment sample from washout during ascent.
The use of small or lightweight samplers, such as the small Birge-Ekman (see page 3-6), petite Ponar,
and mini-Shipek, can be advantageous because of easy handling, particularly from a small vessel
and/or using only a hand line. However, these samplers are not recommended for use in strong
currents or high waves. This is particularly true for the Birge-Ekman sampler, which requires
relatively calm conditions for proper performance. Lightweight samplers generally have the
disadvantage of being less stable during sediment penetration. They tend to fall to one side due to
inadequate or incomplete penetration, resulting in unacceptable samples.
In certain very shallow water applications, such as a stream assessment at a superfund site, it might
be difficult to use even a lightweight sampler to collect a sample. In these cases, it might be
acceptable to collect sediment from depositional areas, using a shovel or other hand implement.
However, such sampling procedures are discouraged as a general rule and the use of a hand corer or
similar device is preferred (see Section 3.2.2).
Figure 3-2 summarizes appropriate grab samplers based on two important site factors, depth and
sediment particle size. This figure also indicates appropriate grab samplers depending on certain
common study constraints such as sample depth and volume desired, and the ability to subsample
directly from the sampler (see also Section 4.3; ASTM, 2000c). Based on all of these factors, the
Ponar or Van Veen samplers are perhaps the most versatile of the grab samplers, hence their common
usage in sediment studies.
Careful use of grab samplers is required to avoid problems such as loss of fine-grained surface
sediments from the bow wave during descent, mixing of sediment layers upon impact, lack of
sediment penetration, and loss of sediment from tilting or washout upon ascent (ASTM, 2000a;
Environment Canada, 1994; Baudo, 1990; Golterman et al., 1983; Plumb,1981). When deploying a
grab sampler, the speed of descent should be controlled, with no "free fall" allowed. In deep waters,
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use of a winching system is recommended to control both the rate of descent and ascent. A ball-
bearing swivel should be used to attach the grab sampler to the cable to minimize twisting during
descent. After the sample is collected, the sampling device should be lifted slowly off the bottom,
then steadily raised to the surface at a speed of about 30 cm/s (Environment Canada, 1994).
Recommendation Box #1
What are appropriate sampling devices given different study
objectives?
Grab or core samplers are preferred over dredges for collecting surficial sediments for
physicochemical or toxicity analyses. Dredges might be acceptable for collecting
macroinvertebrates.
Grab samplers are recommended for surficial sediment analyses where accurate
resolution of surficial sediment depths is not necessary. Core samplers are
recommended for: (a) assessments requiring accurate surficial sediment depth
resolution, (b) historical sediment analyses, (c) detailed sediment quality studies of
vertical sediment profiles, to characterize sediment quality at depth, (d) when
characterizing thick sediment deposits (such as shoals to be excavated), and/or (e)
where it is important to maintain an oxygen-free environment.
In sand, gravel, firm clay, or till sediments, grab samplers might be preferred over core
samplers (when only surface material needs to be collected and samples at depth are
not necessary) because the latter are often less efficient in these sediment types.
Ponar, VanVeen, or Ekman samplers are commonly used and generally preferred for
grab sampling. Ekman samplers, however, are less efficient in deep waters.
The Kajak-Brinkhurst corer is a common core sampler for soft, fine grained sediments
where large volumes or deep cores are not needed. The Phleger corer is commonly
used for a variety of sediments including peat and plant roots but is not appropriate
where large volumes or deep cores are needed.
Box corers are especially recommended for: (a) studies of the sediment-water interface;
(b) collecting larger volumes of sediment from a given depth (generally less than one
meter depth, however); (c) for in-situ studies involving interstitial water characterization;
and (d) collecting subsamples for different analyses from the same station.
Vibracorers are recommended for studies requiring deep cores (> 1 m), or where
sediment consists of very compacted or large grained material (e.g., gravel).
3.2.2 Core Samplers
Core samplers (corers) are used: (1) to obtain sediment samples for geological characterizations and
dating, (2) to investigate the historical input of contaminants to aquatic systems and, (3) to
characterize the depth of contamination at a site. Corers are an essential tool in sediments in which
3-dimensional maps of sediment contamination are necessary. Appendix Table E-2 discusses some
of the advantages and disadvantages of common corers.
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Photos on this page, courtesy of Allen Burton
Vibracorer in use showing extrusion
of the core sample for inspection and
Subsampling.
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Core devices are recommended for projects in which it is critical to maintain the integrity of the
sediment profile, because they are considered to be less disruptive than dredge or grab samplers.
Core samplers should also be used where it is important to maintain an oxygen-free environment
because they limit oxygen exchange with the air more effectively than grab samplers. Cores should
also be used where thick sediment deposits must be representatively sampled (e.g., for dredging
projects).
One limitation of core samplers is that the volume of any given depth horizon within the profile
sample is relatively small. Thus, depending on the number and type of analyses needed, repetitive
sampling at a site might be required to obtain the desired quantity of material from a given depth.
Some core samplers are prone to "plugging" or "rodding" where the friction of the sediment within
the core tube prevents it from passing freely and the core sample is compressed or does not sample to
the depth required. This limitation is more likely with smaller diameter core tubes and heavy clay
sediments. Except for piston corers and vibracorers, there are few core devices that function
efficiently in substrates with significant proportions of sand, gravel, clay, or till.
Coring devices are available in various designs, lengths, and diameters (see Appendix E). With the
obvious exception of hand corers, there are only a few corers that can be operated without a
mechanical winch. The more common of these include the standard Kajak-Brinkhurst corer,
suitable for sampling soft, fine-grained sediments, and the Phleger corer, suitable for a wider variety
of sediment types ranging from soft to sandy, semi-compacted material, as well as peat and plant
roots in shallow lakes or marshes (Mudroch and Azcue, 1995). The Kajak-Brinkhurst corer uses a
larger core tube, and therefore recovers a
greater quantity of sediment, than the
Phleger corer. Both corers can be used
with different liner materials including
stainless steel and PVC. Stainless steel
liners should not be used if trace metal
contamination is an issue.
Gravity corers are appropriate for
recovering up to 3 m long cores from
soft, fine-grained sediments. Recent
models include stabilizing fins on the
upper part of the corer to promote vertical
penetration into the sediment, and
weights that can be mounted externally to
enhance penetration (Mudroch and
Azcue, 1995). A variety of liner
materials are available including stainless
steel; Lexan®, and PVC. For studies in
which metals are a concern, stainless
steel liners should not be used.
Vibracorers are perhaps the most
commonly used coring device in
sampling programs in the U.S. because
they collect deep cores in most types of
sediments, yielding excellent sample
integrity. Vibracorers are one of the only
sampling devices that can reliably collect
Checklist
Corers may consist of the
following components (from
Mudroch and Azcue, 1995)
A hollow metal (or plastic) pipe that serves
as the core barrel
Easily removed plastic liners or core tubes
that fit into the core barrel and retain the
sediment sample
A valve or piston mounted on top of the core
barrel that is open and allows water to flow
through the barrel during descent, but shuts
upon penetration of the corer into the
sediment to prevent the sediment from
sliding through the corer during the ascent
A core catcher to retain the sediment
sample
A core cutter for penetration of the sediment
Removable metal weights (usually lead
coated with plastic) or piston-driven impact
or vibration to increase penetration of the
corer into the sediment
Stabilizing fins to ensure vertical descent of
the corer
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thick sediment samples (up to 10 meters or more). Some programs that rely on vibracorers include
the Puget Sound Estuary Program, the Great Lakes ARCS Program, and the Dredged Materials
Management Program.
Vibracorers have an electric-powered, mechanical vibrator located at the head end of the corer which
applies thousands of vertical vibrations per minute to help penetrate the sediment. A core tube and
rigid liner (preferably of relatively inert material such as cellulose acetate butyrate) of varying
diameter depending on the specific vibrator head used, is inserted into the head and the entire
assembly is lowered in the water. Depending on the horsepower of the vibrating head and its weight,
a vibracorer can penetrate very compact sediments and collect cores up to 6 m long. For example,
the ARCS program in the Great Lakes uses a Rossfelder® Model P-4 Vibracorer (Rossfelder
Corporation, La Jolla, CA) that produces a force of 7,000 Ibs and a mono-directional frequency of
3,400 vibrations per minute (GLNPO, 1994). Cores up to 6 m in length have been routinely collected
using this vibracorer. However, this particular model is relatively heavy (113 kg as compared to 8.1
kg for the more portable Wacker® Model M3000 vibracorers [GLNPO, 1994]). Therefore, use of a
heavy vibracorer requires a large vessel to maintain balance and provide adequate lift to break the
corer out of the sediment and retrieve it (GLNPO, 1994; PSEP, 1997a).
When deployed properly, box corers can obtain undisturbed sediment samples of excellent quality.
The basic box corer consists of a stainless steel box equipped with a frame to add stability and
facilitate vertical penetration on low slopes. Box corers are recommended particularly for studies of
the sediment-water interface or when there is a need to collect larger volumes of sediment from the
depth profile. Because of the heavy weight and large size of almost all box corers, they can be
operated only from a vessel with a large lifting capacity and sufficient deck space. Sediment inside a
box corer can be subsampled by inserting narrow core tubes into the sediment. Thus, they are an
ideal sampler for obtaining acceptable subsamples for different analyses at a given station. Carlton
and Wetzel (1985) describe a box corer that permits the sediment and overlying water to be held
intact as a laboratory microcosm under either the original in situ conditions or other laboratory
controlled conditions. A box corer was developed that enables horizontal subsampling of the entire
sediment volume recovered by the device (Mudroch and Azcue, 1995).
Figure 3-3 summarizes the core samplers that are appropriate given site factors such as depth and
particle size and other study constraints such as sample depth and volume required, and lifting
capacity needed to use the sampling device. Given the factors examined for general monitoring
studies, the Phleger, Alpine, and Kajak-Brinkhurst corers might be most versatile. For dredged
materials evaluations, and projects requiring sediment profile characterizations > 3 m in sediment
depth, the vibracorer or piston corer are the samplers of choice.
Collection of core samples with hand-coring devices should be executed with care to minimize
disturbance and/or compression of sediment during collection. To minimize disruption of the
sediment, core samples should be kept as stationary and vibration-free as possible during transport.
These cautions are particularly applicable to cores collected by divers.
The speed of descent of coring devices should be controlled, especially during the initial penetration
of the sediment, to avoid disturbance of the surface and to minimize compression due to frictional
drag from the sides of the core liner (ASTM, 2000d). In deep waters, winches should be used where
necessary to minimize twisting and tilting and to control the rate of both descent and ascent. With
the exception of piston corers or vibracorers, that are equipped with their own mechanical impact
features, for other corers, only the weight or piston mechanism of the sampler should be used to force
it into the sediment. The sampler should be raised to the surface at a steady rate, similar to that
described for grab samplers. Where core caps are required, it is essential to quickly and securely cap
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the core samples when the samples are retrieved. The liner from the core sampler should be carefully
removed and kept in a stable position until the samples are processed (see Chapter 4). If there is little
to no overlying water in the tube and the sediments are relatively consolidated, it is not necessary to
keep the core sample tubes vertical. Core sample tubes should be quickly capped and taped to secure
the sample. If sediment oxidation is a concern (e.g., due to potential changes in metal bioavailability
or volatile substances), then the head space of the core tube should be purged with an inert gas such
as nitrogen or argon.
Recommendation Box #2
How should sampling devices be used?
The recommended depth of sediment sampling is dependent on the study objectives.
Issues that determine the appropriate depth of sampling include: regulatory objectives
(e.g., depth of dredging for sediment remediation), need to characterize sediments at
depth (e.g., materials to be dredged versus shallow depositional areas in some
superfund sites), historical comparisons, sediment deposition rates, and/or time period of
contamination.
Appropriate winching systems are required to control the rate of ascent and descent of
the samplers.
The sampler should be rinsed thoroughly with water at the sampling station between and
within-station samples, and rinsed with water from the next sapling station before
collecting a sample. Equipment used in the handling of sediment should also be washed
thoroughly between samples.
More rigorous equipment decontamination might be necessary if highly contaminated
sites are sampled or if low level contaminants are a concern.
To reduce the probability of cross-contamination of samples, it is useful to sample
reference or relatively clean sites first and then suspected contaminated sites.
3.3 Sample Acceptability
Only sediments that are correctly collected with grab or core sampling devices should be used for
subsequent physicochemical, biological or toxicity testing. Acceptability of grabs can be ascertained
by noting that the samplers were closed when retrieved, are relatively full of sediment (but not over-
filled), and do not appear to have lost surficial fines. Core samples are acceptable if the core was
inserted vertically in the sediment and an adequate depth was sampled.
A sediment sample should be inspected as soon as it is secured. If a collected sample fails to meet
any of the acceptability conditions listed below for the respective sampling device, then the sample
might need to be rejected and another sample collected at the site. The location of consecutive
attempts should be as close to the original attempt as possible and located in the "upstream" direction
of any existing current. Rejected sediment samples should be discarded in a manner that will not
affect subsequent samples at that station or other possible sampling stations. Illustrations of
acceptable and unacceptable grab samples are provided in Figure 3-4.
Chapter 3: Collection of Whole Sediments
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Checklist
Sample acceptability for grab and core samples
Grab samples should be visually inspected to ensure that the following acceptability conditions
are satisfied (USEPA, 1986b; Environment Canada, 1994):
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Some specific programs (like Superfund) have their own decontamination procedures, which are
legally required.
Acceptable if Minimum
Penetration Requirement Met
and Overlying Water is Present
Unacceptable
(Washed, Rock Caught in Jaws)
Unacceptable (Canted with
Partial Sample)
Unacceptable
(Washed)
Figure 3-4. Illustrations of acceptable and unacceptable grab samples.
3.5 Field Measurements and Observations
Field measurements and observations are critical to any sediment collection study, and specific
details concerning sample documentation should be included in the study plan. Section 2.7
summarizes the types of information commonly recorded in the field during sampling. Several
programs, referenced in this Manual, provide specific guidance on field measurements and
observations necessary (e.g., see the Sampling and Analyses Plan Guidance development by the
Washington Department of Ecology in Appendix B [WDE, 1995 or PSEP, 1997a]). Measurements
and observations should be documented clearly in a bound field logbook (or on pre-printed sample
forms). Preferably, a logbook should be dedicated to an individual project. The investigator's name,
project name, project number, and book number (if more than one is required) should be entered on
Chapter 3: Collection of Whole Sediments
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the inside of the front cover of the
logbook. All entries should be written
in indelible ink, and the date and time of
entry recorded. Additionally, each page
should be initialed and dated by the
investigator. At the end of each day's
activity, or entry of a particular event if
appropriate, the investigator should
enter his or her initials. All aspects of
sample collection and handling as well
as visual observations and field
conditions should be documented in the
field logbooks at the time of sample
collection. Logbook entries should also
include any circumstances that
potentially affected sampling procedures and/or any field preparation of samples. Data entries
should be thorough enough to allow station relocation and sample tracking. Since field records are
the basis for later written reports, language should be objective, factual, and free of personal opinions
or other terminology which might appear inappropriate. In describing characteristics of samples
collected (see below), some cautions should be noted. First, polarized glasses are often worn in the
field to reduce glare, however, they can also alter color vision. Therefore, visual examination or
Checklist
Depending on program
objectives, field measurements
might include the following:
/ temperature and perhaps pH of the sediment
at the sediment-water interface;
/ concentration of dissolved oxygen in the water
overlying the sediment in the system being
sampled.
/ salinity or conductivity of the overlying water.
Recommendation Box #3
What information should be documented for each sample
collected? (PSEP, 1997a; ASTM, 2000a)
+ project title, time and date of collection, sample number, replicate number, site
identification (e.g., name); station number and location (e.g., positioning information);
^ water depth and the sampling penetration depth;
^ details pertaining to unusual events which might have occurred during the operation of the
sampler (e.g., possible sample contamination, equipment failure, unusual appearance of
sediment integrity, control of vertical descent of the sampler, etc.), preservation and
storage method, analysis or test to be preformed;
^ estimate of quantity of sediment recovered by a grab sampler, or length and appearance
of recovered cores;
^ description of the sediment including texture and consistency, color, presence of biota or
debris, presence of oily sheen, changes in sediment characteristics with depth, and
presence/location/thickness of the redox potential discontinuity (RPD) layer (a visual
indication of black is often adequate for documenting anoxia);
^ photograph of the sample is desirable, especially longitudinally-sectioned cores, to
document stratification;
^ deviations from approved work plans or SOPs.
NOTE: Some geological characterization methods might include an odor evaluation of the
sediment as this can provide useful information on physicochemical conditions. However,
sediment odor evaluation is potentially dangerous depending on the chemicals present in the
sediment (ASTM 2000a) and should therefore be done cautiously, if at all.
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characterization of samples should be performed without sunglasses (GLNPO, 1994). Second,
descriptions of sediment texture and composition should rely on a texture-by-feel or "ribbon" test in
addition to visual determinations (GLNPO, 1994). In this test, a small piece of suspected clay is
rolled between the fingers while wearing protective gloves. If the piece easily rolls into a ribbon it is
clay; if it breaks apart, it is silt (GLNPO, 1994).
3.6 Documentation of Sample Collection
Documentation of collection and analysis of sediment and porewater samples requires all the
information necessary to: 1) trace a sample from the field to the final result of analysis; 2) describe
the sampling and analytical methodology; and 3) describe the QA/QC program (Mudroch and Azcue
1995; Keith, 1993). Poor or incomplete documentation of sample collection can compromise the
integrity of the sample(s) and thus, the study. In addition, stations that could not, or were not,
sampled should be documented with an explanation. Samples should be accompanied by chain-of-
custody forms that identify each sample collected and the analyses to be conducted on that sample.
Specific guidance on quality assurance procedures regarding sample chain-of-custody is summarized
in Chapter 7.
Checklist
Project documentation should include...
/ type of vessel used (e.g., size, power, type of engine);
/ notation of the system used to define the position of the sampling site;
/ notation of the system used to identify and track samples;
/ name of personnel collecting the samples;
/ level of personal protective equipment worn;
/ notation of any visitors to the site;
/ sketch of sampling area with photographs, if possible;
/ ambient weather conditions, including wind speed and direction, wave action, current,
tide, vessel traffic, temperature of both the air and water, thickness of ice if present;
/ type of sediment collection device and any modifications made during sampling;
/ calibration data.
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Field Sample Processing, Transport,
and Storage of Sediments
The way in which sediment samples are processed, transported, and stored might alter contaminant
bioavailability and concentration by introducing contaminants to the sample or by changing the
physical, chemical, or biological characteristics of the sample. Manipulation processes often change
availability of organic compounds because of disruption of the equilibrium with organic carbon in
the pore water/sediment system. Similarly, oxidation of anaerobic sediments increases the
availability of certain metals (DiToro et al., 1990; Ankley et al., 1996). Materials and techniques
should be selected to minimize sources of contamination and variation, and sample treatment prior to
testing should be as consistent as possible.
A flowchart is presented in Figure 4-1 that summarizes common sediment processing procedures
discussed in this section as well as issues and objectives relevant to each processing step.
4.1 Sample Containers
Any material that is in contact with a field sample has the potential to contaminate the sample or
adsorb components from the sample. For example, samples can be contaminated by zinc from
glassware, metals from metallic containers, and organic compounds from rubber or plastic materials.
The use of appropriate materials, along with appropriate cleaning procedures, can minimize or
mitigate interferences from sample containers.
Recommendation Box #1
Sample Containers
High density polyethylene (HOPE) or polytetrafluoroethylene (PTFE) containers are
suitable and preferred for most analytical measurements because they are made of
relatively inert material and they are generally unbreakable.
All containers should be pre-cleaned prior to filling with sample.
Consider using certified, pre-cleaned containers, commercially available from many
vendors.
Purge containers with inert gas (e.g., nitrogen) prior to and after filling if anoxic conditions
must be maintained.
Fill containers completely if the sample will not be frozen prior to analysis.
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Issues to
Consider
Check sample
location frequently
to ensure same site
Need adequate
clean space,
utensils, etc.
May need to
perform these
operations in a
glovebox if metals
or volatiles are a
concern; U.V.
shielding may be
needed if PAHs a
concern
May need to
perform these
operations in a
glovebox if metals
or volatiles are a
concern; U.V.
shielding may be
needed if PAHs a
concern
> 1 sampler volume n
per site for analyses?
or
"average" condition to be
measured for each station
within a site?
Composite multiple
samples from a site
Subsampling required
for different types of
analyses?
Sufficient space,
facilities to composite
infield?
Sufficient space,
facilities for
subsampling in
field?
mbine similar sampl
(e.g., similar depth
horizons from cores)
Mix/homogenize
sample
Mix/homogenize
composite sample
Subsample for
separate types of
analyses
Subsample for
separate types of
analyses
Store in cold (1-4° C),
dark and transport to the
lab
Store in cold (1-4°C),
dark and transport to lab
Store in cold (1-4°C),
dark and transport to lab
Composite samples in lab,
homogenize, and subsample
for different types of analyses
Figure 4-1. Flowchart of suggested sediment processing procedures
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4.1.1 Container Material
Borosilicate Glass, and high-density polyethylene, polycarbonate and fluorocarbon plastics should be
used whenever possible to minimize leaching, dissolution, and sorption (ASTM, 2000a; APHA,
1995). Direct contact between sediment samples and the following substances should be avoided:
PVC, natural or neoprene rubber, nylon, talcum powder, polystyrene, galvanized metal, brass,
copper, lead, other metal materials, soda glass, paper tissues, and painted surfaces. Table 4-1
summarizes the appropriate types of sampling containers and allowable holding times for various
types of contaminants associated with sediments.
In general, sediments and pore waters with multiple or unknown chemical types should be stored in
containers made from high density polyethylene plastic or polytetrafluoroethylene (PTFE or
Teflon®) as these materials are least likely to add chemical artifacts or interferences and they are
much less fragile than glass. Samples for organic contaminant analysis should be stored in brown
borosilicate glass containers with PTFE lid liners. If volatile compounds will be analyzed, containers
should have a septum to minimize escape of volatile gases during storage and analysis. Extra
containers should be provided for these analyses in the event that re-analysis of the sample is
required. If samples are contaminated with photoreactive compounds such as PAHs, exposure to
light should be minimized by using brown glass containers or clear containers wrapped tightly with
an opaque material (e.g., clean aluminum foil). Plastic or acid-rinsed glass containers are
recommended when the chemicals of concern are heavy metals.
4.1.2 Container Preparation
Many vendors have commercially available pre-cleaned containers for a variety of applications. For
chemical and toxicological analyses, certified pre-cleaned containers are often a cost-effective way to
limit the potential for container contamination of samples. Thus, manufacturer-supplied pre-cleaned
containers are often a prerequisite in QAPPs.
If new containers are used, Environment Canada (1994) recommends that new glassware and
plasticware should be soaked in 1:1 concentrated acid prior to use. Soaking overnight is adequate for
glassware. For plasticware, the recommended procedure involves soaking for seven days in
hydrochloric acid (HC1), followed by seven days in nitric acid (HNO3), followed by seven days in
deionized water. Shorter soaking times might be satisfactory in most instances (ASTM, 2000a).
Used sample containers should be washed following these steps: (1) non-phosphate detergent wash,
(2) triple water rinse, (3) water-miscible organic solvent wash (acetone followed by pesticide-grade
hexane), (4) water rinse, (5) acid wash (such as 5% concentrated HC1) and (6) triple rinse with
deionized-distilled water. A dichromate-sulfuric acid cleaning solution can generally be used in
place of both the organic solvent and the acid (Steps 3 through 5), but it might attack any silicone
adhesive present in the container. See ASTM (2000a) and USEPA (2000d) for further information.
If a sample is to be refrigerated, the container should be filled to the brim to reduce oxygen exposure.
This is particularly critical for volatile compounds (e.g., AVS). If a sample is to be frozen, the
container should be filled to approximately 90% of its volume (i.e., 10% headspace) to allow for
expansion of the sample during freezing. See Section 4.4 for preservation and storage conditions for
various types of analyses. For studies in which it is critical to maintain the collected sediment under
anoxic conditions (e.g., where metals are the pollutants of concern), the container should be purged
with an inert gas (e.g., nitrogen) before filling and then again before capping tightly.
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Table 4-1. Recommended sampling containers, holding times, and storage
types of sediment analyses (USEPA, 1983;1993; ASTM, 2000a). P=Plastic;
PTFE=Polytetrafluoroethylene; R=refrigerate; F=freeze
conditions for common
G=Glass;
Contaminant
Ammonia
Sulfate
Sulfide
Oil and Grease
Mercury
Metals (except Cr or Hg)
Extractable organics (including
phthalates, airosamines,
organochlorine pesticides, PCBs
aromatics, isophorone, PAHs,
haloethers, chlorinated
hydrocarbons, and TCDD)
Purgables (halocarbons and
aromatics)
Pesticides
Sediment Toxicity (acute and
chronic)
Bioaccumulation testing
Container
P,G
P,G
P,G
G
P,G
P,G
G, PTFE-lined
cap
G, PTFE-lined
septum
G, PTFE-lined
cap
P, PTFE
P, PTFE
Holding Time
28 days
28 days
28 days
28 days
6 weeks
6 months
7 days (until
extraction)
30 days (after
extraction)
14 days
7 days (until
extraction) 30
days (after
extraction)
2 weeks*
2 weeks*
Storage Condition
R;F
R;F
R or NaOH; pH>9
HC1, pH<2
H2SO4, pH<2; R
HNO3, pH<2; F
R;F
R;F
R;F
R, dark
R, dark
*Holding time might be longer depending on the magnitude and type of contaminants present.
See Section 4.5.
All sediment containers should be properly labeled with a waterproof marker prior to sampling.
Containers should be labeled on their sides in addition to or instead of labeling the lids. Each label
should include, at a minimum, the study title, station location and/or sample identification, date and
time of collection, sample type, and name of collector. Blind sample labeling (i.e., a sample code)
should be used, along with a sample log that identifies information about each sample (see Section
2.7) to minimize potential analytical bias. Additional information such as required analyses and any
preservative used might also be included on the label although this information is typically recorded
on the chain-of-custody form (see Section 2.7 and 7.6). Labeled containers should be stabilized in an
upright position in the transport or storage container (see Section 4.4 Transport and Storage for
further information). Extra containers should be carried on each sampling trip.
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4.2 Subsampling and Compositing Samples
The decision to subsample and/or composite sediment samples within or among stations depends on
the purpose and objectives of the study, the nature and heterogeneity of the sediments, the volume of
sediment required for analytical and/or toxicity assessment, and the degree of statistical resolution
that is acceptable. Subsampling and compositing might be accomplished in the field, if facilities,
space, and equipment are available, or alternatively, in a laboratory setting following sample
transport.
Recommendation Box #2
How should sediment samples be subsampled and composited?
Overlying water should be siphoned off, not decanted, from grab samplers prior to
subsampling.
All utensils that are used to process samples should be made of inert materials such as
Teflon(Ehi gh quality stainless steel, or HOPE.
Subsamples should be collected away from the sides of the sampler to avoid potential
contamination.
Sediment samples should be processed prior to long-term storage, within 72 hours (and
preferably within 24 hours) of collection.
Sufficient sample homogenization, prior to placing in containers, is critical for accurate
measurements and correct sediment quality determinations.
If rigorous evaluation of metal contamination is a focus of the study, or if anaerobic
conditions need to be maintained for other reasons, it might be necessary to homogenize,
subsample, and composite samples in an oxygen-free glovebox or other suitable
apparatus.
Similar depth horizons or geologic strata should be subsampled when compositing core
samples.
4.2.1 General Procedures
Subsampling is useful for collecting sediment from a specific depth of a core sample, for splitting
samples among multiple laboratories, for obtaining replicates within a sample, or for forming a
composite sample.
Compositing refers to combining aliquots from two or more samples and analyzing the resulting
pooled sample (Keith, 1993). Compositing is often necessary when a relatively large amount of
sediment must be obtained at each sampling site (for instance, to conduct several different physical,
chemical or biological analyses). Compositing might be a practical, cost-effective way to obtain
average sediment characteristics for a particular site (see Table 2-2), but not to dilute a polluted
sample. Also, if an objective of the study is to define or model physicochemical characteristics of
the sediment, it might be important not to composite samples because of model input requirements
(EPRI, 1999).
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
All utensils (e.g., spoons, scoops, spatulas) which come in direct contact with sediment samples
during handling and processing should be made of non-contaminating materials (e.g., glass, high-
quality stainless steel and/or Teflon®).
Considerations
All handling procedures carry the risk of sample contamination.
Therefore, sediment sample handling should be kept to a
minimum. Potential sample contamination can be caused by
the following common situations...
! making field measurements of sediments using contaminated probes, utensils, or other
instruments.
! contaminated and uncontaminated stations are sampled without appropriate
decontamination of equipment between stations.
! the parameter of interest is volatile (e.g., ammonia, acid volatile sulfides, or volatile organics)
and samples are exposed to air.
! samples are exposed to vessel exhaust fumes, lubricants, or rust.
4.2.2 Grab Samples
If a sediment grab sample is to be subsampled in
the laboratory, the sample should be released
carefully and directly into a labeled container that
is the same shape as the sampler and made of a
chemically-inert material (see Section 4.1 for
recommendations on containers). The container
must be large enough to accommodate the
sediment sample and should be tightly sealed
with the air excluded.
If the grab sample is to be subsampled in the
field, it is desirable to subsample from the
sampler directly to minimize sediment handling
and associated artifacts. Therefore, the sampler
should allow access to the surface of the sample
without loss of water or fine-grained sediment
(see Section 3.1.1 for sampler descriptions). This
typically dictates the use of a grab sampler with
bucket covers that are either removable or hinged
to allow access to the surface of the sediment
sample (e.g., Ponar, Van Veen).
Prior to subsampling from the grab sampler, the
overlying water should be removed by slow
siphoning using a clean tube near one side of the
sampler (WDE, 1995;PSEP, 1997a). If the
overlying water in a sediment sampler is turbid, it
should be allowed to settle if possible.
Considerations
When working with
grab samples...
! decanting the water, or opening the
jaws lightly to let the water run out is
not recommended as these methods
might result in unacceptable
disturbance or loss of fine-grained
sediment and organic matter.
! if metal contamination or sediment
oxygen demand are of concern,
oxidation of sediments could
significantly alter their characteristics.
Process the sample in a glovebox or
similar apparatus under an oxygen-free
environment.
! for samples that are suspected of
heavily elevated polynuclear aromatic
hydrocarbons (PAHs), process
immediately under low light upon
retrieval to minimize ultraviolet light-
activated toxicity of PAHs (Ankley et al.,
1994).
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Checklist
Compositing samples
involves
The general subsampling and compositing
process for grab samples is illustrated in Figure
4-2. Subsampling can be performed using a
spoon or scoop made of inert, non-
contaminating material. Sediment which is in
direct contact with the sides of the grab
sampler should be excluded as a general
precaution against potential contamination
from the device. Subsamples may be combined
or placed into separate clean, pre-labeled
containers. If the sample is to be frozen, it is
advisable to leave approximately 10% head
space in the container to accommodate
expansion and avoid breakage.
There are two alternatives for compositing
sediment samples from grab samplers (see
Figure 4-2): (1) compositing and homogenizing
(mixing) in the field and (2) compositing in the
field and homogenizing in the laboratory.
In some studies (e.g., where metals are the
pollutants of concern), it might be necessary to
subsample a grab sample under oxygen-free
conditions to minimize oxidative changes. In
these cases, it is recommended that a hand-
coring device be used for subsampling. The
core should be inserted immediately upon
retrieval of the sampler, then removed and
placed into a glove box or bag which is flushed
with a constant, controlled volume of inert gas.
The sediment within the core can then be extruded under oxygen-free conditions into deaerated
containers. The presence of oxygen during handling and storage might be relatively unimportant
(Brumbaugh et al., 1994) or very important (Besser et al., 1995), depending on the sediment
characteristics, the contaminants of concern, and the study objectives.
4.2.3 Core Samples
Subsampling sediment core samples is usually done to focus the assessment on a particular sediment
horizon or horizons and/or to evaluate historical changes or vertical extent in contamination or
sedimentation rates. Whenever subsampling of retrieved sediment cores is required, particularly for
analysis of contaminants, the sediment should be extruded from the core liners and subsampled as
soon as possible after collection. This can be accomplished in the field if appropriate facilities and
equipment are available, or in the laboratory after transport.
Systematic subsampling (see Figure 4-3) involves removing the sediment from the core in sections of
uniform thickness. Each incremental core section corresponds to a particular sediment depth
interval. In remedial dredging and geological applications, longer sections (e.g., 25-50 cm) are
typically used to characterize a site.
In the field
/ placing subsamples from individual
grab samples in a clean container to
form a composite sample
/ transporting the composite to a
laboratory
/ homogenizing the sample at the
laboratory to prepare it for testing (See
Section 4.3 for further details)
In the lab
/ placing subsamples from multiple
grabs in a clean container
/ mixing the subsamples to form a
homogeneous composite sample
/ placing the composite sample in one
or more containers, depending on the
number of analyses to be performed
/ transporting the composite sample to
a laboratory (or laboratories) for
testing
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Sample
Retrieval
arid Inspection
Hinged
bucket cover
Surface of retrieved sediment sample
0-2 em layer
"" Grab sampler bucket
Subsampling
Surficial 0-2 cm layer (or other
layer of interest) is scooped or
spooned out of grab.
Scenario 1
Compositing and
homogcnization (mixing) in the
field,
Subsamples
from
multiple
grabs within
a station are
placed in a
bowl.
Subsamples
mixed to
create a
homogenous
composite
sample.
[
Portions of the composite sample are
placed in one or more containers
depending on the analyses to be
conducted. Containers transported to
one or more analytical laboratories.
Scenario 2
Compositing in the field,
homogenixation in the
laboratory.
Subsamples from
multiple grabs
within a station
are placed in a
single container
which is sealed
and transported to
the laboratory.
Composite
sample is
homogenized
w i thin the
laboratory
prior to
analvMs.
Scenario 3
No compositing.
Grab 1
I Sabsaraple 2 -
L.
Subsarnple from
individual grabs
are placed in
individual
* containers and
transported to the
laboratory.
Laboratory analyzes the
samples individually or may
combine them as appropriate to
create compositers.
Figure 4-2. Alternatives for subsampling and compositing sediment grab samples.
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Sample Retrieval
and Inspection
(core descriptions
should be recorded
and core should be
labeled)
Core liner with sediment
sample showing layered
stratigraphy (multiple
sediment units)
Core wilh only two
seclimeni units
Simple Subsatnpling
Suhsampling and Compositing (single core
sample)
L
Subsamples
taken
from uniform
depth
intervals,
placed in
separate
containers,
and analyzed
separately
Each distinct sediment unit is
subsampled, homogenized to create a
depth composite, and analyzed
separately.
Subscripting and
Compositing
(multiple core
samples)
Only horizons with
similar stratigraphy
should be combined to
create homogenous
composite samples for
analysis.
Figure 4-3. Alternatives for subsampling and compositing sediment core samples.
Chapter 4: Field Sampling Processing, Transport, and Storage of Sediments and Interstitial Water
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The depth horizon(s) sampled will depend on the study objectives as well as the nature of the
substrate. For toxicological studies, the biologically active layer and sedimentation rates at the site
might be important factors determining which core sections are sampled. In these studies,
subsampling depth intervals include the 0 to 2 cm layer (for recent deposition) and the 0 to 5 cm or 0
to 15 cm layers (for biological activity, depending on resident organisms). Many programs have
project-specific depths corresponding to study requirements, such as dredging depths for navigation
or remediation dredging. In many regional or national environmental monitoring programs (e.g.,
EMAP), the uppermost surficial layer is sampled because information on the horizontal distribution
of sediment contaminants is desired (USEPA, 2000d).
There are various methods for subsampling sediment cores including gradual extrusion, dissection of
a core using a jig saw, reciprocating saws, use of a segmented gravity corer, a hand corer, or scoops
and spoons. Cutting devices range from stainless steel knives to teflon or nylon string.
A piston-type extruder that applies upward pressure on the sediment is an instrument commonly used
to gradually expose a core for sectioning in some monitoring programs where specific sediment
depths have been defined a priori (Kemp et al., 1971). [Note: For dredged material studies and other
types of remediation projects, where pre-determined depth strata are not necessarily defined, it is
usually important to view the entire core prior to sectioning or compositing.] The capped core liner
containing the sediment and overlying water is uncapped at the lower end and placed vertically on
top of the piston. The top cap is removed and the water is siphoned off to avoid disturbance of the
sediment-water interface. The core liner is then pushed slowly down until the surface of the
sediment is at the upper end of the liner. Sediment sections are collected by pushing the liner down
and cutting the exposed sediment into sections of the desired thickness using a stainless steel or
Teflon® cutter (Environment Canada, 1994; Mudroch and Azcue, 1995). A 1- to 2-mm outer layer
of sediment that has been in contact with the plastic or metal liner should be removed and discarded,
if possible, to avoid contamination. Each sediment subsample should be placed into a labeled, clean
and chemically-inert container, or, if subsamples are being composited, into an appropriately sized
mixing bowl. The size of the container should be as close to the volume of the sediment as possible
to minimize the head space in the container. If it is desirable to maintain an oxygen-free environment
during subsampling, then all handling or manipulations should take place in a glove box or bag filled
with an inert gas and modified to accommodate the core liner through an opening (Environment
Canada, 1994; Mudroch and MacKnight, 1994).
Cores of more consolidated material can be mounted onto a horizontal U-shaped rail and the liner cut
using a saw mounted on a depth-controlling jig. The final cut can then be made with a sharp knife to
avoid contamination of the sediment by liner material, and the core itself can be sliced with Teflon®
or nylon string. The core then becomes two D-shaped halves that can be easily inspected and
subsampled (Mudroch and Azcue, 1995). Sediment in contact with the saw blade should not be used
for toxicity tests or metals analyses due to potential contamination from the saw blade. Another
alternative for sectioning and subsampling is a segmented gravity corer described by Aanderaa
Instruments of Victoria, BC, Canada. The core tube of the sampler consists of a series of rings
placed on top of one another. Subsampling is carried out by rotating the rings around its other axis
so that it cuts sediment layers of similar thickness. This segmented core tube is suitable for sampling
fine-grained sediments and allows one person in the field to subsample the core into 1-cm sections
(Mudroch and Azcue, 1995).
Sediment from box-core samples can be effectively subsampled with a small hand corer after the
overlying water has been carefully siphoned off and discarded. Hand corers with small inner
diameters less than 3 cm tend to compact sediments, so they must be used with care. Spoons or
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scoops have also been used to subsample surface sediments from a box corer (Environment Canada,
1994).
Like grab samples, core samples may be composited or subsampled in the field or laboratory after
evaluating them for acceptability. Although there might be occasions when it is desirable to
composite incremental core depths, it is recommended that only horizons of similar stratigraphy be
composited. Depending on the study objectives and desired sampling resolution, individual horizons
within a single core can be homogenized to create one or more "depth composites" for that core, or
corresponding horizons from two or more cores might be composited (Figure 4-3). Composite
samples must be homogenized prior to analysis or testing.
4.3 Homogenization
Homogenization refers to the complete mixing of sediment to obtain consistency of physicochemical
properties throughout the sample prior to using in analyses. Homogenization is typically performed
on individual samples, as well as on composited samples and can be done either in the field or the
laboratory.
4.3.1 General Procedures
Prior to homogenization, unrepresentative materials (e.g., twigs, shells, leaves, stones, wood chips
and seagrass) are often removed and documented in an appropriate field log (see Section 5.2 for
techniques to remove unrepresentative material). The need for removal of larger matter depends on
the analyses to be conducted.
Mixing should be performed as quickly and efficiently as possible, because prolonged mixing can
alter the particle-size distribution in a sample and cause oxidation of the sediments (Ditsworth et al.,
1990; Stemmer et al., 1990a;b). This can alter the bioavailability of contaminants, particularly
metals, by increasing or decreasing their availability (Ankley et al., 1996). If metal contaminants or
volatile chemicals are a concern, samples should be mixed in a glovebox under an inert atmosphere
and quickly partitioned into sample containers for analysis.
Recommendation Box #3
How should samples be homogenized?
Use a sufficiently large, precleaned glass or stainless steel mixing bowl to homogenize
the sample.
Use clean glass polyethylene, or stainless steel implements (e.g., spoon) to mix sediment.
Mixing should be performed as quickly and efficiently as possible while attempting to
reduce oxidation of the sample.
Intensive manual mixing of wet sediment, in a suitably large container, is usually sufficient
to homogenize the sample (Burton et al., 1989; Ingersoll and Nelson, 1990; Johns et al.,
1991 a; Carrand Chapman, 1992).
Regardless of the mixing method selected, the effectiveness of the method should be
demonstrated using a homogenate replicate.
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Photo courtesy of Nile Kemble
Homogenizing a composited sediment
sample using a mechanical mixer
Photo courtesy of Chris Ingersol
,%.. 1'J
Subsampling sediment for toxicity testing
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Mixing should be performed in a large,
precleaned glass or stainless steel bowl. The
sediment should be thoroughly stirred with a clean
glass, high density polyethylene, or stainless steel
spoon until textural, color, and moisture
homogeneity are achieved (Environment Canada,
1994; PSEP, 1995). Hand mixing has also been
performed by rolling the sediment out flat on a
sheet of plastic or pre-combusted foil and
tumbling the sediment by alternately raising each
corner of the sheet (Mudroch and Macknight,
1994). This procedure, however, is not
recommended where the anaerobic integrity of the
sediment must be maintained.
Considerations
Homogenization of
anaerobic sediments...
! Beware of over-mixing and/or
introducing air to the sample. Such
mixing is likely to change the chemical
characteristics of the sample and yield
unrepresentative results. This is
especially important if samples are
initially anaerobic or if volatile or labile
chemicals are of interest (e.g., AVS).
Mechanical mixers have also been used to homogenize samples (Ditsworth et al., 1990; Stemmer et
al., 1990b; Kemble et al., 1993), including portable cement mixers (bare metal and Teflon-lined) and
portable drills fitted with a variety of stainless steel paddles (Kemble et al., 1994b).
Homogenate replicates consist of two or more subsamples, taken from different locations within a
mixed sample, and then comparing analytical results of the replicate samples. After the sediment has
been homogenized, it is generally partitioned among sample containers. Partitioning sediments for
chemical or toxicity analyses may be accomplished using various methods. In one method, a number
of small portions are removed from random locations in the mixing container and distributed
randomly in all sample jars until the appropriate volume of sediment is contained in each sample jar
for each analysis. During distribution, the sediment is periodically mixed using a glass rod or
porcelain spatula to minimize stratification effects due to differential settling, especially if the
sediment is prone to rapid settling (ASTM, 2000a). An alternative is to use a splitter box designed to
contain and then divide the homogenized sediment.
4.4 Sample Transport and Storage
Transport and storage methods should be designed to maintain structural and chemical qualities of
sediment and pore water samples. Sediments collected using grab samplers are usually transferred
from the sampler to containers that may or may not serve as the storage container. The containers
might be stored temporarily in the field or they might be transported immediately to a laboratory for
storage. If sediment core samples are not sectioned or subsampled in the field, they may be stored
upright, in the core liner, for intact transportation to the laboratory. If sectioning or subsampling
takes place in the field, then the subsamples may also be transferred to sample containers and stored
temporarily. The sample containers with the field-collected sediments are then placed into a
transport container and shipped to the laboratory.
4.4.1 General Procedures
Proper storage conditions (see Table 4-1) should be achieved as quickly as possible after sampling.
For those parameters that are preserved via refrigeration (e.g., toxicity) samples should be stored in
the field in refrigerated units on board the sampling vessel or in insulated containers containing ice or
frozen ice packs. For samples that can be preserved via freezing (e.g., some metal and organic
chemical analyses) dry ice can be used to freeze samples for temporary storage and transport
(USEPA, 1983, 1993). Pelletized dry ice has been used effectively in the dredged materials
management program to store core samples. It is important to know chilling capacities and
Chapter 4: Field Sampling Processing, Transport, and Storage of Sediments and Interstitial Water
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
efficiencies to assure that temperature regulation is adequate. Care should be taken to prevent
refrigerated samples from freezing and to keep frozen samples from thawing. Freezing changes the
sediment volume depending on the water content, and it permanently changes the structure of the
sediment and potentially alters the bioavailability of sediment associated contaminants.
Logistics for sample transport will be specifically tailored to each study. In some cases it is most
efficient to transfer samples to a local storage facility where they can be either frozen or refrigerated.
Depending on the logistics of the operation, field personnel may transport samples to the laboratory
themselves or utilize an overnight courier service. If a freight carrier is employed, the user must be
aware of any potentially limiting regulations (e.g. regarding the use of ice or dry ice). Samples that
have a recommended storage temperature should be cooled to that temperature prior to placement in
the transport container. Light should be excluded from the transport container.
Core samples should be transported as intact core liners (tubes). Prior to sample transport, the entire
space over the sediment in the core liner should be filled with site water, and both ends of the core
liner should be completely sealed to prevent mixing of the sediment inside. The cores should be
maintained in an upright position particularly if the sample is not highly consolidated material, and
secured in either a transport container (e.g., cooler or insulated box) with ice or ice packs, or in a
refrigerated unit that can maintain a temperature near 4°C (Environment Canada, 1994). If the
transport container cannot accommodate long core samples such as from vibracorers or piston corers
(core liners > 1 m), then the core samples can be cut into 1-m lengths, and the ends securely capped
such that no air is trapped inside the liners (see Section 4.3.3).
Impregnating unconsolidated sediment cores with epoxy or polyester resins will preserve sediment
structure and texture (Ginsburg et al., 1966; Crevello et al., 1981) but not sediment chemical
characteristics. Therefore, this procedure is not recommended for transporting or storing sediment
samples for chemical characterization or biological testing (Environment Canada, 1994).
Recommendation Box #4
Sample Transport and Storage
The volume of overlying water in sediment samples should be minimized to reduce the
potential for resuspension of surface sediments during transport.
Care should be taken to retain the surficial floe overlying a core sample.
Cores should be secured in an upright position during transport to minimize disturbance
of the sediment.
Sediment samples collected in the field should be stored in containers without
headspace at 4° C and in the dark to minimize changes in contaminant bioavailability.
Prior to transport, headspace in the core liner should be filled with site water and both
ends of the liner should be completely sealed.
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4.5 Sample Holding
Times
Limits for effective holding times are
governed by sediment type and
contaminant characteristics (ASTM,
2000a). Because these qualities are
not always known, a general
recommendation is to store sediments
and interstitial water in the dark at 4
°C (SETAC, 2001). Preservation and
recommended storage times for
various types of analyses are
summarized in Table 4-1.
Recommendation Box #5
How long should samples be
stored before analysis?
Unless site-specific information is available,
sediment samples should be stored no longer
than two weeks prior to using in toxicity testing.
Preserved samples for chemical analyses
should be stored no longer than the maximum
holding times as defined by the particular
program.
Samples collected for toxicity tests should be used as quickly as possible. Recommended maximum
holding times range from 10 days (NOAA) to two weeks (ASTM, 2000a; USEPA, 2000d), to eight
weeks (USEPA/ACOE, 1991, 1998). Preferred sample storage times reported for toxicity tests have
varied substantially (Dillon et al., 1994; Becker and Ginn, 1990; Carr and Chapman, 1992; Moore et
al., 1996; Sarda and Burton, 1995; Sijm et al., 1997; Defoe and Ankley, 1998), and differences
appear to depend primarily upon the type or class of contaminant(s) present.
Extended storage of sediments that contain high concentrations of labile contaminants (e.g.,
ammonia, volatile organics) might lead to loss of these contaminants and a corresponding reduction
in toxicity. Under these circumstances, the sediment should be tested as soon as possible after
collection, but not later than two weeks (Sarda and Burton, 1995). Sediments that exhibit low to
moderate toxicity might exhibit higher variability in toxicity when tested following storage of short
duration (e.g. two weeks). Testing could actually be more reliable following longer storage for these
types of samples if the longer storage reduces potential interference associated with indigenous
predators (DeFoe and Ankley, 1998). Sediments contaminated with relatively stable compounds
(e.g. high molecular weight compounds such as PCBs) or those that exhibit moderate-to-high
toxicity, do not seem to vary appreciably in toxicity with increased storage time (Moore et al., 1996;
DeFoe and Ankley, 1998). Longer term storage might be acceptable in such cases. Given our
incomplete knowledge on the changes that occur, it is recommended that sediments should be stored
no longer than two weeks for toxicity testing unless site-specific information indicates otherwise.
Periodic measurements of contaminants of concern provide a useful context for interpretation of
toxicity test results when sediments or interstitial waters are stored for extended periods of time, but
this is rarely cost-effective. It might be more efficient to conduct interstitial water toxicity tests
within two weeks of sediment collection, corresponding with the start of sediment tests (Ingersoll et
al., 1993). In general, though, interstitial water should be analyzed as quickly as possible following
sampling to minimize possible changes in contaminant bioavailability.
Sediment cores collected for stratigraphical or geological studies can be stored at 4 °C in a humidity-
controlled room for several months without any substantial changes in sediment properties (Mudroch
and Azcue, 1995).
Chapter 4: Field Sampling Processing, Transport, and Storage of Sediments and Interstitial Water
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Sediment Manipulations
Manipulation of sediments in the laboratory is often required to achieve certain desired
characteristics or forms of material for toxicity testing and chemical analysis. As all manipulation
procedures alter some qualities of field samples, it is critical to evaluate the effect that these changes
might have on the study objective and on each critical measurement endpoint. Therefore, all
procedures used to prepare sediment samples should be explicitly described in the study plan and
fully documented. Generally, manipulation procedures should be designed to maintain sample
representativeness in terms of toxicity and chemistry by minimizing procedural artifacts. Under
certain programs, some analytical procedures and toxicity test protocols necessitate specific
manipulations (e.g., seawater or solvent extractions for effluent toxicity tests, USEPA/ACOE, 1991,
1998). The reader should always consult and follow any program or test-specific guidance.
This chapter discusses methods for several common manipulations performed in the laboratory
including sieving, spiking, organic carbon modification and formulated sediments, sediment dilution,
and elutriate preparation. Other sediment manipulations, such as salinity adjustments or pre-
treatment of sediment ammonia or sulfides (often done in conjunction with toxicity testing in certain
regulatory programs) are not discussed in this manual as these are well documented elsewhere (e.g.,
PSEP, 1995; USEPA/ACOE, 1998). The reader should consult these references for further
information on these procedures. Figure 5-1 presents a flowchart summarizing the laboratory
manipulations discussed in this section, illustrating important issues to be considered for each
manipulation.
5.1 Sieving
In general, sieving is not recommended
because it can substantially change the
physicochemical characteristics of the
sediment sample. For example, wet
sieving of sediment through fine mesh
(<500 |-im openings) has been shown to
result in decreased percent total organic
carbon and decreased concentrations of
total PCBs, which might have been
associated with fine suspended organic
matter lost during the sieving process
(Day et al., 1995). Sieving can also
disrupt the natural chemical equilibrium
by homogenizing or otherwise changing
the biological activity within the
sediment (Environment Canada, 1994).
In some cases, however, sieving might be
necessary to remove indigenous
organisms, which can interfere with
subsequent toxicity testing and confound
Checklist
Sediment samples are
sieved for the following
reasons:
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Field Sediment
• See Chapter 3 for proper
sample collection
procedures and sample
acceptability
Formulated Sediment
• Source of organic carbon
is critical: need to limit
sediment, oxygen demand
and ensure appropriate
particle size range for
study objectives
During compositing, homogenizing,
and subsampling, maintain anoxic
conditions if concerned with:
• Loss of volatile organics,
• Ammonia or sulfides, or metal
oxidation (Note: some metal
oxidation is expected during
sediment or interstitial water toxicity
testing)
Homogenize sample
prior to using
If anoxic sediment
conditions must be
maintained, sieve
in a glovebox or
other suitable
device in an inert
atmosphere.
Formulated sediment needs
to be equilibrated prior to
using for testing or analysis
Sieve sediments only if:
•Indigenous organisms may
interfere with assays
•Large debris will interfere
with assays
•Hand-picking is infeasible.
Figure 4-2. Alternatives for subsampling
Sediment Dilution
• Appropriate particle size
distribution needs to be
maintained as per study
objectives
• Anoxic sediments should
be diluted under an inert
atmosphere
• Organic carbon content
needs to be maintained at
appropriate levels
Spiking of sediments:
•Ensure homogenization and
quasi-equilibration
and compositing sediment grab samples.
Subsample and partition into
toxicity test chambers or
suitable containers for
chemical analyses
Figure 5-1. Flowchart depicting relationships between common sediment manipulations including
important considerations.
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interpretations of analytical results (USEPA, 1994; 2000d; ASTM, 2000e). Indigenous organisms
can be problematic in toxicity testing because they might be similar in appearance to test organisms
or they might prey on the test organisms.
If sieving is performed, it should be done for all samples to be tested, including control and reference
sediments if the objective of the study is to compare results among stations (ASTM, 2000a). It might
be desirable to obtain certain measurements (e.g., dissolved and total organic carbon, acid volatile
sulfide [AVS], and simultaneously extracted metals [SEM]) both before and after manipulation, to
document changes associated with sieving (USEPA, 2000d). In addition, it might be desirable to
document the effect of sieving on the sediment sample by conducting comparative toxicity tests using
sieved and unsieved sediment (Environment Canada, 1994).
Recommendation Box #1
Should sediment be sieved prior to analyses?
In general, sieving is not recommended because it can substantially change the
physicochemical characteristics of the sediment sample. However, sieving might be
necessary in preparing samples for some sediment toxicity testing (i.e., marine amphipod
tests; USEPA, 1994).
Unwanted materials (e.g., large particles and indigenous organisms) can be removed
from the sediment sample using forceps as a preferred alternative to sieving.
5.1.1 Sieving Methods
Press Sieving
If sieving is necessary, press sieving is the preferred method. In this method, sediment particles are
hand-pressed through a sieve using chemically inert paddles (Giesy et al., 1990; Johns et al., 1991).
Matter retained by the screen, such as organisms, shell fragments, gravel, and debris, should be
recorded in a log book and discarded (USEPA/ACOE, 1991). Samples with high debris, vegetation,
or clay content might be difficult to press through a single sieve with a mesh size less than 1 mm;
such samples might need to be pressed through a series of sieves with progressively smaller
openings. Water should not be added to sediment when press sieving, as this could result in changes
in contaminant concentration and bioavailability. Samples that are going to be used for both
chemical analysis and toxicity tests should be sieved together, homogenized, and then split for their
respective analyses.
Wet Sieving
If sediments cannot be press sieved without the addition of pressure, wet sieving might be required,
however, this type of sieving increases the likelihood of contaminant loss. Wet sieving involves
swirling sediment particles within a sieve using water to facilitate the mechanical separation of
smaller from larger particles. A slurry made with water that has separated from the sediment during
storage or transport might be sufficient to wash particles through the sieve. Wet samples that might
have settled during transit should be stirred to incorporate as much field water as possible. In some
cases, addition of a small volume of running site or deionized water might be required (ASTM,
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Photo courtesy of Allen Burton
Sieving a sediment sample for toxicity testing
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2000a). Mechanical shakers or stirring with a nylon brush can also facilitate wet sieving (Mudroch
and MacKnight, 1994).
Recommended Sieves
In general, smaller mesh sieves are
preferred to reduce loss of fines.
Stainless steel, brass, or plastic woven
polymer sieves (e.g., polyethylene,
polypropylene, nylon, and Teflon) with
mesh sizes that vary from 0.24 to 2.0 mm
have been used to sieve sediment for
toxicity tests (Keilty et al., 1988a;b;
Giesy et al., 1990; Lydy et al., 1990;
Stemmer et al., 1990a;b; Johns et al.,
1991; Landrum and Faust, 1991). Non-
metallic sieves are preferred if metals are
of interest. Stainless steel sieves are
acceptable if organic compounds are of
interest. Stainless steel (provided the
mesh is not soldered or welded to the
frame), nylon, or Nitex-type plastic
sieves are recommended when other
inorganic constituents are of concern or
are to be analyzed (ASTM, 2000a; PSEP, 1995).
Considerations
The mesh type and size
should be chosen based on
the following
considerations...
! The type of toxicity test and test organisms to
be used.
! Potential predators and/or competitors present
in the sample.
! Potential adsorption or contamination of the
chemical of interest due to sieving.
! The nature of the sample, including its particle
size distribution, volume, and size of debris.
Recommendation Box #2
What type of sieve should be
used?
Stainless steel or brass sieves are not
recommended when metals are a concern or
are analyzed (ASTM, 2000a).
Smaller mesh sieves (<2.0 mm mesh openings)
are recommended to reduce loss of fine particle
sizes.
Nylon or nitex mesh sieves are recommended
for inorganics analyses (e.g., metals).
Generally, sieving through a 10-mesh
(2-mm openings) sieve is acceptable as
a basis to discriminate between
sediment and other materials (ASTM,
2000a). For toxicity testing, the most
frequently used mesh size is 1.0 mm
(Environment Canada, 1994), which
will remove most adult amphipods.
However, a mesh of 0.25 mm might be
needed to remove immature
amphipods and most macrofauna
(Landrum et al., 1992; Robinson et al.,
1988; Day et al., 1995). In marine
sediments, sieves with a mesh size of
0.5 mm are effective in removing most
of the immature amphipods (Swartz et
al., 1990; PSEP, 1995).
5.1.2 Alternatives to Sieving
Unwanted materials (e.g., large particles, trash, and indigenous organisms), can be removed from the
sediment sample using forceps, prior to or, as an alternative to, sieving. If anerobic integrity of the
sample is not a concern, the sediment could be spread on a sorting tray made of cleaned, chemically-
inert material, and should be hand-picked with forceps. A stereomicroscope or magnifying lens
might facilitate the process, or may be used to determine if sieving is necessary. Hand-picking is
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preferable to sieving because it is less disruptive, but it typically is not practical for large volumes of
sediment. Of course, this process oxidizes the sediment and might alter contaminant bioavailability.
Autoclaving, freezing, and gamma irradiation of sediments are alternatives to physical removal for
inhibiting endemic biological activity in field-collected sediments. These are not generally
recommended procedures. Each method has unique effects on the physicochemical and biological
characteristics of the sediment, and a careful evaluation with respect to the study objectives is
warranted when these methods are considered.
Considerations
In preparing formulated sediments, the following should be
noted...
! Specific material sources should be carefully selected, as characteristics can vary
significantly among product types. A number of suppliers of various sediment components
are listed in ASTM (2000a) and USEPA (2000d).
! A critical component of formulated sediments is the source of organic carbon. It is not clear
that any one source of organic carbon is routinely superior to another. Alpha cellulose
appears to be a promising carbon source for formulated sediments.
! A variety of formulations have been used successfully in sediment toxicity testing (see
ASTM [2000a] and USEPA [2000d]). At this time, no one formulation appears to be
universally better than others.
5.2 Formulated Sediment and Organic Carbon Modification
5.2.1 General Considerations
Formulated sediments (also called
reconstituted, artificial, or synthetic
sediments) are mixtures of materials that
mimic the physical components of natural
sediments. While they have not been used
routinely, formulated sediments potentially
offer advantages over natural sediments for
use in chemical fate and biological effects
testing.
Formulated sediments also have limitations,
however. They do not possess the natural
microbial, meiofaunal, and macrofaunal
communities or the complex organic and
inorganic gradients prevalent in natural
sediments. The lack of biological activity,
diagenesis, and oxidation-reduction (redox)
potential gradients undoubtedly alters some
sorption and desorption properties, which
Checklist
Advantages of
Formulated Sediments
They provide a consistent, reproducible
medium which facilitates comparisons
between different sets of tests.
Eliminates interferences caused by the
presence of indigenous organisms.
Components of the formulated sediment
can be altered to measure the effect of
certain physicochemical characteristics
on chemical fate and bioavailability.
They are useful in spiking experiments
to obtain effect concentrations for
chemicals.
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might in turn alter contaminant fate and effects. The current lack of understanding of
physicochemical controls on bioavailability in different sediment environments precludes broad-scale
use of formulated sediments in definitive ecological risk assessments.
A formulated sediment should: (1) support the survival, growth, or reproduction of a variety of
benthic invertebrates, (2) provide consistent acceptable biological endpoints for a variety of species,
and (3) be composed of materials that have consistent characteristics (USEPA, 2000d; ASTM,
^^^^^^^^^^^^^^^^^^^^^^^^^^^^ 2000a). Characteristics should include:
(1) consistency of materials from batch to
Checklist
Disadvantages of
Formulated Sediments
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
other sources have caused dissolved oxygen concentrations to fall to unacceptable levels (Kemble et
al., 1999).
More about organic carbon modification:
Five studies compared organic carbon sources in formulated sediments. A study of 31 different
organic carbon recipes by Environment Canada (1995) compared effects on sediment
homogeneity, density, and turbidity. Cerophyll and trout chow were selected as the optimal
organic carbon sources with high clay (kaolin at 50 or 75% total concentration) and fine sand.
Ribeiro et al. (1994) recommended use of synthetic alpha-cellulose as a carbon source
amended with humic acid. This compound has since been tested by Kemble et al (1999),
Sawyer and Burton (1994), and Fleming and Nixon (1996). Ribeiro et al. (1994) found that
sorption was dependent on the amount of organic carbon present. Kemble et al. (1999) found
that growth and survival of Chironomus tentans and Hyalella azteca was better in 10% than in
2% alpha-cellulose. Both alpha-cellulose and conditioned red maple leaves were found to be
suitable as organic carbon amendments for reference toxicant testing with Hyallela azteca (96
hr) when spiked with cadmium, zinc, or anthracene (Sawyer and Burton, 1994).
Use of alpha cellulose as a carbon source for sediment-spiking studies has not been adequately
evaluated, but it appears to be promising. Alpha cellulose is a consistent source of organic
carbon that is relatively biologically inactive and low in concentrations of chemicals of concern.
Furthermore, Kemble et al. (1999) reported that conditioning of formulated sediment was not
necessary when alpha cellulose was used as a carbon source for a negative control sediment.
Compared with other sources of organic carbon, alpha cellulose is highly polymerized and would
not serve as a food source, but rather would serve to add texture or provide a partitioning
compartment for chemicals.
Reductions in organic carbon content have been achieved by diluting sediment with clean sand (See
Section 5.4; Clark et al., 1986; Clark et al., 1987; Tatem, 1986; Knezovich and Harrison, 1988).
However, this can change sediment characteristics resulting in non-linear responses in toxicity
(Nelson et al., 1993). Combustion has also been used to remove fractions of organic carbon (Adams
et al., 1985; IJC, 1988). However, this method results in substantial modification of the sediment
characteristics, including oxidization of some inorganic components.
The ratio of carbon to nitrogen to phosphorous might be an important parameter to consider when
selecting an organic carbon source. This ratio can vary widely among carbon sources (ASTM,
2000a; USEPA, 2000d). For example, carbon can range from 30 to 47%, nitrogen from 0.7 to 45
mg/g, and phosphorous from below detection limits to 11 |ig/g for several different carbon sources
(USEPA, 2000d).
A variety of formulations have been used successfully in sediment toxicity testing (see ASTM, 2000a
and USEPA, 2000d). At this time, no one formulation appears to be universally better than others.
5.3 Spiking
Spiking involves adding one or more chemicals to sediment for either experimental or quality control
purposes. Spiking environmental samples is used to document recoveries of an analyte and thereby
analytical bias. Spiked sediments are used in toxicity tests to determine effects of material(s) on test
species. Spiking tests can also provide information concerning chemical interactions and
transformation rates. The design of spiking experiments, and interpretation of results, should always
consider the ability of the sediment to sequester contaminants, recognizing that this governs many
chemical and biological processes (O'Donnel et al., 1985; Stemmer et al., 1990a;b; ASTM, 2000a;
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Northcott and Jones, 2000). In preparation for toxicity and bioaccumulation tests, references
regarding the choice of test concentrations should be consulted (USEPA, 2000d; ASTM, 2000a;
Environment Canada, 1995). Program specific guidance documents should also be consulted as
appropriate.
Several issues regarding sediment spiking are addressed in this section. First, several methods have
been used to spike sediments but the appropriate method needs to be selected carefully depending on
the type of material being spiked (e.g., soluble in water or not), its physical-chemical form, and
objectives of the particular study. Second, spiked material should be uniformly distributed
throughout the sediment. Otherwise, analyses or toxicity tests are likely to yield highly variable
results, depending on the concentration of spiked material present. Third, the spiked material needs
to be at equilibrium between the sediment and the interstitial water to ensure that all relevant
exposure phases are appropriately considered in chemical analyses or toxicity testing. The time it
takes to reach this equilibrium is a critical factor that needs to be considered and documented.
Recommendation Box #3
How should sediments be spiked with a chemical or other test
material?
Regardless of the spiking technique used, care should be taken to ensure complete and
homogenous mixing.
Replicate subsamples should be analyzed to confirm homogeneous mixing.
Moisture content should be determined on triplicates for each sample so that the spike
concentration can be normalized on a dry weight basis.
Wet spiking is recommended over dry spiking methods.
Generally speaking, the jar rolling method is more suitable than hand mixing for spiking
larger batches of sediment.
To ensure chemical equilibrium between the sediment and pore water in toxicity testing,
spike sediments should be stored for at least one month, unless other information is
available for the spiking material and sediment type.
Direct addition of organic solvent carriers should be avoided because they might alter
sediment chemistry and affect contaminant bioavailability. Shell coating methods should
be used instead as this eliminates many of the disadvantages of solvent carriers.
5.3.1 Preparation for Spiking
Debris and indigenous organisms should be removed from sediment samples as soon as possible after
collection to reduce deterioration of sediment quality due to decomposition of organic debris and
dying infauna. If sediments are to be stored prior to spiking, they should be kept in sealed containers
at4°C.
Regardless of the spiking technique used, care should be taken to ensure complete and homogenous
mixing (See Section 4.4). It is recommended that chemical analyses be conducted to verify that
concentrations of the spiked contaminants are uniform throughout the mixed material. Three or more
subsamples of the spiked sediment should be randomly collected to determine the concentration of
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the substance being tested. In general, the coefficient of variation (C V) should be < 20% for
homogeneity of mixing to be considered sufficient (ASTM, 2000a; Northcott and Jones, 2000).
Temperatures should be kept cool during spiking preparation (e.g., 4° C) due to rapid
physicochemical and microbiological alterations which might occur in the sediment that, in turn,
might alter bioavailability and toxicity (ASTM, 2000a; Environment Canada, 1995). If spiking PAH
compounds, it might be important to conduct spiking in the dark, or at least under low light as PAH
toxicity has been shown to increase under ultraviolet light (Ankley et al., 1994).
It is recommended that a subsample of the spiked sediment be analyzed for at least the following
parameters: moisture content, pH, ammonia, total organic carbon (TOC), acid volatile sulfide
(AVS), particle size distribution, and background levels of the chemical(s) to be spiked. Further
characterization may include analyses of total volatile residue, pore water salinity (before and after
any sieving), chemical oxygen demand, sediment oxygen demand, oxidation-reduction potential (Eh),
metals, total chlorinated organic content, chlorinated organic compounds, and polycyclic aromatic
hydrocarbons (see Appendix G for more information on physicochemical parameters often measured
on sediments). It is particularly important to determine the TOC concentration if the sediment is to
be spiked with a nonionic organic compound, as organic carbon is the primary binding phase for such
compounds (DiToro et al., 1990). Similarly, the concentration of AVS (the primary binding phase
for cationic metals in anoxic sediments) and TOC should be measured after spiking with a cationic
metal (Ankley et al., 1996; Leonard et al., 1999).
The sediment moisture content measurement is used to standardize the amount of chemical spiked on
a dry weight basis (see Appendix G). Generally, the moisture content should be determined on
triplicates for each sample by measuring the weight lost following 24 h of oven-drying at 105 °C.
After drying, the samples should be cooled to room temperature in a desiccator before taking dry
weight measurements (Yee et al., 1992). The mean wet density, expressed as mg water/cm3, is
measured by using the same drying method on known sediment volumes. This allows spiking to be
normalized from a volume basis to an equivalent dry weight basis.
5.3.2 Methods for Spiking
Spiking of both wet and dry sediments is common, but wet spiking is recommended because drying
might reduce the representativeness of the sample by changing its physicochemical characteristics
(ASTM, 2000a). Methods differ mainly in the amount of water present in the mixture during
spiking, the solvent used to apply the toxicant, and the method of mixing. Generally speaking, the jar
rolling method is more suitable than hand mixing for spiking larger batches of sediment.
In addition to the above techniques, sediments may be spiked by hand stirring using a scoop or
spatula, as long as the homogeneity of the mixture is verified. Eberbach and gyro-rotary shakers
have also been used effectively to mix spiked sediments (Stemmer et al., 1990a). Less commonly,
chemical(s) are added to the water overlying the sediment and allowed to sorb with no mixing
(Stephenson and Kane, 1984; O'Neill et al., 1985; Crossland and Wolff, 1985; Pritchard et al., 1986).
Sediment Rolling
One of the recommended wet sediment rolling techniques requires a specific jar-rolling apparatus,
first described by Ditsworth et al. (1990). Many other jar-rolling apparatuses are available, ranging
in size and options available. This "rolling mill" method has been used to homogenize large volumes
of sediments spiked with metals and non-ionic organic compounds. The primary disadvantage of this
method is that the mixing apparatus must be constructed or purchased.
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The jar-rolling apparatus used by Ditsworth et al. (1990) consists of eight parallel, horizontal rollers
powered by an electric motor through a reduction gear, belts, and pulleys, which rotate cylindrical
vessels containing the substrate mixtures. Mixing is accomplished gravimetrically by slowly rolling
the jars (gallon-sized jars can be rolled at approximately 15 rpm). Optimally wetted, individual
substrate particles adhere to each other and to the wall of the revolving jar until they cascade or
tumble down the surface of the substrate mass. Dilution water may be added to the substrate before
rolling to adjust the sediment-to-water ratio for
optimal mixing. If oxidation is a concern (for
example, if the sample will be analyzed for metals),
jar contents might need to be maintained in an inert
atmosphere. If PAHs are of concern then jars should
be shielded from light (Ankley et al., 1994).
Considerations
When rolling
sediment:
Jars should not be overfilled, as
this will reduce mixing efficiency.
Prolonged rolling (e.g., > 1 wk)
should be avoided to minimize
physicochemical changes to the
sediment.
Each jar should be loaded with the required amount of
wet base sediment (with a calculated mass of dry
sediment required for the test) prior to introduction of
the toxicant. Several 1-cm diameter holes of different
depths should be punched into the sediment to provide
more surface area for the initial distribution of the test
material. A predetermined volume of the stock
solution or a serial dilution of the stock should be
used to spike each jar load of sediment. A volumetric
pipette should be used to distribute each aliquot onto the top surface and into the holes of the
sediment in each jar. Sediments should be spiked sequentially, proceeding from low to high
concentrations of test material, to minimize cross-contamination. Control substrates should be
prepared by adding an equivalent volume of dilution water to a jar loaded with unspiked sediment.
After spiking, all jars and their contents should be processed identically.
Typically, jars should be rolled for greater than two hours to achieve sample homogeneity. Jars
should be closely monitored during the first hour of rolling to ensure proper mixing of substrates.
After rolling for approximately 15 min, mixing efficiencies of the substrates can be judged visually.
If a sediment displays excessive cohesiveness, as indicated by agglomerating or balling, the jars
should be opened and an aliquot of appropriate dilution water (50 mL of either saltwater or
freshwater depending on the source of the sediment) added to each substrate to increase the fluidity.
This procedure should be repeated as necessary until the operator visually observes that all substrates
are tumbling without forming balls. Adding water in small rather than large aliquots can prevent
over-saturation of the sediment. Over-saturation is undesirable because excess water must be
decanted following rolling, prior to sediment testing.
After rolling, the jars should be gently shaken to settle sediment that adhered to the walls. They may
be set upright and stored overnight in the dark at room temperature or at an alternate temperature
(e.g., 4° C) depending on the study objectives. After equilibration (see Section 5.3.3) and prior to
distributing the sample to test chambers, additional rolling for two hours will help integrate
interstitial water into the sediment.
Sediment Suspension Spiking
The sediment suspension technique (Cairns et al., 1984; Schuytema et al., 1984; Stemmer et al.,
1990a; b; Landrum and Faust, 1991; Landrum et al., 1992) is the simplest of the three spiking
techniques and requires the least equipment. The method involves placing dilution water and
sediment together in a 1-L beaker. The desired amount of toxicant, dissolved in dilution water, is
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added to the beaker. The mixture should be stirred at a moderate speed with a stir bar, or mechanical
stirrer, for a minimum of four hours. The sediment in the beakers should then be allowed to settle
and equilibrated at the appropriate test temperature as specified in the test method. The excess water
overlying the sediment is decanted and discarded, and the sediment is distributed to the test
containers (Environment Canada, 1995).
Slurry Spiking
The slurry technique (Birge et al., 1987; Francis et al., 1984; Landrum and Faust, 1991; Landrum et
al., 1992) requires a minimum of equipment and involves less water than the sediment suspension
technique. A 250-g dry weight sample of sediment is placed in a 500-mL Erlenmeyer flask. Via a
25 -mL aliquot of distilled, deionized water, a sufficient concentration of the materials of interest is
added to obtain the desired sediment concentration (mg/kg, dry weight basis). Control (unspiked)
sediment receives a 25-mL aliquot of distilled, deionized water having no added materials. The
sealed flask may be mixed using various methods such as continuous agitation in a shaker for five
days (Birge et al., 1987) or vigorous shaking for 60 seconds, twice daily for seven days (Francis et
al., 1984). Following mixing, the sediment suspensions should be centrifuged to remove water. The
moisture content of the sediment should be approximately 15% to 20% after centrifugation. After
removal of excess water, the prepared sediment can be placed in the exposure chambers and covered
with dilution water according to the specific test methods. This procedure often yields sediment
having its original moisture content.
5.3.3 Equilibration Times
Prior to distributing the spiked sediment to containers for toxicity testing or chemical analyses, the
spiked sediments should be stored for a sufficient time to approach chemical equilibrium in the
test material between the sediment and interstitial water. Equilibration times for spiked sediments
vary widely among studies (Burton, 1991), depending on the spiking material and sediment type. For
metals, equilibration time can be as short as 24 h (Jenne and Zachara, 1984; Nebecker et al., 1986),
but one to two weeks is more typical (ASTM, 2000a). For organic compounds with low octanol-
water partition coefficients (K^), equilibration times as short as 24 h have been used (Dewitt et al.,
1989). Some organic contaminants might undergo rapid microbiological degradation depending on
the microbial population present in the sample. In these cases, knowledge of microbial effects might
be important in defining an appropriate equilibration period. Organic compounds with a high
partition coefficient might require two months or more to establish equilibrium (Landrum et al.,
1992). Boundaries for the sorption time can be estimated from the partition coefficient, using
calculations described by Karickhoff and Morris (1985a, b). It is important to recognize that the
quantity of spiked chemical might exceed the capacity of the test sediment system, prohibiting
equilibrium.
For research purposes, unless definitive information is available regarding equilibration time for a
given contaminant and sediment concentration, a one-month equilibration period is recommended,
with consideration that two months might be needed in some instances (USEPA, 2000d). For
regulatory programs, however, sample holding time should not exceed 2 weeks. Therefore, for these
programs spiking equilibration time should not exceed 2 weeks. Periodic monitoring during the
equilibration time is highly recommended to empirically establish stability of interstitial water
concentrations (USEPA, 2000d). Sediment and interstitial water chemical concentrations should also
be monitored during long-term bioassay tests to determine the actual chemical concentrations to
which test organisms are exposed, and to verify that the concentrations remain stable over the
duration of the test.
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5.3.4 Use of Organic Solvents
Direct addition of organic solvents should be avoided if possible, because they might dramatically
affect sediment geochemistry and alter bioavailability (USEPA, 2000d). However, many organic
materials require use of a solvent to adequately mix with the sediment. If an organic solvent is to be
used, the solvent should be at a concentration that does not affect test organisms and should be
uniform across treatments. Further, both solvent control and negative control sediments should be
included in tests with solvents. The solvent concentration in the control should equal the treatment
concentration and should be from the same batch used to make the stock solution (ASTM, 2000a).
To reduce the possibility of solvent-related artifacts, the spiking process should include a step which
allows the solvent to evaporate before addition of sediment and water followed by rolling (McLeese
et al., 1980; Muir et al.,1982; Adams et al., 1985). Highly volatile organic compounds have been
spiked into sediments using co-solvents followed by shaking in an aqueous slurry. When highly
volatile compounds are used, immediate testing in covered flow-through systems is recommended
(Knezovich and Harrison, 1988).
There is some uncertainty concerning artifacts introduced by the use of solvents. The use of a polar,
water soluble carrier such as methanol was found to have little effect on the partitioning of nonpolar
compounds to dissolved organic matter at concentrations up to 15% carrier by volume (Webster et
al., 1990). However, another study showed that changes in partitioning by a factor of approximately
two might occur with 10% methanol as a co-solvent for anthracene sorption (Nkedi-Kizza et al.,
1985). The effect of carrier volume on partitioning of organic chemicals in sediments is equivocal.
However, because solvents might be either directly or indirectly toxic to the test organisms, caution
should be taken to minimize the amount of carrier used. In addition, the use of a carrier such as
acetone might result in faster equilibration of spiked organic compounds (Schults et al., 1992).
Shell coating techniques which introduce dry chemical(s) to wet sediment have also been developed,
principally to eliminate the potential disadvantages of solvent carriers. The chemical may be either
coated on the inside walls of the container (Ditsworth et al., 1990; Burgess et al., 2000) or coated
onto silica sand (Driscoll et al., 1997; Cole et al., 2000). In each shell coating method, the chemical
is dissolved in solvent, placed in a glass spiking container (with or without sand), and the solvent is
slowly evaporated prior to addition of the wet sediment. Wet sediment then sorbs the chemical from
the dry surfaces. It is important that the solvent be allowed to evaporate prior to adding sediment or
water.
5.4 Preparation of Sediment Dilutions
Spiked or field-contaminated sediments can be diluted with whole sediment to obtain different
contaminant concentrations for concentration-effects testing. The diluent sediment should have
physicochemical characteristics similar to the test sediment, including organic carbon content and
particle size, but should not contain concentrations of contaminants above background levels
(ASTM, 2000a; Burton, 1991). Diluent sediment has included formulated sediment as well as known
reference site sediment. Diluted sediment samples should be homogenized and equilibrated in
accordance with procedures described in Sections 4.4 and 5.3.3, respectively.
The diluent sediment should be combined with the test sediment in ratios determined on a dry weight
basis to achieve the desired nominal dilution series (DeWitt, personal communication). Volume to
volume dilutions have also been performed (e.g., Schlekat et al.,1995; Johns et al., 1985), but weight
to weight dilutions are preferred because they provide more accurate control and enable a more
straightforward calculation of dose-response curves.
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Results from dilution experiments should be interpreted with care. There are often non-linear
responses due to non-equilibrium, non-linear sorption-desorption processes that cannot always be
adequately controlled (Nelson et al., 1993). Nelson et al. (1993) found that analyses of diluted
sediments did not match nominal concentrations as estimated by physical characteristics. They
suggested that chemical characterization is needed to determine effects of manipulations (i.e.,
mixing) and resulting changes (i.e., oxygenation of complexing agents such as acid volatile sulfides).
5.5 Preparation of Sediment Elutriates
Many studies of sediment toxicity have evaluated aqueous extractions of suspended sediment called
elutriates. The elutriate method was initially developed to assess the effects of dredging operations
on water quality (U.S. ACOE, 1976). Elutriate manipulations are also applicable to any situation
where the resuspension of sediment-bound toxicants is of concern, such as bioturbation and storms,
that might disturb sediments and affect water quality (USEPA/ACOE, 1991, 1998; Ankley et al.,
1991). USEPA/ACOE (1998) lists eighteen freshwater and saltwater aquatic organisms as
candidates for elutriate toxicity testing. Standard effluent toxicity test procedures are also
appropriate for elutriates, including tests with various vascular and non-vascular plant species
(Ingersoll, 1995).
Elutriate tests are not intended to reflect the toxicity of interstitial waters or whole sediments, as
there are differences in contaminant bioavailability in the two types of media (Harkey et al., 1994).
In general, elutriates have been found to be less toxic than bulk sediments or interstitial water
fractions (Burgess et al., 1993; Ankley et al., 1991), although in some studies elutriates have been
found to be more toxic (Hoke et al., 1990) or equally as toxic (Flegel et al., 1994) relative to
interstitial water.
While there are several procedural variations, the basic method for elutriate preparation involves
combining various mixtures of water and sediment (usually in the ratio of 4 parts water to 1 part
sediment, by volume) and shaking, bubbling or stirring the mixture for 1 hour (Ross and Henebry,
1989; Daniels et al., 1989; Ankley et al., 1991; Burgess et al., 1993; USEPA/USACOE, 1991, 1998).
It is likely that chemical concentrations will vary depending on the elutriate procedure used. Specific
program guidance should be consulted as appropriate. The water phase is then separated from the
sediment by settling and/or centrifugation (Note: the dredging remediation program does not always
require centrifuging elutriates). Once an elutriate has been prepared, it should be analyzed or used in
biological tests immediately, or as soon as possible thereafter. It should be stored at 4 °C for not
longer than 24 h, unless the test method dictates otherwise (Environment Canada, 1994;
USEPA/ACOE, 1991, 1998). For toxicity test exposures exceeding 24 h, fresh elutriate should be
prepared daily.
Filtering the elutriate is generally discouraged, but it might be prescribed for some toxicity tests.
Filtration can reduce the toxicity of sediment elutriates due to sorption of dissolved chemicals on the
filtration membrane and retention of colloids. If colloidal material needs to be removed, serial or
double centrifugation is generally a preferred alternative. If an elutriate must be filtered, it is
recommended that only pre-treated filters be used and that the first 10 to 15 mL of the elutriate to
pass through the filter be discarded (Environment Canada, 1994). Testing with a filtered elutriate
should include an assessment to determine the extent of analyte adsorption/desorption to/from the
filter.
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Recommendation Box #4
How should sediment elutriates be performed?
Combine 1 part sediment with 4 parts water unless a specific program stipulates
otherwise.
The sediment-water mixture should be vigorously shaken, bubbled or stirred for 1 hour.
Centrifugation is a useful means for isolating the water phase (elutriate).
Once prepared, the elutriate should be analyzed or tested as soon as possible.
Store elutriate at 4° C with little or no headspace until analysis.
Filtering the elutriate (to remove colloidal material) is generally not recommended. Use
double (serial) centrifugation if appropriate.
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Collection of Interstitial Water
Sediment interstitial water, or pore water, is defined as the water occupying the spaces between
sediment particles. Interstitial water might occupy about 50% (or more) of the volume of a
depositional (silt-clay) sediment. The interstitial water is in contact with sediment surfaces for
relatively long periods of time and therefore, might become contaminated due to partitioning of the
contaminants from the surrounding sediments. In addition, interstitial waters might reflect ground
water - surface water transition zones in upwelling or downwelling areas. In these areas their
chemistry might be more reflective of ground or surface waters at the site. Therefore, flow, residence
time and other physicochemical factors (e.g., pH, temperature, redox potential, organic carbon,
sulfides, carbonates, mineralogy) might have varying roles in determining whether interstitial waters
are contaminated.
In many depositional sediments, interstitial waters are relatively static, and therefore contaminants in
the interstitial water and in the solid phase are expected to be at thermodynamic equilibrium. This
makes interstitial waters useful for assessing contaminant levels and associated toxicity. Interstitial
water is often isolated to provide either a matrix for toxicity testing and/or to provide an indication of
the concentration and/or partitioning of contaminants within the sediment matrix.
6.1 General Procedures
Interstitial water sampling has become especially important in regulatory programs because
interstitial water toxicity tests yield additional information not currently provided by solid-phase,
elutriate, or sediment extract tests (Carr and Chapman, 1992; SETAC, 2001). Furthermore,
interstitial water toxicity tests have proven to be useful in sediment toxicity identification evaluation
(TIE) studies (e.g., Burgess et al., 1996; Carr, 1998; Burton, 2001) as test procedures and sample
manipulation techniques are generally cheaper, faster, and easier to conduct than solid-phase tests
(SETAC, 2001). Thus, the collection of interstitial water has become increasingly important in
sediment quality monitoring and remediation programs.
Interstitial water sampling is most suitable for sediment types ranging from sandy to uncompacted
silt-clays (Sarda and Burton, 1995; SETAC, 2001). Such sampling is not typically performed on
sediments with coarse particle size (such as gravel) or on hard, compacted clays, as the potential for
interstitial water contamination in these sediment types is relatively low.
As with all sampling discussed in this manual, the principle aim is to use procedures that minimize
changes to the in situ condition of the water. It should be recognized that most sediment collection
and processing methods have been shown to alter interstitial water chemistry (e.g., Schults et al.,
1992; Bufflap and Allen, 1995; Sarda and Burton, 1995), thereby potentially altering contaminant
bioavailability and toxicity.
Laboratory-based methods (e.g., centrifugation, pressurization, or suction) are commonly used as
alternatives to in-situ interstitial water collection (see Section 6.2). While these methods have been
shown to alter interstitial water chemistry, they're sometimes necessary or preferred, especially when
larger sample volumes are required (e.g., for toxicity testing).
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
As both in-situ and laboratory-based or ex-situ (e.g., methods might be appropriate for many study
objectives, it is critical that the same procedures are used for all stations sampled in a study, or
program, so that appropriate sample comparisons can be made. Furthermore, the sediment depth
at which interstitial water is sampled (either using in-situ or ex-situ extraction methods) should
match the depth of interest in the study (SET AC, 2001). For example, samples for dredging
remediation should be sampled to the depth to be disturbed by dredging activity, whereas samples for
a status and trends survey should be collected at the biologically active depth (often < 15 cm).
Figure 6-1 summarizes the major considerations for selecting in-situ or ex-situ procedures in a given
study.
The two major issues of concern regarding interstitial water sample integrity are: 1) the ability of the
sampling device to maintain physicochemical conditions in the natural state by minimizing
adsorption/leaching of chemicals to/from the device, and 2) the ability to maintain the sample in the
redox state existing at the site. Precautions required to reduce the likelihood of sample artifacts will
vary with each study as indicated in the following sections.
6.2 In-situ Collection
In situ methods might be superior to ex-situ methods for collecting interstitial water, as they are less
subject to sampling/extraction related artifacts and therefore, might be more likely to maintain the
chemical integrity of the sample (Sarda and Burton, 1995; ASTM 2000a; SET AC, 2001). jjowever,
in situ methods have generally produced relatively small volumes of interstitial water, and often
limited to wadeable or diver-accessible
water depths. These logistical
constraints have limited their use and
applicability in sediment monitoring
studies.
The principal methods for in situ
collection of interstitial water involve
either deployed "peepers" (Bufflap and
Allen, 1995; Brumbaugh et al., 1994;
Adams, 1991; Carignan and Lean, 1991;
Carignan et al., 1985; Bottomley and
Bayly, 1984) or suction techniques
(Watson and Prickers, 1990; Knezovich
and Harrison, 1988; Howes et al., 1985).
A summary of these methods is
provided in Table 6-1. Both methods
have a high likelihood of maintaining in
situ conditions. In cases where in situ
deployment is impractical, peepers or
suction devices can be placed in
relatively undisturbed sediments
collected by core or grab samplers (see
Chapter 3).
Recommendation Box #1
In-situ interstitial water
collection
Use peepers for sampling interstitial waters,
rather than (or in addition to) grab or core
sediment extractions if site conditions, volume
requirements, and logistical considerations
allow.
Reduce potential for oxygenation of samples
by proper deployment and retrieval
procedures.
Allow adequate equilibration of peepers prior
to sampling.
Minimize handling and processing of field-
collected interstitial waters.
Field collected interstitial water samples should
be stored in containers, without headspace at
4° C in the dark, until analyzed/tested.
Samples for certain chemical analyses (e.g.,
pesticides, phenols), should be frozen or
preserved immediately.
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There is a need for interstitial
water chemistry or toxicity
information if study objectives
include any of the following:
Verify effect based on sediment quality
guidelines (e.g.,Ecotox Thresholds, ER-M)
Build a weight-of-evidence conclusion
Assess exposures and/or effects in a more
bioavailable compartment
Use water column-based assays
Apply Toxicity Identification Evaluation
methods
Assess upwelling, downwelling, or
dynamic interstitial water conditions
Use peepers if:
If peepers are
not feasible:
Station is shallow and
peeper can be manually
deployed
Minimal pore water
volume needed
Expertise available
Sediment depth of
concern matches peeper
exposure
Equilibration time can
be met
Use least destructive
sediment sampling
method: Core>Ekman>
Ponar>Van Veen (see
Chapter 3).
Isolate interstitial water
by centrifugation, or by
squeezing or suction in
that order of preference
Peeper Design:
Equilibration time is dependent on:
sediment, chamber size, mesh size
Increase mesh size to speed
equilibration and allow transport of
larger particles.
Pre-purge system if oxidation is a
concern.
See
Figure 6-2
Figure 6-1. Considerations for selecting the appropriate type of interstitial water sampling method.
Chapter 6: Collection of Interstitial Water
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Photo and illustration on this page, courtesy of Allen Burton
, *' ," ';-.%
Peepers deployed in the field
General peeper design with in-situ sample extraction
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6.2.1 Peeper Methods
Peepers are small chambers with membrane or mesh walls containing either distilled water or clean
water of the appropriate salinity or hardness. Samples are collected by burying the devices in
sediments and allowing surrounding interstitial waters to infiltrate. In principle, dissolved solutes
will diffuse through the porous wall into the peeper and the contained water will reach equilibrium
with the ambient interstitial water. The design concept for sediment peepers originated as
modifications of the dialysis bag technique used by Mayer (1976) and Hesslein (1976), and has been
modified successfully for use in laboratory sediment toxicity tests (Doig and Liber, 2000). The
initial designs consisted of either a flat base plate or a cylindrical dialysis probe (Bottomley and
Bayly, 1984) with compartments covered by dialysis membranes and a manifold for collection of
multiple samples at various depths in the sediment profile (Figure 6-2). Further modifications to
these designs have incorporated sampling ports, large sample compartments, and various types of
membranes with different pore sizes. These modifications are usually required based on specific
project objectives regarding sample volumes and contaminants of interest.
Table 6-1. In-situ interstitial water collection methods (Sarda and Burton, 1995; SET AC, 2001).
Device
Peeper
In situ
Suction
Sediment
Depth
(cm)
0.2-10
0.2-30
Sample
Volume
(L3)
<0.5
< 0.25
Advantages
Most accurate method, reduced
artifacts, no lab processing;
relatively free of effects from
temperature, oxidation, and
pressure; inexpensive and easy to
construct; some selectivity
possible depending on nature of
sample via specific membranes;
wide range of membrane/mesh
pore sizes, and/or internal solutes
or substrates available.
Reduced artifacts, gradient
definition; rapid collection, no lab
processing; closed system which
prevents contamination; methods
include airstone, syringes, probes,
and core-type samplers.
Disadvantages
Requires deployment by hand, thus
requiring diving in > 0.6 m depth
water; requires hours to days for
equilibration (varies with site and
chamber); methods not
standardized and used infrequently;
some membranes such as
dialysis/cellulose are subject to
biofouling; must deoxygenate
chamber and materials to prevent
oxidation effects; some
construction materials yield
chemical artifacts; some chambers
only allow small sample volumes;
care must be used on collection to
prevent sample oxidation.
Requires custom, non-standard
collection devices; small volumes;
limited to softer sediments; core
airstone method; difficult in some
sediments and in deeper water (>1
m); method might require diving for
deployment in deep waters;
methods used infrequently and by
limited number of laboratories.
Note: Incorporation of filtration into any collection method might result in loss of metal and organic
compounds.
Various peeper devices have been recently used effectively to collect interstitial water. For example,
a simplified design using a 1 jam polycarbonate membrane over the opening of a polyethylene vial
was successful in capturing elevated levels of copper and zinc (Brumbaugh et al., 1994). Other
Chapter 6: Collection of Interstitial Water
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Dialyzer Sampler or "Peeper"
Assembled
£1 „
Sampler components
Base plate
Dialysis
membrane
Cover
INTERNAL UN(T SAMPLER fWackeO
LENGTH. 4cm
•« GROOVE FOR O-RINQ
.,3 OPEN PORTS
(membrane removed)
-RUBBER SEPTUM
Figure 6-2. Front view and components of peeper sampling devices (top: plate device;
bottom: cylindrical probe)
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designs have been used to collect nonpolar organic compounds in a variety of aquatic systems
(Bennett et al., 1996; Axelman et al., 1999) and in overlying water (Huckins et al., 1990).
Peepers have also been used to expose organisms to sediments in situ (Burton et al., 2001). Burton et
al. (1999) successfully introduced organisms to aerobic sediments using peepers. However, anoxic
sediments are not amenable to in situ organism exposure.
Different materials might be advisable in constructing peepers depending on the contaminants of
concern. For example, for many contaminants, peepers constructed from acrylic material appear to
yield interstitial water samples with minimal chemical artifacts (Burton et al., 2001). Some polymer
materials might be inappropriate for studies of certain nonpolar organic compounds. Cellulose
membranes are also unsuitable, as they decompose too quickly. Plastic samplers can contaminate
anoxic sediments with diffusible oxygen (Carignan et al., 1994).
In preparation for interstitial water collection, peeper chambers should be filled with deoxygenated
water, which can be prepared by nitrogen purging for 24 hours prior to insertion. If sediment
oxidation is a concern, the peepers should be transported to the deployment site in a sealed oxygen-
free water bath to avoid potential changes to the sediment-water equilibrium caused by dissolved
oxygen interactions. However, during peeper equilibration periods, anoxic conditions are likely to be
quickly reestablished. In addition, when samples are collected and processed, exposure to oxygen
should be minimized.
Following initial placement, the equilibration time for peepers may range from hours to a month, but
a deployment period of one to two weeks is most often used (Adams, 1991; Call et al., 1999; Steward
and Malley, 1999). Equilibration time is a function of sediment type, study objectives, contaminants
of concern, and temperature (e.g., Skalski and Burton, 1991; Carr et al., 1989; Howes et al., 1985;
Simon et al., 1985; Mayer, 1976). Membrane pore size also affects equilibration time, with larger
pore sizes being used to achieve reduced equilibration times (Sarda and Burton, 1995). For example,
using a peeper with a 149-|am pore size, Adams (1991) reported equilibration of conductivity within
hours of peeper insertion into the sediment. Thus, it appears that equilibration time is a function of
the type of contaminant, sediment type, peeper volume, and mesh pore size.
Peepers with large-pored membranes, while shortening equilibration time, also allow particulates to
enter the chamber. The larger solids tend to settle to the bottom of the peeper chamber, and caution
should be used to avoid collecting the solids when retrieving the water sample from the chamber.
Colloidal particles will remain suspended in the sample and thereby present an artifact, but the
concentration of such particles is typically lower than that found in laboratory- centrifuged samples
(Chin and Gschwend, 1991).
In several studies, analysis of interstitial water from replicate peepers has demonstrated from low to
high heterogeneity in water quality characteristics (Frazier et al., 1996; Sarda and Burton, 1995).
The potential for high variability in interstitial water chemical characteristics should be taken into
account when developing the sampling design.
6.2.2 Suction Methods
There are a variety of suction devices for collecting interstitial water. A typical suction device
consists of a syringe or tube of varying length, with one or more ports located at the desired sampling
positions (ASTM, 2000a). The device is inserted into the sediment to the desired depth and a
manual, spring-operated, or vacuum gas suction is applied to directly retrieve the water sample. A
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variation on this approach employs a peeper-like porous cup or perforated tube with filters. The unit
is inserted into the sediment for a period of time, allowing interstitial water to infiltrate the chamber
before suction is applied. The samples are then retrieved by suction. Another variation that has been
used successfully employs an airstone embedded into the sediment which forces interstitial water
upward where it can be collected via syringe or tube. All of these suction methods generally yield
smaller quantities of interstitial water than peepers and chemical (lexicological) artifacts are more
likely due to greater potential exposure of interstitial water to oxygen (ASTM, 2000a).
6.2.3 Processing of Field-Collected Interstitial Water Sample
Following sample retrieval, interstitial water might need to be recovered and stabilized quickly to
prevent oxidative changes or volatilization (Carignan, 1984). Containers should be filled, with no
headspace to minimize changes in dissolved oxygen and contaminant bioavailability. Procedures for
stabilization are dependent on the analyses to be performed. When non-volatile compounds are the
target analytes, acidification is often stipulated, while organic carbon and methane may be stabilized
with saturated mercury chloride (Mudroch and MacKnight, 1994).
Samples to be analyzed for toxicity, are normally cooled to 4° C as soon as possible for transport to
the laboratory. EPA methods for toxicity testing of surface waters and effluents (USEPA 1991)
recommend that samples not be frozen in storage or transport. However, recent information suggests
that freezing of interstitial water may not affect toxicity in some cases (Ho et al., 1997; Carr and
Chapman, 1995; SETAC, 2001). Unless a demonstration of acceptability is made for the sites of
interest, interstitial water samples should not be frozen prior to biological testing. Samples for
chemical analyses should be preserved immediately, if appropriate, or cooled to 4° C as soon as
possible.
6.3 Ex-situ Extraction of Interstitial Water
Ex-situ interstitial water collection methods are often necessary when relatively large volumes of
interstitial water are required (such as for toxicity testing), when in-situ collection is not viable or
when a brief sampling time is critical. While these extraction methods can be done in the field or in
the laboratory, extraction in the laboratory, just prior to analysis or testing, is preferable so that the
sample is maintained as close to its original state as much as possible during transport and storage
(SETAC, 2001). Guidance in this chapter reflects recommendations presented in several recent
publications including proceedings from two workshops devoted entirely to interstitial water
extraction methods, water handling, and use in toxicity applications: (1) a dredged materials
management program workshop on interstitial water extraction methods and sample storage in
relation to tributyltin analysis (Hoffman, 1998) and (2) a Pellston workshop on interstitial water
toxicity testing including interstitial water extraction methods and applications (SETAC, 2001).
Figure 6-3 summarizes many of the issues associated with laboratory isolation of interstitial water
discussed in this section.
6.3.1 General Procedures
Centrifugation and squeezing are the two most common techniques for collecting interstitial water,
and are generally preferred when large volumes are required. Other methods include pressurization
(e.g., vacuum filtration) devices, which can be used to recover small volumes of interstitial water.
Regardless of the method used, interstitial water should be preserved immediately for chemical
analyses, if appropriate, or analyzed as soon as possible after sample collection if unpreserved (such
as for toxicity testing; Hoffman, 1998; SETAC, 2001). Significant chemical changes can occur even
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Mix water that has
separated from sediment
during storage into
sediment
May need to minimize
oxidation as in whole
sediments.
Double (repeated)
centrifugation improves
particle (colloidal)
removal.
Complete extraction as
soon as possible.
Centrifuge at high speed (e.g., 8000-
10,000 x g) for 30 minutes is
suggested for toxicity testing unless
study-specific information or
objectives dictate otherwise.
Temperature should either
approximate in situ sediment
temperature or 4° C, depending on
study objectives.
Interstitial water should
be analyzed as soon as
possible.
Store at 4° C with no
headspace or apply inert
gas in headspace for
toxicity studies.
Preserve with appropriate
preservative for chemical
analyses immediately.
Figure 6-3. Summary of recommended procedures and considerations for laboratory isolation of
interstitial water*
*Note: Emphasis should be placed on minimizing the duration of all sample manipulations whenever
possible
when interstitial water is stored for periods as short as 24 h (Hulbert and Brindle, 1975; Watson et
al., 1985; Kemble et al., 1999; Sarda and Burton, 1995; SETAC, 2001).
If sediments are anoxic, as most depositional sediments are, sample processing, including mixing of
interstitial water that has separated from the sediment, should be conducted in an inert atmosphere or
with minimal atmospheric contact. Exposure to air can result in oxidation of contaminants, thereby
altering bioavailability (Bray et al., 1973; Lyons et al., 1979; Howes et al., 1985). Air exposure can
also result in loss of volatile sulfides, which might increase the availability of sulfide-bound metals
(Allen et al., 1993; Bufflap and Allen, 1995). In addition, iron and manganese oxyhydroxides are
quickly formed upon exposure to air. These compounds readily complex with trace metals, thus
Chapter 6: Collection of Interstitial Water
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altering metals-related toxicity (Bray et al., 1973; Troup et al., 1974; Burton, 1991; Bufflap and
Allen, 1995). Maintaining anoxic processing conditions is not necessary when study objectives are
concerned with exposures to aerobic sediments, or if target contaminants are unaffected by oxidation
in short-term toxicity or bioaccumulation testing.
Interstitial water filtration should be avoided (SETAC, 2001). Numerous studies have shown that
filters reduce toxicity and contaminant concentrations by retaining contaminant-associated particles
and also by contaminant sorption onto the filter matrix (Bray et al., 1973; Troup et al., 1974; Sasson-
Brickson and Burton, 1991; Schults et al., 1992). If filtration is stipulated by a test method, treated
filters (e.g., pre-soaked in distilled, deionized water, or combusted at 400° C overnight for glass fiber
filters) should be used, and an unfiltered sample should also be tested for toxicity and contaminant
concentrations. The characteristics of filters and the filtering apparatus should also be carefully
considered, as different filters have different sorptive capacities for different contaminants.
Recommendation Box #2
Extraction of interstitial water
Centrifugation is the generally preferred laboratory method for the extraction of interstitial
water.
Extraction of interstitial water should be completed as soon as possible.
Interstitial water that has accumulated on the surface of the homogenized sediment
sample should be mixed into the sediment before the sample is partitioned among
centrifuge bottles.
Unless other program-specific guidance is available, sediments should be centrifuged at
high speed (e.g., 8000-10,000 x g) for 30 minutes.
Unless site-specific information suggests otherwise, centrifuging should be at 4° C to
minimize temperature-mediated biological and chemical processes.
Interstitial water should be preserved immediately for chemical analyses or analyzed as
soon as possible after extraction, unpreserved. For toxicity testing, interstitial water
should be stored at 4° C for not longer than 24 hours, unless the test method dictates
otherwise.
Filtration should be avoided unless required by a test method because it might reduce
interstitial water toxicity. Double (serial) centrif ugation (low speed followed by high speed)
should be used instead.
If filtering is required by a test method, pre-treated filters should be used to reduce
potential contamination.
6.3.2 Centrifugation
Centrifugation is the generally preferred laboratory method for collection of interstitial water
(SETAC, 2001). It is a relatively simple procedure that allows rapid collection of large volumes of
interstitial water. It also facilitates the maintenance of anoxic conditions (if required). However,
Centrifugation, like other ex-situ procedures might yield chemical and/or toxicological artifacts due to
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the extraction procedures themselves, which might alter the natural equilibrium between interstitial
water and sediment.
Prior to centrifugation, the sediment sample is homogenized (see Section 4.3) and partitioned among
centrifuge bottles. If the homogenized sample is stored prior to centrifugation, interstitial water
might accumulate on the surface of the sediment. This overlying water should be mixed into the
sediment before subsampling for centrifugation. Samples are then partitioned among centrifuge
bottles. In general, approximately 50% of sediment moisture content can be extracted as interstitial
water. If interstitial water volume requirements are lower, smaller sediment subsamples may be
used.
For more information about centrifugation:
Interstitial waters have been isolated over a range of centrifugal forces and durations (Landrum
etal., 1987; Giesyetal., 1988; Schults et al., 1992; Burgess etal., 1993; Ankley et al., 1990;
Schubauer-Berigan and Ankley, 1991; Ankley and Schubauer-Berigan, 1994). For toxicity
testing of interstitial waters, some sources recommend that sediments be centrifuged at 10,000
x g for a 30 min period (ASTM, 2000a; Environment Canada, 1994). Such high speed
centrifugation is often necessary to remove most colloids and dispersible clays (Adams, 1991;
Chin and Gschwend, 1991; Brownawell and Farrington, 1986; Ankley and Schubauer-Berigan,
1994), which can introduce interferences to chemical or toxicological analysis. However, such
high speed centrifuges are not commonly available. Furthermore, many materials (glass,
plastic) are not able to withstand high centrifugation speeds. Finally, it should be noted that
toxicity is typically reduced with high speed centrifugation due to the removal of particle-
associated contaminants (Sasson-Brickson and Burton 1991; Schults et al., 1992; Ankley and
Schubauer-Berigan, 1994; Buff lap and Allen, 1995).
Based on research to date, both slower and faster centrifugation speeds (and associated differences in
colloid/suspended solids removal) may be appropriate depending on the study objectives. For many
programs that are interested in characterizing site toxicity, high speed centrifugation may not be
appropriate because one is interested in toxicity potential of the interstitial water in its entirety (i.e.,
including colloidal material). However, if one is interested in comparing interstitial water
contaminant concentrations to specific sediment quality values, or model exposure compartments for
example (EPRI, 2000), then high speed centrifugation might be necessary. As our knowledge is still
limited in this area, it is perhaps most important to note that centrifugation speed often has a dramatic
effect on observed sample toxicity and chemical characteristics. Therefore, in any sediment
monitoring study, one centrifugation protocol (including speed and time) should be identified and
used throughout for all samples.
Centrifugation has been performed at various temperatures. ASTM (2000a) recommends that the
centrifugation temperature reflect the in situ sediment temperature to ensure that the equilibrium
between the particulate and interstitial water is not altered. Alternatively, a temperature of 4° C may
be preferred to minimize temperature-mediated chemical and biological processes (Environment
Canada, 1994).
When centrifuging coarse sand, it might be desirable to use a modified centrifuge bottle to aid
interstitial water recovery (USEPA/ACOE, 1998). The modified bottle is equipped with an internal
filter that can recover 75% of the interstitial water, as compared to 25 - 30% recovery from squeezing
(Saagerefa/., 1990).
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As discussed in Section 4.2, all containers have limitations with regards to adsorption or leaching of
chemicals, ease of use, and reliability. For example, polytetrafluororthylene (FTP) bottles have been
used successfully up to 2500 x g when filled to 80% of capacity, but collapse at 3000 g (Burgess et
al., 1993). Polycarbonate bottles have been used successfully for tributytin analyses in interstitial
water (Hoffman, 1998). If small volumes of water are required for testing, higher speed
centrifugation can be performed with glass tubes (up to 10,000 g, Word et al., 1987). Larger glass
tubes, however, can not be centrifuged at such high speeds. If metal toxicity is not a concern, then
high speed centrifugation in larger stainless steel centrifuge tubes is suitable. If test samples are
contaminated with photoreactive compounds such as PAHs, exposure of the sample to light should
be minimized to limit degradation or alteration of potentially toxic compounds. This can be
accomplished by using reduced lighting.
6.3.3 Sediment Squeezing
Isolation of interstitial water by squeezing has been performed using a variety of procedures and
devices (Reeburgh, 1967; Kalil and Goldhaker, 1973; Jahnke, 1988; Carr et al., 1989; Long et al.,
1990; Watson and Prickers, 1990; Adams, 1991; Carr and Chapman, 1995; Carr, 1998). Inexpensive
low pressure mechanical squeezers can be constructed, and may provide specialized capacities such
as collection of interstitial water profiles from core samples (Bender, et al, 1987). In all cases, the
interstitial water is passed through a filter that is a part of the squeezing apparatus.
Squeezing has been shown to produce a number of artifacts due to shifts in equilibrium from
pressure, temperature, and gradient changes (e.g., Froelich et al., 1979; Kriukov and Manheim, 1982;
Bellinger et al., 1992; Schults, 1992). Squeezing can affect the electrolyte concentration in the
interstitial water particularly with a decrease in chemical concentrations near the end of the
squeezing process. However, others reported that squeezing did not produce artifacts in interstitial
water toxicity studies (Carr and Chapman, 1995; Carr, 1998; SPTAC, 2001). It is therefore
recommended that if squeezing is performed, moderate pressures be applied along with electrolyte
(conductivity) monitoring during extraction (Kriukov and Manheim, 1982). Squeezing should also
be performed at in situ ambient temperatures, as significant alterations to interstitial water
composition can occur when squeezing is conducted at temperatures different from ambient
conditions (e.g., Mangelsdorf et al., 1969; Bischoff et al., 1970; Sayles et al., 1973).
Other sources of interstitial water alteration during squeezing are: contamination from overlying
water; internal mixing of interstitial water during extrusion; and solid-solution reactions as interstitial
water is expressed through the overlying sediment. As interstitial waters are displaced into upper
sediment zones, they come in contact with solids with which they are not in equilibrium. This inter-
mixing causes solid-solution reactions to occur. Most interstitial water chemical species are rapidly
transformed, as observed with ammonia and trace metals (Rosenfield, 1979; Santschi et al., 1997).
Bellinger et al. (1992) found elevated levels of several ions and dissolved organic carbon in squeezed
samples as compared to samples collected by in situ peepers. The magnitude of the artifact will
depend on the pollutant sediment characteristics and redox potential.
6.3.4 Pressurized and Vacuum Devices
Other methods for extraction of interstitial water from sediment samples can include vacuum
filtration (Jenne andZachara, 1987; Knezovich and Harrison, 1987; Winger andLasier, 1991), gas
pressurization (Reeburgh, 1967), and displacement (Adams, 1991). These methods typically recover
only small volumes of interstitial water and are not commonly used.
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Photo courtesy of Allen Burton
Sediment squeezing apparatus for extracting interstitial water
Chapter 6: Collection of Interstitial Water
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Use of a hand vacuum with an aquarium stone is an effective vacuum filtration method (Winger and
Lasier, 1991; Sarda and Burton, 1995). The procedure typically involves attaching the air stone to a
50 mL syringe via plastic tubing, inserting it into the sediment to the desired depth, and then
applying suction. This method can recover relatively large volumes of interstitial water; Santschi et
al. (1997) used this procedure to extract up to 1,500 mL from 4 L of sediment. Sarda and Burton
(1995) found that ammonia concentrations in water obtained by this procedure were similar to those
collected by in situ peepers. Drawbacks to this method include loss of equilibrium between the
interstitial water and the solids, filter clogging, and oxidation (Brinkman et al., 1982).
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Quality Assurance and Quality
Control
7.1 General Procedures
Quality assurance activities provide a formalized system for evaluating the technical adequacy of
sample collection and laboratory analysis activities. These quality assurance activities begin before
samples are collected and continue after laboratory analyses are completed, requiring ongoing
coordination and oversight. The
quality assurance program should
integrate management and technical
practices into a single system to
Checklist
QA practices within a
laboratory should address all
activities that affect the quality
of the final data, such as
sediment sampling and handling
condition and operation of equipment
instrument calibration
replication
use of standards
record keeping
data evaluation
provide data that are sufficient,
appropriate, and of known and
documented quality.
Developing and maintaining a quality
assurance (QA) program requires an
ongoing commitment by project
management and also includes the
following: (1) appointment of a
quality assurance officer with the
responsibility and authority to develop
and maintain a QA program,
(2) preparation of a Quality Assurance
Project Plan with Data Quality
Objectives, (3) preparation of written
descriptions of Standard Operating
Procedures (SOPs) for sediment
sampling and manipulations, instrument calibration, sample chain-of -custody, laboratory sample
tracking system, and (4) provision of adequate, qualified technical staff and suitable space and
equipment to assure reliable data. Program specific guidance for developing and maintaining a QA
program should be followed as appropriate. Examples of program guidance for developing a quality
assurance program can be found in USEPA (1994; 1995; 2000d), PSEP (1997a), WDE (1995), and
USEPA/ACOE (1991, 1998).
Quality control (QC) practices consist of more focused, routine, day-to-day activities carried out
within the scope of the overall QA program. QC is the routine application of procedures for
obtaining data that are accurate (precise and unbiased), representative, comparable, and complete.
QC procedures include activities such as identification of sampling and analytical methods,
calibration and standardization, and sample custody and record keeping. Audits, reviews, and
complete and thorough documentation are used to verify compliance with predefined QC procedures.
Project-specific QA plans (QAPP; see Section 7.3 below) provide a detailed plan for activities
performed at each stage of the study and outline the data quality objectives that should be achieved.
Through periodic reporting, QA activities provide a means to track progress and milestones,
performance of measurement systems, and data quality. A complete project-specific QA/QC effort
has two major components: a QA program implemented by the responsible agency (i.e., the data
Chapter 7: Quality Assurance and Quality Control
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
user) and QC programs implements by the parties responsible for collection and analyses (i.e., the
data generators).
7.2 QA/QC Procedures for Sediment Collection and Manipulation
Checklist
QA/QC procedures for
sample collection should
include the following
principal elements:
implementing a sound sampling approach
based on the intended use of the data.
use of sampling methodologies which allow
the collection of representative samples
based upon data needs.
use of sampling devices that minimize the
disturbance or alteration to the media's
chemical composition.
employing decontamination procedures
which reduce cross-contamination potential
between sampling points.
use of proper sample containers and
preservation techniques that maximize the
integrity of samples.
To ensure the appropriateness of the
sample collection protocol for sample
integrity and data of suitable quality, a
program of scheduled field QC samples,
such as field replicates (duplicates, splits,
field spikes), field blanks (rinsate
equipment), bottle, trip, and background
(upgradient) samples is critical. All field
QC samples should be handled exactly as
the sediment samples and should be
treated as blind samples so as to
minimize bias in the analysis. A random
portion of the samples should also be
analyzed by a third party to evaluate the
primary laboratory's performance. QC
replicates (duplicates, splits) should be
collected for analysis by the primary
laboratory to determine analytical
variability (USEPA 1995).
The procedures for sediment
manipulations described in Chapter 4
should maintain the sample in a chemical
condition as similar as possible to that at
the time of collection. QA procedures
are established to assure that SOPs are followed and that contamination is neither introduced to nor
lost from the manipulated sample. For example, samples to be analyzed for trace metals should not
come in contact with metal surfaces (except stainless steel). Sample tracking sheets should document
date, time, and investigator related to removal and replacement of samples from storage. Specific
manipulation procedures should follow established SOPs that minimize chemical alteration of the
sample (excepting chemical spiking), maintain sediment physical properties, and include replication
and blank samples.
7.3 The Quality Assurance Project Plan (QAPP)
The Quality Assurance Project Plan (QAPP) is a project-specific document that specifies the data
quality and quantity requirements needed for the study as well as all procedures that will be used to
collect, analyze, and report those data.
The QAPP uses input from the sampling design derived from the Data Quality Objectives Process
(see Chapter 2 specifically Measurement Quality Objectives discussion, Section 2.4, and USEPA,
2000a) to specify the above elements. This Plan should be reviewed by an independent person (e.g.,
quality assurance officer or staff member not involved in the project directly) for accuracy and
completeness. A key element of a QAPP is Standard Operating Procedures (see Section 7.4).
Further information on preparing a QAPP and resources necessary can be found in USEPA (2000e).
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7.4 Standard Operating Procedures
Standard operating procedures are written descriptions of routine methods and should be provided
for all methods used. A large number of field and laboratory operations can be standardized and
presented as standard operating procedures. General types of procedures that benefit from standard
operating procedures include field measurements ancillary to sample collection (e.g., water quality
measurements or mixing model input measurements); chain-of-custody, sample handling, and
shipment; and routine analytical methods for chemical analyses and toxicological analyses. Standard
operating procedures ensure that all persons conducting work are following the same procedures and
that the procedures do not change over time. All personnel should be thoroughly familiar with the
standard operating procedures before work is initiated. Deviations from standard operating
procedures might affect data quality and integrity. If it is necessary to deviate from approved
standard operating procedures, these deviations must be documented and approved through an
appropriate chain-of-command.
7.5 Sediment Sample Documentation
Bound field logbooks should be used for the maintenance of field records. All entries should be
dated and time of entry recorded. All aspects of sample collection and handling as well as visual
observations should be documented in the field logbooks. Documentation should be recorded in pre-
numbered bound notebooks using indelible ink pens in sufficient detail so that decision logic may be
traced back, once reviewed.
Checklist
Quality Assurance Project Plans vary in content depending on
program needs, but should address the following elements:
/ a description of the project organization and responsibilities
/ definition of data quality objectives (see Section 2.1)
/ sampling, analysis, and measurement procedures
J instrument calibration procedures
J procedures for recording, reducing, validating, and reporting data
J procedures for performing quality assurance verification and internal quality control
checks
J preventive maintenance schedules
J specific routine procedures to evaluate precision, accuracy, and completeness
J definitions of deviations and appropriate corrective actions
/ information on appropriate training
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Proper field sheet, sample labeling, chain-of-custody, and sample tracking documentation should be
maintained as appropriate. Specific details concerning sample documentation and sample
management should be included in planning documents and reviewed by the sampling team prior to
initializing the sampling program.
7.6 Sample Tracking Documentation
Samples delivered to the laboratory should be accompanied by a chain-of-custody record that
includes the name of the study, location of collection, date and time of collection, type of sample,
sample name or number, number of containers, analysis required, and the collector's signatures.
When turning over possession of samples, the relinquisher and the receiver sign, date and record the
time on the record sheet. The record sheet allows the transfer of a group of samples at one time.
When the laboratory takes possession of the samples, each should be assigned a unique laboratory
identification designation. This assures a consistent system for tracking within the laboratory. If the
samples arrive at the laboratory when designated personnel are not there to receive them, the samples
are put into a secure location and the transfer is conducted when the appropriate personnel are
present.
Upon arrival at the laboratory, samples are
inspected for condition and temperature, and
sample container labels are verified against
the chain-of-custody record or sample
tracking form. Sample information is
entered on a laboratory log-in data sheets
used to maintain information regarding
sample: receipt, shipping, collection date,
and storage. To allow for accurate
identification of samples, information
contained on sample tracking forms must
match identically with information
contained on the sample container labels.
The tracking form lists both the collector's
and the laboratory's identification
designations. Verified tracking forms are
signed by the laboratory personnel with date
and time in ink. Missing and/or
compromised samples (e.g., inappropriate
preservation to maintain integrity,
inappropriate containers, and unlabeled or
mislabeled containers) are documented on
the tracking forms.
When samples are removed from storage,
the sample tracking form accompanies it and
documents data, time, and investigator
associated with any manipulations. The
manipulation type is noted on the form in
detail or by reference to an approved
laboratory SOP. Any deviation from the
SOP are also noted. Should the sample be
Checklist
Sample documentation
should include:
project name, and analysis or test to be
performed
sampling locations
dates and times
sampling personnel present
level of personal protective equipment
worn
weather or any environmental condition
that might affect samples
equipment used to collect samples, and
sample container preparation
calibration data
deviations from approved work plans or
SOPs
sketch of sampling area
notation of the system identifying and
tracking samples
notation of any visitors to the site
initials and date on each page
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modified in such a way that additional subsamples are created, additional tracking forms must also be
created.
7.7 Record Keeping
Proper record keeping is essential to the scientific defensibility of a sediment sampling and
manipulation program. A separate file should be maintained for each sampling/manipulation event
or closely related events. This file should contain field logs, chain-of-custody forms, sample tracking
forms, storage records, and any QA/QC documentation and records. Original documentation should
be signed and dated by the originator.
7.8 QA Audits
In addition to the QA/QC procedures conducted on
a routine basis, quality audits (i.e., performance and
quality systems audits) might be conducted.
Performance audits refer to independent checks to
evaluate the quality of data produced during testing.
There are three types of performance audits:
sampling; test; and data processing. These audits
are independent of normal quality control checks
performed by the operator.
A systems audit is an on-site inspection and review
of the quality assurance system. The systems audit is
performed to verify that the organization is
following the policies and procedures described in
its QA/QC plan and in appropriate SOPs. Systems
audits are performed by an auditor typically from an
accrediting body.
Checklist
Performance
auditing procedures
are:
/ sample auditing - the auditor
uses a separate set of calibrated
standards to check the sample
collection system.
J test auditing - the auditor is
provided with set of a duplicate
sample or split portion.
/ data processing audit - the
auditor spot checks calculations
or a dummy set of raw data is
inserted followed by review of
validated data.
7.9 Corrective Action (Management of Non-conformance Events)
The QA Officer and the responsible manager are responsible for reviewing the circumstances of all
instances of occurrence of nonconformities, to determine whether corrective action should be taken.
The manager is responsible for determining if new samples are required, if the customer should be
notified, if additional testing is necessary, or whether the results should be confirmed. A good
communication plan is invaluable in helping to identify interactions among labs, clients, and agencies
during corrective actions.
Corrective action might take two forms: that of addressing technical problems associated with
project activities and that of addressing QA/QC infractions based upon performance. Technical
problems in meeting project objectives may range in magnitude from failure to meet minor
procedural requirements, to major problems associated with inappropriate methods or data loss.
Established procedures for corrective action of minor technical problems are often included in the
SOPs for cases where performance limits or acceptance criteria have been exceeded. On-the-spot
corrective actions are noted on data sheets. Major or recurrent QA/QC problems which require long-
term corrective action, such as modification of SOPs, are reported. Depending upon the nature and
Chapter 7: Quality Assurance and Quality Control
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
severity of the problem, an approach might be developed. Any corrective action is documented by
management.
Infractions of QA/QC policies by staff are identified and addressed by the management. Minor
infractions are corrected through additional training and/or closer supervision. Major or recurrent
infractions are corrected through re-assignment of technical personnel.
Corrective actions relative to sample collection and manipulation may include, but are not limited to,
review of the data and calculations, flagging and/or qualification of suspect data, or possible re-
sampling. A review that provides a preliminary check of all "out of limit" events is performed as
soon as the data for a given parameter or test is tabulated and verified for accuracy. "Out of limit"
events are flagged to determine whether new samples are required.
7.10 Data Reporting
In addition to reporting the raw data from a given sediment quality study or analysis, the data report
should include additional quality assurance information to ensure the data user that sample handling
and analyses are in accordance with the project plan. The quality assurance information also
documents procedures taken to ensure accurate data collection. Data are to be presented
electronically as well as in hardcopy for many regulatory programs. Required electronic format
should be explicitly outlined as a data quality objective during the planning process.
Checklist
Quality Assurance Reporting
/ A copy of the sample chain of custody record, including documentation of sample
collection date and time
/ Documentation of the laboratory certification number
/ Documentation of the analysis method used
J Documentation of analysis date and time (or testing period in the case of toxicity tests)
J Documentation that data for spikes, duplicates, standards, etc meets laboratory QA/QC
requirements for chemical analytes
J Documentation that reference toxicant test data meets laboratory QA/QC requirements
for toxicity tests.
/ Documentation of any deviations in sample preparation or analysis protocols
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References
Adams, D.D. 1991. Sampling Sediment Pore Water In: CRC Handbook of Techniques for Sediment
Sampling. Mudroch, A and MacKnight, S.D. (eds.). CRC Press, Inc., Boca Raton, FL.
Adams, W.U., R.A. Kimerle, and R.G. Mosher. 1985. Aquatic safety assessment of chemicals sorbed
to sediments. Aquatic Toxicol. and Hazard Assessment, Seventh Symposium, ASTM STP 854,
ASTM, Philadelphia, PA. pp. 429-453.
Alldredge, J.R. 1987. Sample size for monitoring of toxic chemical sites. Environmental Monitoring
and Assessment. 9:143-154.
Allen, T., 1975. Particle Size Measurement. John Wiley & Sons, New York, 452 pp.
Allen, H.E., G. Fu and B. Deng. 1993. Analysis of acid-volatile sulfide (AVS) and simultaneously
extracted metals (SEM) for the estimation of potential toxicity in aquatic sediments. Environmental
Toxicology and Chemistry 12:1441-1453.
American Public Health Association (APHA). 1995. Standard Methods for the Examination of Water
and Wastewater. 18th ed, Washington, D.C.
ASTM. 2000a. E 1391 - 94 Standard guide for collection, storage, characterization, and manipulation
of sediments for toxicological testing, p. 768-788. In: 2000 ASTM Standards on Environmental
Sampling, Vol. 11.05 Conshohocken, PA.
ASTM. 2000b. D4687 Guide for general planning of waste sampling. In 2000 ASTM Standards on
Environmental Sampling, Vol. 11.04 Conshohocken, PA.
ASTM. 2000c. D4387. Standard guide for selecting grab sampling devices for collecting benthic
macroinvertebrates. In: 2000 ASTM Standards on Environmental Sampling, Vol. 11.05
Conshohocken, PA.
ASTM. 2000d. D4823-95. Guide for core-sampling submerged, unconsolidated sediments. In: 2000
ASTM Standards on Environmental Sampling, Vol. 11.05 Conshohocken, PA.
ASTM. 2000e. D3976-92 Standard practice for preparation of sediment samples for chemical
analysis, pp.163-165, In: 2000 ASTM Standards on Environmental Sampling, Conshohocken, PA.
ASTM. 2000f. E729-96 Standard guide for conducting acute toxicity tests with fishes,
macroinvertebrates, and amphipods. p. 218-238. In: 2000 Annual Book of ASTM Standards, Vol.
11.05, Conshohocken, PA.
ASTM. 2000g. D1426-93 Standard test methods for ammonia nitrogen in water, pp. 151-155. In:
2000 ASTM Standards on Environmental Sampling, Vol. 11.04 Conshohocken, PA.
Chapter 8: References 8-1
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
ASTM. 2000h. D1067-92 Standard test methods for acidity and alkalinity of water, pp. 82.88. In:
2000 ASTM Standards on Environmental Sampling, Vol. 11.04 Conshohocken, PA.
ASTM. 20001. Dl 126-92 Standard test method for hardness in water, pp. 107-109. In: 2000 ASTM
Standards on Environmental Sampling, Vol. 11.04 Conshohocken, PA.
Ankley, G.T. and Schubauer-Berigan. 1994. Comparison of techniques for the isolation of pore water
for sediment toxicity testing. Archives of Environmental Contamination & Toxicology 27:507-512.
Ankley, G.T., A. Katko and J.W. Arthur. 1990. Identification of ammonia as a major sediment-
associated toxicant in the lower Fox River and Green Bay, Wisconsin. Environmental Toxicology
and Chemistry 9:313-322.
Ankley, G.T., M.K. Schubauer-Berigan, and J.R. Dierkes. 1991. Predicting the toxicity of bulk
sediments to aquatic organisms with aqueous test fractions: Pore water vs. elutriate. Environmental
Toxicology and Chemistry 10:925-939.
Ankley, G.T., S.A. Collyard, P.D. Monson, and P.A. Kosian. 1994. Influence of ultraviolet light on
the toxicity of sediments contaminated with polycyclic aromatic hydrocarbons. Environmental
Toxicology and Chemistry 11:1791-1796.
Ankley, G.T., D.M. DiToro, D.J. Hansen, and W.J. Berry. 1996. Technical basis and proposal for
deriving sediment quality criteria for metals. Environmental Toxicology and Chemistry 15: 2056-
2066.
Axelman, J., N. Carina, D. Broman, and N. Kristoffer, 1999: Accumulation of polycyclic aromatic
hydrocarbons in semipermeable membrane devices and caged mussels (Mytilus edulis 1.) In relation
to water column phase distribution. Environmental Toxicology and Chemistry 18(11):2454-2461
Barbour, M.T., J. Gerritsen, B.D. Snyder, J.B. Stribling. 1999. Rapid bioassessment protocols for use
in streams and wadeable rivers: periphyton, benthic macroinvertebrates, and fish, 2nd ed. EPA 841-B-
99-002. U.S. Environmental Protection Agency, Office of Water, Washington, D.C.
Barth, D.E. and T. Starks. 1985. Sediment sampling quality assurance user's guide. Prepared for
Environmental Monitoring Systems Laboratory, U.S. Environmental Protection Agency, 99 p.
Bascomb, C.L. 1964. Rapid method for the determination of cation exchange capacity of calcareous
and non-calcareous soils. /. Sci. FoodAgric. 15: 821-823.
Bates, RG. 1981. The modern meaning of pH. CRC Critical Reviews in Analytical Chemistry 10:247.
Baudo, R. 1990. Sediment Sampling, Mapping and Data Analysis. In: J.P. Giesy and H. Muntau
(eds.), Sediments: Chemistry and Toxicity of In-Place Pollutants. Lewis Publishers, Inc., Chelsea,
MI, pp. 15-60.
Becker, D.S. and T.C. Ginn. 1990. Effects of sediment holding time on sediment toxicity. Prepared
by PTI Environmental Services, Inc. for the U.S. EPA, Region 10 Office of Puget Sound, Seattle,
WA. EGA 910/9-90-009.
Bender, M., W. Martin, J. Hess, F. Sayles, L. Ball, and C. Lambert. 1987. A whole-core squeezer for
interfacial pore-water sampling. Limnology and Oceanography 32:1214-1225.
8-2 US Environmental Protection Agency
-------
Technical Manual
Bennett ER, C.D. Metcalfe, and T.L. Metcalfe. 1996. Semi-permeable membrane devices (SPMDs)
for monitoring organic contaminants in the Otonabee River, Ontario. Chemosphere 33:363-375.
Berner, R.A. 1963. Electrode studies of hydrogen sulphide in marine sediments. Geochimica et
Cosmochimica Acta. 27:563.
Berry, W.J., M.G. Cantwell, P.A. Edwards, J.R. Serbst and D.J. Hansen. 1999. Predicting toxicity of
sediments spiked with silver. Environmental Toxicology and Chemistry 18:40-48.
Besser, J.M., J.A. Kubitz, C.G. Ingersoll, W.E. Braselton, and J.P. Giesy. 1995. Influences of copper
bioaccumulation, growth, and survival of the midge Chironomus tentaus in metal-contaminated
sediments. Journal of Aquatic Ecosystem Health 4:157-168.
Birge, W.J., Black, J., Westerman, S. and Francis, P. 1987. Toxicity of sediment-associated metals to
freshwater organisms: Biomonitoring procedures. In: Fate and Effects of Sediment-Bound
Chemicals, Aquatic Systems. Pergamon Press, NY, pp. 199-218.
Bischoff, J.L., R.E. Greer, and A.O. Luistro. 1970. Composition of interstitial waters of marine
sediments: Temperature of squeezing effect. Science 167:1245-1246.
Black, C. A. (ed.) 1965. Methods of Soil Analysis. American Society of Agronomy, Agronomy
Monograph No. 9, Madison, WI.
Boehm, P.O., D.S Page, E.S. Gilfillan, W.A Stubblefield,. and E.J Harner. 1995. Shoreline Ecology
Program for Prince William Sound, Alaska, Following the Exxon Valdez Oil Spill: Part 2-Chemistry
and Toxicology. Exxon Valdez Oil Spill: Fate and Effects in Alaskan Waters, ASTM STP 1219, Peter
G. Wells, James N. Butler, and Jane S. Hughes, Eds., American Society for Testing and Materials,
Philadelphia, PA.
Bellinger, R., H. Brandl, P. Hohener, K.W. Hanselmann, and R. Bachofen. 1992. Squeeze-water
analysis for the determination of microbial metabolites in lake sediments-comparison of methods.
Limnology and Oceanography 37:448-455.
Borga, P., T. Elowson and K. Liukko. 1996. Environmental loads from water-sprinkled softwood
timber: 2: influence of tree species and water characterizations on wastewater discharges.
Environmental Toxicology and Chemistry. 15: 1445-1454
Bottomly, E.Z. and I.E. Bayly. 1984. A sediment pore water sampler used in root zone studies of the
submerged macrophyte, Myriophyllum spicatum. Limnology and Oceanography 29:671-673.
Bower and Holm-Hansen. 1980. Canadian Journal of Fisheries and Aquatic Sciences 37:794-798.
Bray, J.T., O.P. Bricker, and B.N. Troup. 1973. Phosphate in interstitial waters of anoxic sediments:
Oxidation effects during sampling procedure. Science 180:1362-1364.
Bregnard, T. B-A., P Hohener, A. Haner and J. Zeyer. 1996. Degradation of weathered diesel fuel by
microorganisms from a contaminated aquifer in aerobic microcosms. Environmental Toxicology and
Chemistry 15:299-307.
Brinkman, A.G., W. van Raaphorst, and L. Lijklema. 1982. In situ sampling of interstitial water from
lake sediments. Hydrobiologia 92:659-663.
Chapter 8: References 8-3
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Brownawell, B.J., and J.W. Farrington. 1986. Biogeochemistry of PCBs in interstitial waters of a
coastal marine sediment. Geochimica et CosmochimicaActa. 50:157-169.
Brumbaugh, W.G., C.G. Ingersoll, N.E. Kemble, T.W. May and J.L. Zajicek. 1994. Chemical
characterization of sediments and pore water from the Upper Clark Fork River and Milltown
Reservoir, Montana. Environmental Toxicology and Chemistry 13:1971-1973.
Buchanan, J.B. 1984. Sediment analysis. In: Methods for the Study of Marine Benthos. N.A. Holme
and A.D. Mclntyre (eds). Blackwell Scientific Publications, Boston, MA, pp. 41-65.
Bufflap, W.E. and H.E. Allen. 1995. Sediment pore water collection methods: A review. Water
Research 29:165-177.
Burgess, R.M., M.G. Cantwell, M.C. Pelletier, K.T. Ho, J.R. Serbst, H.F. Cook, and A. Kuhn. 2000.
Development of a toxicity identification evaluation procedure for characterizing metal toxicity in
marine sediments. Environmental Toxicology and Chemistry 19(4):982-991.
Burgess, R.M. and R.A. McKinney. 1997. Effects of sediment homogenization on interstitial water
PCB geochemistry. Archives of Environmental Contamination & Toxicology 33:125-129.
Burgess, R.M. 1996. Enrichment of marine sediment colloids withpolychlorinatedbiphenyls: Trends
resulting from PCB solubility and chlorination. Environmental Science and Technology 30(8):2556-
2566.
Burgess, R.M., K.A. Schweitzer, R.A. McKinney and D.K. Phelps. 1993. Contaminated marine
sediments: water column and interstitial toxic effects. Environmental Toxicology and Chemistry
12:127-138.
Burton, G.A. Jr. 1991. Assessment of freshwater sediment toxicity. Environmental Toxicology and
Chemistry 10:1585-1627.
Burton, G.A. Jr. 1992. Sediment collection and processing: factors affecting realism. In: G.A.
Burton, Jr. (ed.), Sediment Toxicity Assessment. Lewis Publishers, Chelsea, MI, pp. 37-67.
Burton GA, Jr., Rowland CD, Greenberg MS, Lavoie DR, Nordstrom JF, Eggert LM. 2001. A tiered,
weight-of-evidence approach for evaluating aquatic ecosystems. Aquatic Ecosystem Health and
Management (in press).
Burton, G.A. Jr., C. Rowland, D. Lavoie, and N. Nordstrom. 1999. Assessment of In Situ Stressors
and Sediment Toxicity in the Lower Housatonic River. Final Report to R.F. Weston.,
Manchester, NH.
Burton, G.A., Jr., B.L. Stemmer, K.L. Winks, P.E. Ross, and L.C. Burnett. 1989. A multitrophic level
evaluation of sediment toxicity in Waukegan and Indiana Harbors. Environmental Toxicology and
Chemistry 8:1057-1066.
Cairns, M.A., A.V. Nebeker, J.H. Gakstatter, and W.L. Griffis. 1984. Toxicity of copper-spiked
sediments to freshwater invertebrates. Environmental Toxicology and Chemistry 3:435-445.
8-4 US Environmental Protection Agency
-------
Technical Manual
Call, D. J., N. Christine, T.P. Polkinghorne, L.T. Markee, D.L. Brooke, J.W. Geiger,. K. Gorsuch,
and N. Robillard, 1999: Silver toxicity to Chironomus tentans in two freshwater sediments.
Environmental Toxicology and Chemistry 18(l):30-39.
Carignan, R. 1984. Interstitial water sampling by dialysis: Methodological notes. Limnology and
Oceanography 29:667-670.
Carignan, R. and D.R.S. Lean. 1991. Regeneration of dissolved substances in a seasonally anoxic
lake: The relative importance of processes occurring in the water column and in the sediments.
Limnology and Oceanography 36:683-703.
Carignan, R., F. Rapin, and A. Tessier. 1985. Sediment Pore water sampling for metal analysis: A
comparison of techniques. Geochimica et Cosmochimica Acta. 49:2493-2497.
Carignan, R., S. St. Pierre, and R. Gachter. 1994. Use of diffusion samplers in oligotrophic lake
sediments: Effects of free oxygen in sampler material. Limnology and Oceanography 39:468-474.
Carlton, R.G., and R.G. Wetzel. 1985. A box corer for studying metabolism of epipelic
microorganisms in sediment under in situ conditions. Limnology and Oceanography 30:422.
Carr, R.S. 1998. Marine and estuarine porewater toxicity testing. In: Wells, PG, K. Lee, C. Blaise,
eds. Microscale testing in aquatic toxicology: advances, techniques, and practice. CRC Press, Boca
Raton, FL., p. 523-538.
Carr, R.S. and D.C. Chapman. 1992. Comparison of solid-phase and pore-water approaches for
assessing the quality of marine and estuarine sediments. Chemical Ecology 7:19-30.
Carr, R.S. and D.C. Chapman. 1995. Comparison of methods for conducting marine and estuarine
sediment porewater toxicity tests - Extraction, storage, and handling techniques. Archives of
Environmental Contamination & Toxicology 28:69-77.
Carr, R.S., J.W. Williams, and C.T.B. Fragata. 1989. Development and evaluation of a novel marine
sediment pore water toxicity test with the polychaete Dinophilus gyrociliatus. Environmental
Toxicology and Chemistry 8:533-543.
Chao, T.T. and L. Zhou. 1983. Extraction techniques for selective dissolution of amorphous iron
oxides from soils and sediments. Soil Science Society of America Journal 47:225-232.
Chin, Y., and P.M. Gschwend. 1991. The abundance, distribution, and configuration of pore water
organic colloids in recent sediment. Geochimica et Cosmochimica Acta. 55:1309-1317.
Clark, J.R., J.M. Patrick, J.C. Moore, and J. Forester. 1986. Accumulation of sediment-bound PCBs
by fiddler crabs. Bulletin of Environmental Contamination and Toxicology 36:571-578.
Clark, J.R., J.M. Patrick, Jr., J.C. Moore, and E.M. Lores. 1987. Waterborne and sediment-source
toxicities of six organic chemicals to grass shrimp (Palaemonetes pugio) and Amphioxus
(Branchiostoma caribaeum). Archives of Environmental Contamination & Toxicology 16:401-407.
Cole, F.A., B.L. Boese, R.C. Schwatz, J.D. Lanberson and T.H. Dewit. 2000. Effects of storage on
the toxicity of sediments spiked with fluoranthene to the amphipod, Rhepoxyinius abronius.
Environmental Toxicology and Chemistry 19(3):744-748.
Chapter 8: References 8-5
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Crecelius, E.A., E.A. Jenne and J.S. Anthony. 1987. Sediment quality criteria for metals:
optimization of extraction methods for determining the quantity of sorbents and adsorbed metals in
sediments. Report prepared by Battelle for U.S. Environmental Protection Agency, Criteria and
Standards Division, Washington, D.C.
Crevello,, P.D., J.M. Rine, and D.E. Lanesky. 1981. A method for impregnating unconsolidated cores
and slabs of calcareous and terrigenous muds. Journal of Sediment Petrology 51:658-660.
Grassland, N.O., and C.J.M. Wolff. 1985. Fate and biological effects of pentachlorophenol in
outdoor ponds. Environmental Toxicology and Chemistry 4:73-86.
Daniels, S.A., M. Munawar, and C.I. Mayfield. 1989. An improved elutriation technique for the
bioassessment of sediment contaminants. Hydrobiologia 188/189:619-631.
Day, P. R. 1965. Particle Fractionation and Particle-Size Analysis, pp. 562-566 In: Hydrometer
Method of Particle Size Analysis. Monograph No. 9 American Society of Agronomy, Madison, WI.
Day, K.E., R.S. Kirby, and T.B. Reynoldson. 1995. The effect of manipulations on freshwater
sediments on responses of benthic invertebrates in whole-sediment toxicity tests. Environmental
Toxicology and Chemistry 14:1333-1343.
Defoe, D.L and G.T. Ankley. 1998. Influence of storage time on toxicity of freshwater sediments to
benthic macroinvertebrates. Environmental Pollution In Press.
Dewitt, T.H., R.J. Ozretich, R.C. Swartz, J.O. Lamberson, D.W. Schults, G.R. Ditsworth, J.K. Jones,
L. Hoselton, and L.M. Smith. 1992. The influence of organic matter quality on the toxicity and
partitioning of sediment-associated fluoranthene. Environmental Toxicology and Chemistry
11(2): 197-208.
Dewitt, T.H., R.C. Swartz, and J.O. Lamberson. 1989. Measuring the acute toxicity of estuarine
sediments. Environmental Toxicology and Chemistry 8:1035-1048.
Diamond, J., A. Richardson, and C. Daley. 1999. Ecological effects of sediment-associated
contaminants in inner Burlington Harbor, Lake Champlain. In: T. Manley and P. Manley (eds.). Lake
Champlain in Transition: From Research Toward Restoration. American Geophysical Union,
Washington, D.C. pp. 261-276.
Dillon, T.M., D.W. Moore, and A.S. Jarvis. 1994. The effects of storage temperature and time on
sediment toxicity. Archives of Environmental Contamination & Toxicology 27:51-53.
Di Toro, D.M., Mahony, J.H., Hansen, D.J., Scott, K.J., Hicks, M.B., Mayr, S.M., and Redmond, M.
1990. Toxicity of cadmium in sediments: The role of acid volatile sulfides. Environmental
Toxicology and Chemistry 9:1487-1502.
Di Toro, D.M., C.S. Zarba, D.J. Hansen, W.J. Berry, R.C. Swartz, C.E. Cowan, S.P. Pavlou, H.E.
Allen, N.A. Thomas, and P.R. Paquin. 1991. Technical basis for establishing sediment quality
criteria for nonionic organic chemicals using equilibrium partitioning. Environmental Toxicology and
Chemistry 10:1541-1583.
Ditsworth G.R., D.W. Schults, and J.K.P. Jones. 1990. Preparation of benthic substrates for sediment
toxicity testing. Environmental Toxicology and Chemistry 9:1523-1529.
8-6 US Environmental Protection Agency
-------
Technical Manual
Doig, P., K. Liber. 2000. Dialysis minipeeper for measuring porewater metal concentrations in
laboratory sediment toxicity and bioavailability tests. Environmental Toxicology and Chemistry
19:2882-2889.
Driscoll SK and P.P. Pandrum. 1997. A comparison of equilibrium partitioning and critical body
residue approaches for predicting toxicity of sediment-associated fluoranthene to freshwater
amphipods. Environmental Toxicology and Chemistry 16:2179-2186
Duncan, G. A., and G.G. Pattaie. 1979. Size Analysis Procedures Used in the Sedimentology
Paboratory, NWRI Manual. National Water Research Institute, Canada Centre for Inland Waters.
Pnvironment Canada. 1994. Guidance document on collection and preparation of sediments for
physicochemical characterization and biological testing. Pnvironmental Protection Series. Report
PPS l/RM/29, December 1994, 132 pp.
Pnvironment Canada. 1995. Guidance Document on Measurement of Toxicity Test Precision Using
Control Sediments Spiked with a Reference Toxicant. Report PPS l/RM/30.
Plectrical Power Research Institute (PPRI). 1986. Speciation of selenium and arsenic in natural
waters and sediments. Prepared by Battelle Pacific Northwest Paboratories. Vol. 2. PPRI PA-4641.
PPRI. 1999. Review of Sediment Removal and Remediation Technologies atMGP and Other
Contaminated Sites, PPRI, Palo Alto, CA, and Northeast Utilities, Berlin, CT: 1999. TR-113106.
PPRI. 2000. MARS model, Beta v.l, 2000.
Ferraro, S.P., F.A. Cole, W.A. DeBen, and R.C. Swartz. 1989. Power-cost efficiency of eight
macrobenthic sampling schemes in Puget Sound, Washington, USA. Canadian Journal of Fisheries
and Aquatic Sciences 46(10):2157-2165.
Ferraro, S.P., R.C. Swartz, F.A. Cole, and W.A. DeBen. 1994. Optimum macrobenthic sampling
protocol for detecting pollution impacts in the southern California Bight. Environmental Monitoring
and Assessment 29:127'-153.
Flegel, A.R., R.W. Risebrough, B. Anderson, J. Hunt, S. Anderson, J. Oliver, M. Stephenson, and R.
Pickard. 1994. San Francisco Estuary Pilot Regional Monitoring Program Sediment Studies, San
Francisco Bay Regional Water Quality Control Board/State Water Resources Control Board,
Oakland, CA.
Fleming, R. and S. Nixon. 1996. Sediment Toxicity Tests. Interim Report No. DoF, Contract 8864.
Department of Environment. Buckinghamshire, Fngland.
Folk, R.P. 1968. Petrology of sedimentary rocks. Hemphill Publishing Co., Austin, TX. 172 p.
Francis, P.C., W. Birge, and J. Black. 1984. Fffects of cadmium-enriched sediment on fish and
amphibian embryo-larval stages. Ecotoxicology and Environmental Safety 8:378-387.
Frazier, B.F., T.J. Naimo, and M.B. Sandheinrich. 1996. Temporal and vertical distribution of total
ammonia nitrogen and un-ionized ammonia nitrogen in sediment pore water from the upper
Mississippi River. Environmental Toxicology and Chemistry 15:92-99.
Chapter 8: References 8-7
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Fredette, T.J., J.E. Clausner, D.A. Nelson, E.B. Hands, T. Miller-Way, J.A. Adair, V.A. Sotler, and
F.J. Anders. 1990. "Selected Tools and Techniques for Physical and Biological Monitoring of
Aquatic Dredged Material Disposal Sites", Technical Report, U.S. Army Engineer Waterways
Experiment Station, Vicksburg, MS. D-90-11.
Froelich, P.M., G.P. Klinkhammer, M.L. Bender, N.A. Luedtke, G.R. Heath, D. Cullen, P. Dauphin,
D. Hammond, B. Hartmann, and V. Maynard. 1979. Early oxidation of organic matter in pelagic
sediments of the eastern equatorial Atlantic: Suboxic diagenesis. Geochimica et Cosmochimica Acta
43:1075-1090.
Gambrell, R.P, R.A. Khalid and W.H. Patrick, Jr. 1976. Physicochemical parameters that regulate
mobilization and immobilization of toxic heavy metals. Proceedings on Speciality Conference on
Dredging and Its Environmental Effects, Mobile, AL (New York: American Society of Civil
Engineering).
Gee, G.W. and J.W. Bauder. 1986. Particle-size analysis. In: A. Klute (ed.), Methods of Soil
Analysis, 2nd ed. Part 1. Physical and Mineralogical Methods. American Society of Agronomy,
Madison, WI, pp. 383-411.
Giesy, J.P., R.L. Graney, J.L. Newstead, C.J. Rosiu, A. Benda, R.G. Kreis, Jr, and F.J. Horvath.
1988. Comparison of three sediment bioassay methods using Detroit River sediments. Environmental
Toxicology and Chemistry 7:483-493.
Giesy, J.P., C.J. Rosiu, R.L Graney, and M.G. Henry. 1990. Benthic invertebrate bioassays with toxic
sediment and pore water. Environmental Toxicology and Chemistry 9:233-248.
Gilek, M., M. Bjork, D. Broman, N. Kautsky and C. Naf. 1996. Enhanced accumulation of PCB
congeners by Baltic Sea blue mussels, Mytilus edulis, with increased algae enrichment.
Environmental Toxicology and Chemistry 15:1597-1605.
Gilfillan, E.S., Page, D.S., Harner, E.J., Boehm, P.O., "Shoreline Ecology Program For Prince
William Sound, Alaska, Following the Exxon Valdez Oil Spill: Part 3-Biology," Exxon Valdez Oil
Spill: Fate and Effects in Alaskan Waters, ASTM STP 1219, Peter G. Wells, James N. Butler, and
Jane S. Hughes, Eds., American Society for Testing and Materials, Philadelphia, PA, 1995.
Gillfillan, E.S., Harner, E.J., O'Reilly, J.E., Page, D.S., and Burns, W.A. 1999. A Comparison of
Shoreline Assessment Study Designs Used for the Exxon Valdez Oil Spill. Elsevier Science Ltd.
pubs., Great Britain, 1999. Marine Pollution Bulletin 38(5):380-388.
Ginsburg, R.N., H.A. Bernard, R.A. Moody, and E.E. Daigle. 1966. The shell method of
impregnating cores of unconsolidated sediments. Journal of Sediment Petrology 36:1118-1125.
GLNPO 1994. Assessment and remediation of Contaminated Sediments (ARCS) Program.
Assessment Guidance Document. EPA 905-B94-002, Great Lakes National Program Office, Chicago,
111.
Golterman, H.L., P.G. Sly and R.L. Thomas. 1983. Study of the Relationship between Water Quality
and Sediment Transport, UNESCO, Mayenne, France.
8-8 US Environmental Protection Agency
-------
Technical Manual
Gonzalez. A.M. 1995. A laboratory formulated sediment incorporating synthetic acid volatile sulfide.
Abstr. PHI35, p. 310. Annu. Meet. Society for Environmental Toxicology and Chemistry,
Vancouver, B.C.
Green, R.H., "Power Analysis and Practical Strategies for Environmental Monitoring," Environ.
Res., 50: 195-205 (1989).
Guigne, J.Y., N. Rudavina, P.H. Hunt, and J.S. Ford. 1991. An acoustic parametric array for
measuring the thickness and stratigraphy of contaminated sediments. Journal of Great Lakes
Research 17(1): 120-131.
Gustafsson, O., F. Haghesta, C. Chan, J. MacFarlane, and P.M. Gschwend. 1997. Quantification of
the dilute sedimentary soot phase: Implication for PAH speciation and bioavailability. Environmental
Science and Technology 31(1):203-209.
Hakenson, L. 1984. Sediment sampling in different aquatic environments: Statistical Aspects. Water
Resource Research. 20:41-46.
Hedges, J.I. and J.H. Stern. 1984. Carbon and nitrogen determination of carbonate-containing solids.
Limnology and Oceanography 29:657-663.
Harkey, G.A, P.P. Landrum, and S.J. Kaine. 1994. Comparison of whole-sediment elutriate, and
pore-water exposures for use in assessing sediment-associated organic contaminants in bioassays.
Environmental Toxicology and Chemistry 13:1315-1329.
Hedges, J.I. and J.H. Stern. 1984. Carbon and nitrogen determination of carbonate-containing solids.
Limnology and Oceanography 29:657-663.
Hermann, R. 1996. The daily changing pattern of hydrogen peroxide in New Zealand surface waters.
Environmental Toxicology and Chemistry 15:652-662.
Hesslein, R.H. 1976. An in-situ sampler for close interval pore water studies. 21: 912-914.
Ho, K.T., A. Kuhn, M.C. Pelletier, R.M. Burgess, and A. Helmstetter. 1999. Use ofulva lactuca to
distinguish pH-dependent toxicants in marine waters and sediments. Environmental Toxicology and
Chemistry 18(2): 207-212.
Ho, K.T., R. McKinney, A. Kuhn, M. Pelletier, and R. Burgess. 1997. Identification of acute
toxicants in New Bedford Harbor sediments. Environmental Toxicology and Chemistry 16(3):551-
558.
Hoffman, E. 1998. Tributyltin analysis: Clarification of interstitial water extraction methods and
sample storage - Interim. DMMP Clarification Paper, Dredged Material Management Program,
USEPA Region 10, Seattle, WA.
Hoke, R. A., J. P. Giesy, G. R. Ankley, J. L. Newsted and J. Adams. 1990. Toxicity of sediments
from Western Lake Erie and Maumee River at Toledo, Ohio, 1987: implications for current dredged
material disposal practices. Journal of Great Lakes Research 16:457-470.
Holland, A.F. 1985. Long-term variation of macrobenthos in a mesohaline region of the Chesapeake
Bay. Estuaries 8(2a):93-113
Chapter 8: References 8-9
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Hoss, S., M. Haitzer, W. Traunspurger and C.E.W. Steinberg. 1999. Growth and fertility of
Caenorhabditis elegans (Nematoda) in unpolluted freshwater sediments: Response to particle size
distribution and organic content. Environmental Toxicology and Chemistry 18(12): 2921-2925.
Howes, B.L., J.W.H. Dacey, and S.G. Wakeham. 1985. Effects of sampling technique on
measurements of porewater constituents in salt marsh sediments. Limnology and Oceanography
30:221-227.
Huckins, J.N, M.Q. Ruvwefwn, F.K. Mnuqwwe. 1990. Semipermeable membrane devices containing
model lipid: a new approach to monitoring the bioavailability of lipophilic contaminants and
estimating their bioconcentration potential. Chemosphere 20:533-552.
Hulbert, M.H., and M.P. Brindel. 1975. Effects of sample handling on the composition of marine
sedimentary pore water. Geological Society of America, Bulletin 86:109-110.
Ingersoll, C.G. 1995. Sediment Tests. In: G.M. Rand (ed.), Fundamentals of Aquatic Toxicology, 2nd
Edition. Taylor and Francis, Washington, D.C. pp. 231-255.
Ingersoll, C.G. and M.K. Nelson. 1990. Testing sediment toxicity with Hyalella azteca (Amphipoda)
and Chironomis riparius (Diptera), pp. 93-109, In: Aquatic Toxicology and Risk Assessment, vol.
13, W.G. Landis and W.H. van der Schalie (Eds.) ASTM STP 1096, American Society for Testing
and Materials, Philadelphia, PA.
Ingersoll, C.G., D.R. Buckler, E.A. Crecelius, and T.W. La Point. 1993. U.S. Fish and Wildlife
Service and Battelle final report for the U.S. EPA GLNPO assessment and remediation of
contaminated sediment (ARCS) project: Biological assessment of contaminated Great Lakes
sediment. EPA-905-R93-006, Chicago, IL.
International Joint Commission (IJC). 1988. Procedures for the assessment of contaminated sediment
problems in the great lakes. IJC, Windsor, Ont, Canada, p. 140.
ITFM. 1995. The strategy for improving water quality monitoring in the United States. Final Report
of the Intergovernmental Task on Monitoring Water Quality, US Geological Survey, Office of Water
Data Coordination, Reston, VA, OFR 95-742.
Jackson, M. L. 1958. Soil Chemical Analysis. Prentice-Hall, Inc., Englewood Cliffs, NJ.
Jahnke, R.A. 1988. A simple, reliable, and inexpensive pore-water sampler. Limnology and
Oceanography 33:483-487.
Jenne, E.A. and J.M. Zachara. 1984. Factors influencing the sorption of metals. Fate and Effects of
Sediment-bound Chemicals in Aquatic Systems. Pergamon Press, NY pp 83-98.
Johns, D.M., R. Gutjahr-Gobell, and P. Schauer. 1985. Use of bioenergetics to investigate the impact
of dredged material on benthic species: A laboratory study with polychaetes and Black Rock Harbor
material. Field Verification Program. Prepared for EPA and USCOE, monitored by WES, Vicksburg,
MI.
Johns, D.M., R.A. Pastorok, and T.C. Ginn. 1991. A sublethal sediment toxicity test using juvenile
Neanthes sp. (Polychaeta: Nereidae), pp. 280-293, In: Aquatic Toxicology and Risk Assessment, vol.
8-10 US Environmental Protection Agency
-------
Technical Manual
14, M.A. Mayes and M.G. Barren (Eds.) ASTM STP 1124, American Society for Testing and
Materials. Philadelphia, PA.
Kalil, E.K., and M. Goldhaker. 1973. A sediment squeezer for removal of pore waters without air
contact. Journal of Sediment Petrology 43:554-557.
Kaplan, I., S.T. Lu, R.P. Lee and G. Warrick. 1996. Polycyclic hydrocarbon biomarkers confirm
selective incorporation of petroleum in soil and kangaroo rat liver samples near an oil well blowout
site in the western San Joaquin Valley, California. Environmental Toxicology and Chemistry 15:696-
707.
Karickhoff, S.W., and K.R. Morris. 1985a. Sorption dynamics of hydrophobic pollutants in sediment
suspensions. Environmental Toxicology and Chemistry 4:469-479.
Karickhoff S.W., and K.R. Morris. 1985b. Sorption of hydrophobic pollutants in natural sediments.
Analysis, Chemistry, Biology 2:193-205.
Keilty, T.J., D.S. White, and P.P. Landrum. 1988a. Short-term lethality and sediment avoidance
assays with endrin-contaminated sediment and two oligochaetes from Lake Michigan. Archives of
Environmental Contamination & Toxicology 17:95-101.
Keilty, T.J., D.S. White, and P.P. Landrum. 1988b. Sublethal responses to endrin in sediment by
Stylodrilius heringianus (Lumbriculidae) as measured by a 137Cesium marker layer technique.
Aquatic Toxicology 13:227-250.
Keith, L. H. 1993. Principles of Environmental Sampling. ACS Professional Reference Book,
American Chemical Society, 458 pp.
Kemble, N.E., J.M. Besserr, W.G. Brumbaugh, E.L. Brunson, T.J. Canfield, J.J. Coyle, F.J. Dwyer,
J.F. Fairchild, C.G. Ingersoll, T.W. La Point, J.C. Meadows, D.P. Mondo, B.C. Poulton, D.F.
Woodward, and J.L. Zajicek. 1993. Sediment toxicology, pp. 2-1 to 2-100, In: Effects of Metal-
Contaminated Sediment, Water, and Diet on Aquatic Organisms, C.G. Ingersoll, W.C. Brumbaugh,
A.M. Farag, T.W. La Point, and D.F. Woodward (Eds.), May 10, 1993, U.S. Environmental
Protection Agency, Helena, MT.
Kemble, N.E., W.G. Brumbaugh, E.L. Brenson, F.J. Dwyer, G. Ingersoll, D.P. Monda, and D.F.
Woodward. 1994. Toxicity of metal contaminated sediments from the Upper Clark Fork River
Mountain, to aquatic invertebrates in laboratory exposures. Environmental Toxicology and Chemistry
13:1985-1997.
Kemble, N. E., F. J. Dwyer, C. G. Ingersoll, T.D. Dawson, and T. J. Norberg-King, 1999. Tolerance
of freshwater test organisms to formulated sediments for use as control materials in whole-sediment
toxicity tests. Environmental Toxicology and Chemistry 18:222-230.
Kemp, A.L.W., H.A. Saville, C.B. Gray, and A. Mudrochova. 1971. A simple corer and method for
sampling the mud-water interface. Limnology and Oceanography 16:689.
Kersten, M. and U. Forstner. 1987. Cadmium associations in freshwater and marine sediment.
Cadmium in the Aquatic Environment, John Wiley & Sons, New York. pp. 51-88.
Chapter 8: References 8-11
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Klemm, D.J., P.A. Lewis, F. Fulk, and J.M. Lazorchak. 1990. Macroinvertebrate Field and
Laboratory Methods for Evaluating the Biological Integrity of Surface Waters. US Environmental
Protection Agency, EPA-600-4-90-030, Environmental Monitoring and Support Laboratory,
Cincinnati, OH
Klute, A. Ed. 1986. Methods of Soil Analysis. Part 1. Physical and Mineralogical Methods, 2nd ed.
American Society of Agronomy, Madison, WI, USA.
Knezovich, J.P., and F.L. Harrison. 1988. The bioavailability of sediment-sorbed chlorobenzenes to
larvae of the midge chironomus decorus. Ecotoxicology and Environmental Safety 15:226-241.
Knezovitch, J.P. and F.L. Harrison. 1987. A new method for determining the concentration of
volatile organic compounds in sediment interstitial water. Bulletin of Environmental Contamination
and Toxicology 38:937-940.
Kosian, P.A., C.W. West, M.S. Pasha, J.S. Cox, D.R. Mount, R.J. Huggett, and G.T. Ankley. 1999.
Use of nonpolar resin for reduction of fluoranthene bioavailability in sediment. Environmental
Toxicology and Chemistry 18(2):201-206.
Kratochvil, B. and J.K. Taylor, "Sampling for chemical Analysis," Anal. Chem., 53: 924A-938A
(1981).
Kristensen, E. and T.H. Blackburn. 1987. The fate of organic carbon and nitrogen in experimental
marine sediment systems: Influence of bioturbation and anoxia. Journal of Marine Research 45:231-
257.
Kriukov, P.A., and F.T. Manheim. 1982. Extractions and investigative techniques for study of
interstitial waters of unconsolidated sediments: A review. The Dynamic Environment of the Ocean
Floor., K.A. Fanning and F.T. Manheim (Eds.). Lexington Books, Washington D.C. pp. 3-26.
Landrum, P.F. and W.R. Faust. 1991. Effect of variation in sediment composition on the uptake rate
coefficient for selected PCB and PAH congeners by the amphipod, Diporeia sp., pp. 263-279, In:
Aquatic Toxicology and Risk Assessment, vol. 14, M.A. Mayes and M.G. Barren (Eds.) ASTM STP
1124, American Society for Testing and Materials. Philadelphia, PA.
Landrum, P.F., S.R. Nihart, B.J. Eadie, and L.R. Herche. 1987. Reduction in bioavailability of
organic contaminants to the amphipod Pontoporeia hoyi by dissolved organic matter of sediment
interstitial waters. Environmental Toxicology and Chemistry 6:11-20.
Landrum, P.F., B.J. Eadie, and W.R. Faust. 1992. Variation in the bioavailability of polycyclic
aromatic hydrocarbons to the amphipod Diporeia (spp.) with sediment aging. Environmental
Toxicology and Chemistry 11:1197-1208.
Leonard, E. 1991. Standard operating procedures for total organic carbon analysis of sediment
samples. U.S. Environmental Protection Agency, Office of Research and Development,
Environmental Research Laboratory, Duluth, MN.
Leonard, E.N., D.R. Mount, and G.T. Ankley. 1999. Modification of metal partitioning by
supplementing acid volatile sulfide in freshwater sediments. Environmental Toxicology and
Chemistry 18(5):858-864.
8-12 US Environmental Protection Agency
-------
Technical Manual
Leppard, G. G. 1986. The Fibrillar Matrix Component of Lacustrine Biofilms. Water Research
20:697-702.
Leppard, G. G., J. Buffle, R.R. De Vitore, and D. Pereet. 1988. The infrastructure and Physical
Characteristics of a Distinctive Colloidal Iron Particulate Isolated from a Small Eutrophic Lake.
Archives Hydrobiology 113:405-424.
Long, E.R., M.F. Buchman, S.M. Bay, R.J. Breteler, R.S. Carr, P.M. Chapman, J.E. Hose, A.L.
Lissner, J. Scott, and D.A. Wolfe. 1990. Comparative evaluation of five toxicity tests with sediments
from San Francisco Bay and Tomales Bay, California. Environmental Toxicology and Chemistry
9:1193-1214
Long, E.R., D.D. MacDonald, S.L. Smith, F.D. Calder. 1995. Incidence of adverse biological effects
within ranges of chemical concentrations in marine and estuarine sediments. Environmental
Management 19(l):81-97.
Long, E.R., A. Robertson, D.A. Wolfe, J. Hameedi, and G.M. Sloane. 1996. Estimates of the spatial
extent of sediment toxicity in major U.S. estuaries. Environmental Science and Technology 30:3585-
3592.
Lydy, M.J., K.A. Bruner, K.A. Fry, and S.W. Fisher. 1990. Effects of sediment and the route of
exposure on the toxicity and accumulation of neutral lipophilic and moderately water soluble
metabolizable compounds in the midge, Chironomus riparius, pp. 140-164, In: Aquatic Toxicology
and Risk Assessment, vol. 13, W.G. Landis and W.H. van der Schalie (Eds.) ASTM STP 1096,
American Society for Testing and Materials. Philadelphia, PA.
Lyons, W.B., J. Gaudette, and G. Smith. 1979. Pore water sampling in anoxic carbonate sediments:
Oxidation Artifacts. Nature 277:48-49.
MacDonald, L.H., A.W. Smart, and R.C. Wissmar. 1991. Monitoring Guidelines to Evaluate effects
of Forestry Activities on Streams in the Pacific Northwest and Alaska EPA 910/9-01-001. USEPA
Region 10, Seattle, WA.
MacLeod, W., Jr., D. Brown, A. Friedman, O. Maynes and R. Pierce. 1985. Standard analytical
procedures of the NOAA National Analytical Facility, 1984-85, extractable toxic organic
compounds. Prepared for the NOAA National Status and Trends Program. NOAA Technical
Memorandum NMFS F/NWC-64.
Mangelsdorf, P.C. and T.R.S. Wilson. 1969. Potassium enrichments in interstitial waters of recent
marine sediments. Science 165:171.
Mayer, L.M. 1976. Chemical water sampling in lakes and sediments with dialysis bags. Limnology
and Oceanography 21:909.
McCave, I. M., and Jarvis, J. 1973. Use of the Model T Coulter Counter in Size Analysis of Fine to
Coarse Sand. Sedimentology 20:305-315.
McLeese, D. W., Metcalfe, C. D., and Pezzack, D. S. 1980. Uptake of PCB's from Sediment by
Nereis virens and Crangon septemspinosa. Archives of Environmental Contamination & Toxicology
9:507-518.
Chapter 8: References 8-13
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Metro. 1981. (Revised 1983). Analytical support and data validity: organics. Prepared for Toxicant
Pretreatment Planning Study. Municipality of Metropolitan Seattle, Seattle, WA.
Milton, J.S., J.J. Corbet, and P.M. McTeer, Introduction to Statistics, D.C. Heath and Company,
Toronto, Ontario, 517 p. (1986)
Moore, D.W., T.M. Dillon, andE.W. Gamble. 1996. Long-term storage of sediments: Implications
for sediment toxicity testing. Environmental Pollution 89:341-342.
Morris, J.C. and W. Stumm. 1967. Redox equilibria and measurements in the aquatic environment.
Advances in Chemistry Series 67:270.
Mudroch, A. and S.D. MacKnight. 1994. CRC Handbook of Techniques for Aquatic Sediment
Sampling, 2nd Ed., CRC Press, Boca Raton, FL, 210 pp.
Mudroch, A. and J.M. Azcue. 1995. Manual of Aquatic Sediment Sampling. CRC/Lewis, Boca
Raton, FL.
Muir, D.C.G, N.D. Griff, B.E. Townsend, D.A. Matner, and W.L. Lockhart. 1982. Comparison of the
uptake and bioconcentration of fluridone and terbutryn by rainbow trout and Chironomus tentans in
sediment and water systems. Archives of Environmental Contamination & Toxicology 11:595-602.
Murray, J.W., V. Grunamanis and W.M. Smethie, Jr. 1978. Interstitial water chemistry in the
sediments of Saanich Inlet. Geochimica et Cosmochimica Acta. 42:1011-1026.
Myers, D.E. 1988. Some aspects of multivariate geostastical analysis. In: C. F. Chung (ed)
Quantitative Analysis of Mineral and Energy Resources, D. Reidel Publishing Co., Dordrecht,
Germany, pp 669-687
National Oceanic and Atmospheric Administration (NOAA). 1991. NOAA diving manual: diving for
science and technology. Prepared by U.S. Department of Commerce, NOAA, Office of Undersea
Research. Publication number VM981.U6228.
Nebecker, A.V., S.T. Onjukka, M.A. Cairns, and D.F. Kraweztk. 1986. Survival of Daphinia magna
and Hyalella azteca in cadmium spiked water. Environmental Toxicology and Chemistry 5:933-938.
Nelson, M.K., P.F. Landrum, G.A. Burton, Jr., S.J. Klaine, E.A. Crecelius, T.D. Byl, D.C. Gossiaux,
V.N. Tsymbal, L. Cleveland, C.G. Ingersoll, and G. Sasson-Brickson. 1993. Toxicity of
contaminated sediments in dilution series with control sediments. Chemosphere 27:1789-1812.
Nelson, D.W. and L.E. Sommers. 1996. Total carbon, organic carbon, and organic matter. In:
Methods of Soil Analysis: Part 3 Chemical Methods, D.L. Sparks et al. (eds.). Soil Science Society
of America, Inc. Madison, WI.
Nkedi-Kizza, P., P.S.C. Rao, A.G. Hornsby. 1985. Influence of organic cosolvents on sorption of
hydrophobic organic chemicals by soils. Environmental Science and Technology 19:975-979.
Northcott, G.L. and K.C. Jones. 2000. Spiking hydrophobic organic compounds into soil and
sediment: a review and critique of adopted procedures. Environmental Toxicology and Chemistry
19:2418-2430.
8-14 US Environmental Protection Agency
-------
Technical Manual
O'Donnel, J.R., B.M. Kaplan, and H.E. Allen. 1985. Bioavailability of trace metals in natural waters.
Aquatic Toxicol. And Hazard Assessment: Seventh symposium. ASTM STP 854, ASTM,
Philadelphia, PA. pp. 485-501.
O'Neill, E.J., C.A. Monti, P.H. Prichard, A.W. Bourquin, and D.G. Ahearn. 1985. Effects of
Lugworms and seagrass on kepone (chlordecone) distribution in sediment-water laboratory systems.
Environmental Toxicology and Chemistry 4:453-458.
Page, A. L., Miller, R. H., and Keeney, D. R. (eds.). 1982. Methods of Soil Analysis Parts 1 and 2.
Amer. Soc. Agron., Madison, WI.
Page, D.S., P.D. Boehm, G.S. Douglas, and A.E Bence. 1995a. Identification of Hydrocarbon
Sources in the Benthic Sediments of Prince William Sound and the Gulf of Alaska Following the
Exxon Valdez Oil Spill. Exxon Valdez Oil Spill: Fate and Effects in Alaskan Waters, ASTM STP
1219, Peter G. Wells, James N. Butler, and Jane S. Hughes, Eds., American Society for Testing and
Materials, Philadelphia, PA.
Page, D.S., E.S Gilfillan,. P.D Boehm, and E.J Harner. 1995b. Shoreline Ecology Program for Prince
William Sound, Alaska, Following the Exxon Valdez Oil Spill: Part I-Study Design and Methods.
Exxon Valdez Oil Spill: Fate and Effects in Alaskan Waters, ASTM STP 1219, James N. Butler, and
Jane S. Hughes, Eds., American Society for Testing and Materials, Philadelphia, PA.
Pascoe, G.A. and J. A. DalSoglio. 1994. Planning and implementation of a comprehensive ecological
risk assessment at the Milltown Reservoir-Clark Fork River superfund site, Montana. Environmental
Toxicology and Chemistry 13:1943-1956.
Patrick, W. H. Jr. 1958. Modification of Method Particle Size Analyses. Proceedings of the Soil
Science Society of America 4:366-367.
Phillips, B.M., B.S. Anderson, and J.W. Hunt. 1997. Measurement and distribution of interstitial and
overlying water ammonia and hydrogen sulfide in sediment toxicity tests. Marine Environmental
Research 44(2): 117-126.
Plumb, R. H., 1981. Procedures for Handling and Chemical Analysis of Sediment and Water
Samples. Environmental Protection Agency/Corps of Engineers Technical Committee on Criteria for
Dredged and Fill Material, Contract EPA-4805572010.
Prichard, P.H., C.A. Monti, E.J. O'Neill, J.P. Connolly, D.G. Ahearn. 1986. Movement of kepone
(chlorodecone) across an undisturbed sediment-water interface in laboratory systems. Environmental
Toxicology and Chemistry 5:667-673.
Puget Sound Estuary Program (PSEP). 1997a. Recommended guidelines for sampling marine
sediment, water column, and tissue in Puget Sound. U.S. Environmental Protection Agency, Region
10, Seattle, WA and Puget Sound Water Quality Authority, Olympia, WA.
PSEP. 1987b. Recommended Guidelines for Measuring Metals in Puget Sound Water, Sediment, and
Tissue Samples. Prepared for U.S. Environmental Protection Agency Region 10, Office of Puget
Sound, Seattle, WA and Puget Sound Water Quality Authority, Olympia, WA. Water Quality
Authority, Olympia, WA.
Chapter 8: References 8-15
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
PSEP. 1987c. Recommended Guidelines for Measuring Organic Compounds in Puget Sound
Sediment, and Tissue Samples. Prepared for U.S. Environmental Protection Agency Region 10,
Office of Puget Sound, Seattle, WA and Puget Sound Water Quality Authority, Olympia, WA. Water
Quality Authority, Olympia, WA.
PSEP. 1996. Recommended Guidelines for measuring selected environmental variables in Puget
Sound. Prepared for U.S. Environmental Protection Agency Region 10, Office of Puget Sound,
Seattle, WA and Puget Sound Water Quality Authority, Olympia, WA. Water Quality Authority,
Olympia, WA.
PSEP. 1995. Recommended guidelines for conducting laboratory bioassays on Puget Sound
sediments. Prepared for U.S. Environmental Protection Agency Region 10, Office of Puget Sound,
Seattle, WA and Puget Sound Water Quality Authority, Olympia, WA. Water Quality Authority,
Olympia, WA.
Reeburgh, W.S. 1967. An improved interstitial water sampler. Limnology and Oceanography 12:163-
165.
Resendes, J., W.Y. Shiu, and D. Mackay. 1992. Sensing the fugacity of hydrophobic organic
chemicals in aqueous systems. Environmental Science and Technology 26:2381-2387.
Ribeiro, R., R. Margalho, F. Goncalves, and A. Scares. 1994. Abstr. HC07, p. 224. Annu. Meet. Soc.
Environmental Toxicology and Chemistry, Denver, CO.
Robinson, A.M., J.O. Lamberson, F.A. Cole, and R.C. Swartz. 1988. Effects of culture conditions on
the sensitivity of a phoxocephalid amphipod, Rhepoxynius abronius, to cadmium in sediment.
Environmental Toxicology and Chemistry 7:953-959.
Rosenfeld, J.K. 1979. Ammonia absorption in nearshore anoxic sediments. Limnology and
Oceanography 24:356-364.
Ross, P.E., and M.S. Henebry. 1989. Use of four microbial tests to assess the ecotoxicological hazard
of contaminated sediments. Toxicity Assessment 4:1-21.
Rukavina, N. A. and Duncan, G. A. 1970. F.A.S.T.- Fast Analysis of Sediment Texture. Proceedings
of the Conference on Great Lakes Research, pp. 274-281.
Saager, P.M., J-P Sweerts, and H.J Ellermeijer. 1990. A simple pore-water sampler for coarse, sandy
sediments of low porosity. Limnology and Oceanography 35:747-751.
Sanford, R. B., and D.J.P. Swift. 1971. Comparisons of Sieving and Settling Techniques for Size
Analysis, Using a Benthos Rapid Sediment Analyzer. Sedimentology 17:257-264.
Santschi, P.H., J. Lenhart, and B.D. Honeyman. 1997. Heterogeneous processes affecting trace
contaminant distribution in estuaries: The role of natural organic matter. Marine Chemistry
58:99-125.
Sarda, N. and G.A. Burton Jr. 1995. Ammonia variation in sediments: Spatial, temporal and method-
related effects. Environmental Toxicology and Chemistry 14:1499-1506.
8-16 US Environmental Protection Agency
-------
Technical Manual
Sasson-Brickson, G. and G.A. Burton, Jr. 1991. In situ and laboratory sediment toxicity testing with
Ceriodaphnia dubia. Environmental Toxicology and Chemistry 10:201-207.
Sawyer, L.N. and G.A. Burton, Jr. 1994. Validation of various formulated sediment recipes for use in
toxicity assessments. Abstr. HC06, p. 224. Annual Meeting of the Society for Environmental
Toxicology and Chemistry. Denver, CO.
Sayles, F.L., T.R.S. Wilson, D.N. Hume, and P.C. Mangelsdorf Jr. 1973. In situ sampler for marine
sedimentary pore waters: Evidence for potassium depletion and calcium enrichment. Science
180:154-156.
Schlekat, C.E., K.J. Scott, R.C. Swartz, B Albrecht, L. Anrim, K. Doe, S. Douglas, J.A. Ferretti, D.J.
Hansen, D.W. Moore, C. Mueller, and A. Tang. 1995. Interlaboratory comparison of a 10-day
sediment toxicity test method using Ampelisca Abdita, Eohaustorius Estuarius, and Leptocheirus
Plumulosus. ET&C, 14:2163-2174.
Schubauer-Berigan, M.K. and G.T. Ankley. 1991. The contribution of ammonia, metals and nonpolar
organic compounds to the toxicity of sediment interstitial water from an Illinois River tributary.
Environmental Toxicology and Chemistry 10:925-939.
Schults, D.W. S.P. Ferraro, L.M. Smith, F.A. Roberts, and C.K. Poindexter. 1992. A comparison of
methods for collecting interstitial water for trace organic compounds and metals analyses. Water
Research 26:989.995.
Schuytema, G.S., P.O. Nelson, K.W. Malueg, A.V. Nebeker, D.F. Krawczyk, A.K. Ratcliff, and J.H.
Gakstatter. 1984. Toxicity of cadmium in water and sediment to Daphnia magna. Environmental
Toxicology and Chemistry 3:293-308.
SET AC. 2001. Porewater Toxicity Testing: Biological, Chemical, and Ecological Considerations
with a Review of Methods and Applications, and Recommendations for Future areas of Research.
SET AC Technical Workshop. Society for Environmental Toxicology and Chemistry, Pensacola, FL.
Sijm, R.T.H., M. Haller, and S.M. Schrap. 1997. Influence of storage on sediment characteristics and
drying sediment on sorption coefficients of organic contaminants. Bulletin of Environmental
Contamination and Toxicology 58:961-968.
Simon, N.S., M.M. Kennedy, and C.S. Massoni. 1985. Evaluation and use of a diffusion controlled
sampler for determining chemical and dissolved oxygen gradients at the sediment-water interface.
Hydrobiologia 126:135-141.
Singer, J. K., Anderson, J. B., Ledbetter, M. T., McCave, I. N., Jones, K. P. N., and Wright, R. 1988.
An Assessment of Analytical Techniques for the Size Analysis of Fine-Grained Sediments. Journal
of Sediment Petrology 58:534-543.
Skalski, C. and G. A. Burton. 1991. Laboratory and In Situ Sediment Toxicity Evaluations Using
Early Life Stages of Pimephales promelas. M.S. thesis. Wright State University, Dayton, OH.
Solomon, K.R, Ankley G.T, Baudo R, Burton G.A, Ingersoll C.G, Lick W, Luoma S, MacDonald
DD, Reynoldson TB, Swartz RC, Warren-Hicks WJ. 1997. Work group summary report on
methodological uncertainty in sediment ecological risk assessment. In: Ingersoll CG, Dillon T,
Chapter 8: References 8-17
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Biddinger RG (editors). Ecological risk assessment of contaminated sediment. Pensacola FL: SET AC
Press, p 271-296.
Stemmer, B.L., G.A. Burton, Jr., and S. Leibfritz-Frederick. 1990a. Effects of sediment test variables
on selenium toxicity to Daphnia magna. Environmental Toxicology and Chemistry 9:381-389.
Stemmer, B.L., G.A. Burton, Jr., and S. Leibfritz-Frederick. 1990b. Effect of sediment spatial
variance and collection method on Cladoceran toxicity and indigenous microbial activity
determinations. Environmental Toxicology and Chemistry 9:1035-1044.
Stephenson, R.R., and D.F. Kane. 1984. Persistence and effects of chemicals in small enclosures in
ponds. Arch. Environmental Toxicology and Chemistry 13:313-326.
Sternberg, R. W., and J.S. Creager. 1961. Comparative Efficiencies of Size Analysis by Hydrometer
and Pipette Methods. Journal of Sediment Petrology 31:96-100.
Stewart, A.R., and D.F. Malley. 1999. Effect of metal mixture (Cu, Zn, Pb, and Ni) on cadmium
partitioning in littoral sediments and its accumulation by the freshwater macrophyte Eric/cation
septanguare . Environmental Toxicology and Chemistry 18(3):436-447.
Swartz, R.S., D.W. Schults, T.H. DeWitt, G.R. Ditsworth, and J.O. Lamberson. 1990. Toxicity of
fluoranthene in sediment to marine amphipods: A test of the equilibrium partitioning approach to
sediment quality criteria. Environmental Toxicology and Chemistry 9:1071-1080.
Swift, D.J.P., J.R. Schubel, and R.W. Sheldon. 1972. Size analysis of fine-grained suspended
sediments: A review. Journal of Sedimentary Petrology 42:122-134.
Tatem, H.E. 1986. Bioaccumulation of poly chlorinated biphenyls and metals from contaminated
sediment by freshwater prawns, Macrobracium rosenbergii, and clams, Corbiculafluminea. Archives
of Environmental Contamination & Toxicology 15:171-183.
Thurston, R.V., R.C. Russo and G.A. Vinogradov. 1981. Ammonia toxicity to fishes. Effect of pH on
the toxicity of the unionized ammonia species. Environmental Science and Technology 15:837-840.
Tinsley, I.J. 1979. Chemical Concepts in Pollution Behavior. Wiley-Interscience, New York, p. 92.
Troup, B.N., O.P. Bricker, and J.T. Bray. 1974. Oxidation effect on the analysis of iron in the
interstitial water of recent anoxic sediments. Nature 249:237.
Truax, D.D., A. Shindala and H. Sartain. 1995. Comparison of two sediment oxygen demand
measurement techniques. Journal of Environmental Engineering, September, pp. 619-624.
Tye, R., R. Jepsen and W. Lick. 1996. Effects of colloids, flocculation, particle size, and organic
matter on the adsorption of hexachlorobenzene to sediments. Environmental Toxicology and
Chemistry 15:643-651.
Uchrin, C.G. and W.K. Ahlert. 1985. In situ sediment oxygen demand determinations in the Passaic
River (NJ) during the late summer/early fall 1983. Water Resources 19:1141-1144.
8-18 US Environmental Protection Agency
-------
Technical Manual
U.S. Army Corps of Engineers. 1976. Ecological evaluation of proposed discharge of dredged or fill
material into navigable waters. Miscellaneous Paper D-76-17, Waterways Experiment Station,
Vicksburg, MS.
U.S. Environmental Protection Agency. 1979. Chemistry Laboratory Manual for Bottom Sediments
and Elutriate Testing. EPA-905-4-79-014 (NTIS PB 294596) EPA Region V, Chicago, IL.
U.S. Environmental Protection Agency. 1983. Methods for the chemical analysis of water and
wastes. EPA 600/4-79-020. U.S. Environmental Protection Agency, Environmental Monitoring and
Support Laboratory, Cincinnati, OH. 460pp.
U.S. Environmental Protection Agency. 1986a. Occupational health and safety manual. Office of
Administration, Washington, DC.
U.S. Environmental Protection Agency. 1986b. Test methods for evaluating solid waste (SW-846):
physical/chemical methods. U.S. Environmental Protection Agency, Office of Solid Waste,
Washington, DC.
U.S. Environmental Protection Agency. 1987. Quality Assurance/Quality Control (QA/QC) for
301 (h) Monitoring Programs: Guidance on Field and Laboratory Methods. U.S. EPA 430/9-86-004.
U.S. Environmental Protection Agency. 1991. Methods for measuring the acute toxicity of effluents
and receiving waters to freshwater and marine organisms. Fourth edition. EPA-600/4-90/027F,
Cincinnati, OH.
U.S. Environmental Protection Agency. 1993. U.S. EPA Contract Laboratory Program - statement of
work for organic analysis, multi-media, multi-concentration. Document ILMO1.0-ILMO-1.9, 1993.
U.S. Environmental Protection Agency, Washington, DC.
U.S. Environmental Protection Agency. 1994. Methods for measuring the toxicity of sediment-
associated contaminants with estuarine and marine amphipods. EPA-600/R-94/025, Narragansett, RI.
U.S. Environmental Protection Agency. 1995. QA/QC Guidance for Sampling and analysis of
sediments, water, and tissues for dredged material evaluations (chemical evaluations). EPA 832-B-
95-002. Office of Water, Washington, D.C.
U.S. Environmental Protection Agency. 1996. Eco Update Ecotox Thresholds. EPA 540-F-95-0380
U.S. Environmental Protection Agency. 1997a. The incidence and severity of sediment
contamination in surface waters of the United States. Volume 1: National Sediment Quality Survey.
EPA 823-R-97-006. Office of Science and Technology, Washington, DC.
U.S. Environmental Protection Agency. 1997b. EPA Diving safety manual - August, 1997. Prepared
by the Office of Administration and Resource Management, Safety, Health, and Environmental
Management Division.
U.S. Environmental Protection Agency. 1997 f. Ecological Risk Assessment Guidance for Superfund:
Process for Designing and Conducting Ecological Risk Assessments. EPA 540-R-97-006.
U.S. Environmental Protection Agency. 1998. Contaminated sediment management strategy. EPA
823-R-98-001. Office of Water, Washington, DC.
Chapter 8: References 8-19
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
U.S. Environmental Protection Agency. 1999. 1999 Update of Ambient Water Quality Criteria for
Ammonia. EPA-823-F-99-013, Office of Water, Washington, DC.
U.S. Environmental Protection Agency. 2000a. Guidance for the Data Quality Objectives Process.
EPA QA/G-4. EPA/600/R-96/055. Office of Environmental Information, Washington, D.C.
U.S. Environmental Protection Agency. 2000b. Guidance for Choosing a Sampling Design for
Environmental Data Collection.
U.S. Environmental Protection Agency. 2000c. Estuarine and Near Coastal Marine Waters:
Bioassessment and Biocriteria Technical Guidance. EPA-822-B-00-004.
U.S. Environmental Protection Agency. 2000d. Methods for measuring the toxicity and
bioaccumulation of sediment-associated contaminants with freshwater invertebrates. Second Edition.
EPA/600/R-99/064, Duluth, MN.
U.S. Environmental Protection Agency. 2000e. Guidance in the Use and Development of a Quality
Assurance Project Plan. EPA QA/G-5S, Office of Environmental Information, Washington, D.C. In
Press.
U.S. Environmental Protection Agency/Army Corps of Engineers. 1991. Evaluation of dredged
material proposed for ocean disposal: Testing manual. EPA-503/8-91/001, Office of Water,
Waterways Experiment Station, Vicksburg, MS.
U.S. Environmental Protection Agency/ Army Corps of Engineers. 1998. Evaluation of dredged
material proposed for discharge in waters of the U.S. - testing manual. EPA-823-B-98-004,
Washington, DC.
U.S. Geological Survey. 1969. Techniques of Water-Resources Investigations of the U.S.G.S. Chp.
Cl, Harold P. Guy, p. 58, Laboratory Theory and Methods for Sediment Analysis; Book 5,
Laboratory Analysis, U.S.G.S. Arlington, VA.
Vanderpleog, H. A. 1981. Effect of the Algal Length/Aperture Length Ratio on Coulter Analyses of
Lake Seston. Canadian Journal of Fisheries & Aquatic Science 38:912-916.
Vecchi, M., T.B. Reynoldson, A. Pasteris and G. Bonomi. 1999. Toxicity of copper-spiked sediments
to Tubifex tubifex (Oligochaeta, Tubificidae): Comparison of the 28-day reproductive bioassay with
an early-life-stage biassay. Environmental Toxicology and Chemistry 18(6): 1144-1148.
Warwick, R.M., and K.R. Clarke. 1991. A comparison of some methods for analyzing changes in
benthic community structure. Journal of the Marine Biological Association of the United Kingdom
71:225-244.
Walters, D.B. and C.W. Jameson. 1984. Health and safety for toxicity testing. Butterworth
Publications, Woburn, MA.
Washington Department of Ecology (WDE). 1995. Sediment Sampling Analysis Plan Appendix:
Guidance on the Development of Sediment Sampling and Analysis Plans Meeting the Requirements
of the Sediment Management Standards. Ecology Publication No. 95-XXX, Washington Department
of Ecology, Seattle, WA.
8-20 US Environmental Protection Agency
-------
Technical Manual
Waters, D.B. (Ed.). 1980. Safe handling of chemical carcinogens, mutagens, teratogens and highly
toxic substances. Ann Arbor Science, Ann Arbor, MI.
Watson, P.O., and T.E. Prickers. 1990. A multilevel, in situ pore-water sampler for use in intertidal
sediments and laboratory microcosms. Limnology and Oceanography 35:1381-1389.
Watson, P.O., P. Prickers, and C. Goodchild. 1985. Spatial and seasonal variations in the chemistry
of sediment interstitial waters i the Tamar estuary. Estuaries and Coastal Shelf Science. 21:105-119.
Watzin, M., A. Mclntosh, E. Brown, R. Lacey, D. Lester, K. Newbrough, and A. Williams. 1997.
Assessing sediment quality in heterogeneous environments: a case study of a small urban harbor in
Lake Champlain, Vermont USA. Environmental Toxicology and Chemistry 16:2125-2135.
Webster, G.R.B., M.R. Servos, G.G. Choudhry, L.P. Sarna, and G.C.G. Muir. 1990. Methods for
dissolving hydrophobics in water for studies of their interactions with dissolved organic matter.
Advances in Chemistry Series, Presented at the 193rd National Meeting of the American Chemical
Society, Division of Environmental Chemistry. Extended Abstracts. 28:191-192.
Weliky, K., E. Suess and C.A. Ungerer. 1983. Problems with accurate carbon measurements in
marine sediments and particulate in seawater: a new approach. Limnology and Oceanography
28:1252-1259.
West, R.J. and S.J. Gonsior. 1996. Biodegradation of triethanolamine. Environmental Toxicology and
Chemistry 15:472-480.
Whitfield, M. 1969. Eh as an operational parameter in estuarine studies. Limnology and
Oceanography 14:547.
Whitfield, M. 1978. The Hydrolysis of Ammonium Ions in Sea Water—Experimental Confirmation of
Predicted Constants at One Atmosphere Pressure. Journal of the Marine Biological Association of
the United Kingdom 58:781-787.
Winger, P.V. and P.J. Lassier. 1991. A vacuum-operated porewater extractor for estuarine and
freshwater sediments. Archives of Environmental Contamination & Toxicology 21:321-324.
Word, J.Q., J.A. Ward, L.M. Franklin, V.I. Cullinan, and S.L. Kiesser. 1987. Evaluation of the
equilibrium partitioning theory for estimating the toxicity of the nonpolar organic compound DDT to
the sediment dwelling amphipod Rhepoxynius abronius. Battelle/Marine Research Laboratory
Report, Task 1, WA56, Sequim, WA. 60pp.
Wright, L.D., D.B. Prior, C.H. Hobbs, R.J. Byrne, J.D. Boon, L.C. Schaffner, and M.O. Green. 1987.
Spatial variability of bottom types in the lower Chesapeake Bay and adjoining estuaries and inner
shelf. Estuarine, Coastal, and Shelf Sciences 24:765-784.
Yamamuro, M. and H. Kayanne. 1995. Rapid direct determination of organic carbon and nitrogen in
carbonate-bearing sediments with a Yanaco MT-5 CHN analyzer. Limnolology and Oceanography
40:1001-1005.
Chapter 8: References 8-21
-------
Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Yee, S., M. Van Rikxoort, and D. McLeay. 1992. The effect of holding time on Eohaustorius
washingtonianus during ten-day sediment bioassays and reference toxicant tests. Report prepared for
Environment Canada and the Inter-Governmental Aquatic Toxicity Group, North Vancouver, BC
53p.
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APPENDIX A
EXAMPLES OF SEDIMENT QUALITY
SAMPLING DESIGNS USING THE DATA
QUALITY OBJECTIVES (DQO) PROCESS
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Technical Manual
The Data Quality Objectives (DQO) process is a logical progression of steps that define the question
to be answered and identifies qualitatively and quantitatively the procedures and decisions necessary
to address the question posed. USEPA (2000a) discusses a 7-step DQO process that leads one
through each of the decision points to help ensure a successful study or program outcome.
Sediment quality monitoring studies, whether for regulatory or non-regulatory purposes, would
benefit from following USEPA's DQO process in order to:
• reduce the likelihood of collecting improper or inappropriate samples
• increase the likelihood of collecting representative samples for the question asked
• decrease the chances of introduced measurement artifacts or interference due to sampling or
sample processing techniques
• increase the likelihood that data, and decisions based on those data, will be scientifically
defensible and accepted by those involved.
The following tables are hypothetical examples demonstrating how the DQO process could be used
in addressing a few common purposes for collecting sediment quality data. The purpose of the study,
or question needing to be answered, drives the input for all subsequent steps in the DQO process.
Thus, sampling design, how samples are collected and manipulated, and the types of analyses chosen,
should all stem from the overall purpose of the study. Many national and regional programs (e.g.,
NOAA's Status and Trends, USEPA's Dredge Materials Management Program, or Puget Sound
Estuary Program) already have a particular purpose identified, thus giving rise to the particular
sampling protocols they each use.
Appendix A: Examples of Sediment Quality Sampling Designs A-3
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Example 1. Objective: Determine whether certain point and nonpoint sources are associated with
sediment contamination in a lake, estuary, or river segment
DQO Element
Issues/Concerns/Information
Certain point and nonpoint sources of concern
Enough resources for a small-moderate survey depending on number
of analyses per station
1. State problem/available
resources
2. Identify questions to be
addressed
How does sediment quality near these sources compare with other
locations and with Ecotox Thresholds (USEPA, 1996)? How toxic are
they?
3. Identify information/
measurements needed
• Use available data, source information, BPJ to identify contaminants
of concern
Measurements could include the following:
• lOd whole sediment toxicity tests
• Acute or chronic toxicity tests using interstitial water
• Benthic macroinvertebrate analyses
• Contaminant analyses (e.g., PAHs, PCBs, metals, pesticides)
• Particle size, AVS (if metals a concern), TOC, % moisture, pH,
ammonia measured for each sample
• Water, pH, oxygen, conductivity/salinity overlying sediment at each
site
4. Define spatial/temporal
boundaries
Sample during one index period
Surficial sediment (top 0 to 2 or up to 15 cm) of most interest
Concentrate sampling near suspected contaminant sources with some
reference stations (locations removed from potential sources) as well
5. Define thresholds or
decision rule for
parameters of interest
Ecotox Thresholds (USEPA, 1996), and/or other sediment threshold
values for contaminants
Toxicity effect level: e.g., significantly lower survival than reference
stations or survival < 50%
6. Limits on decision errors
Precision: < 40% C.V. among field replicates for contaminants and
toxicity
Test for differences between suspect and reference sites at p = 0.05
and power = 80%
Field blanks for contaminants < detection limit
Lab duplicates for contaminants yield < 25% C.V. Toxicity test
replicates < 35% C.V.
Tox test controls meet EPA minimum performance requirements.
7. Optimize the design
Choose targeted sampling design including reference stations
Sample when conditions most favorable for gear efficiency and
personnel safety
Use grab sampler - Ponar, Van Veen, or Petersen (see Table E-l for
advantages and disadvantages)
Use GPS for site positioning (± 10m)
Composite several (determined by number of contaminant analyses
desired) grabs at each site for a single sample
Take 3 replicate samples at 10% of the sites, selected at random
See flowchart for Selecting a Grab Sampler Based on Site-Specific
Factors (Figure 3-2).
A-4
US Environmental Protection Agency
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Example 2. Objective: Determine the status of sediment quality in a site (e.g., lake, estuary, or river
segment)
DQO Element
Issues/Concerns/Information
1. State problem/available
resources
Sediment quality unknown or status was determined in the past and
there is a need to determine how the quality may have changed.
Enough resources for a moderate survey depending on number of
analyses per station.
2. Identify questions to be
addressed
How does sediment quality compare with Ecotox Thresholds (USEPA,
1996)? How toxic are sediments now as compared to historically?
Identify information/
measurements needed
• Use available data, source information, BPJ to identify contaminants
of concern
Measurements could include the following:
• lOd whole sediment toxicity tests
• Acute or chronic toxicity tests using interstitial water
• Benthic macroinvertebrate analyses
• Contaminant analyses (e.g., PAHs, PCBs, metals, pesticides)
• Particle size, AVS (if metals a concern), TOC, % moisture, pH,
ammonia measured for each sample
• Water, pH, oxygen, conductivity/salinity overlying sediment at each
site
4. Define spatial/temporal
boundaries
Sample during one season (index period)
Sample surficial as well as deeper sediments to obtain historical
record.
Sample stations representative of the entire site or, if site contains
different subareas of interest (e.g., areas having very different salinity
zones or different geology/sediment particle size), representative
samples of each subarea.
5. Define thresholds or
decision rule for
parameters of interest
Ecotox Thresholds (USEPA, 1996), and/or other sediment threshold
values for contaminants
Toxicity effect level: e.g., significantly lower survival than reference
stations or survival < 50%
6. Limits on decision errors
Precision: < 40% C.V. among field replicates for contaminants and
toxicity
Test for differences between suspect and reference sites at p = 0.05
and power = 80%
Field blanks for contaminants < detection limit
Lab duplicates for contaminants yield < 25% C.V. Toxicity test
replicates < 35% C.V.
Tox test controls meet EPA minimum performance requirements.
7. Optimize the design
Choose probabilistic sampling design; use stratified random or multi-
stage random design if interested in comparing quality with respect to
certain habitat features or subareas of site, respectively.
Use a corer sampler to obtain vertical (historical) profiiles of sediment
at each station. Collect and analyze samples of strata of interest. Use
of a larger corer (e.g., box corer) will mean fewer cores needed per
station (see Table E-2 for advantages and disadvantages of different
corers.)
Use GPS for station positioning (± 10 m).
Take 3 replicates for each type of analysis at 10% of the stations.
See Flowchart for Selecting Core Samplers Based on Site-Specific
Factors (Figure 3-3).
Appendix A: Examples of Sediment Quality Sampling Designs
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Example 3. Objective: Determine the need for or locations of site remediation (e.g., superfund)
DQO Element
Issues/Concerns/Information
1. State problem/available
resources
Site known or suspected to contain contaminated sediments that pose
an ecological and/or human health risk
Resources are available for a moderate-intensive survey
2. Identify questions to be
addressed
Does the site need to be remediated? Where at the site is sediment
remediation warranted?
3. Identify information/
measurements needed
Use previously collected data, if available, to identify contaminants of
concern. If no information is available, a pilot survey, using a random
sampling design, may be useful to identify potential contaminants of
concern.
Measurements could include:
- Contaminants of concern in whole sediment and/or interstitial
water
- 10 d whole sediment toxicity tests
Acute or chronic interstitial water toxicity tests
Benthic macroinvertebrate analyses
Particle size, AVS (if metals a concern), TOC, % moisture, pH,
ammonia to help interpret chemical or toxicological data.
4. Define spatial/temporal
boundaries
Sample over one or more index periods depending on assumed or
measured rates of sediment or contaminant movement.
Surficial as well as deeper sediments may need to be sampled
depending on depth of contamination.
Sampling all areas of the site may be necessary to locate areas in need
of remediation unless more information is available.
5. Define thresholds or
decision rule for
parameters of interest
Contaminant levels exceed Ecotox Thresholds (USEPA, 1996).
Toxicity effect level: e.g., significantly lower survival than reference
sediment and < 50%.
6. Limits on decision errors
Precision: < 40% C.V. among field replicates for contaminants and
toxicity
Test for differences between suspect and reference sites at p = 0.05
and power = 80%
Field blanks for contaminants < detection limit
Lab duplicates for contaminants yield < 25% C.V. Toxicity test
replicates < 35% C.V.
Tox test controls meet EPA minimum performance requirements.
7. Optimize the design
Choose systematic or grid sampling design if no previous information
available on areas of contamination.
Choose targeted design if information is already available on areas of
contamination within the site.
Choose multi-stage design if more than one area of contamination
within the site is known but locations of contamination within each
area are not precisely known.
Use grab sampler if remediation will involve only surficial sediments,
or sediment depth is known to be shallow (see Table E-l and Figure
3-2).
Use corer if remediation is likely to involve deeper sediments. For
areas in which remediation may entail very deep sediments (> 2 m),
consider using a vibracorer or piston corer (see Table E-2 and the
Flowchart for Selecting Core Samplers Based on Site-Specific Factors
(Figure 3-3).
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APPENDIX B
EXAMPLES OF MEASUREMENT QUALITY
OBJECTIVES USED IN SEDIMENT QUALITY
MONITORING STUDIES
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In the Data Quality Objectives (DQO) framework (discussed in Chapter 2 and examples presented in
Appendix A of this Manual), a key element of this process is defining the thresholds or decision rules
(Step 5, Figure 2-2) and the limits on errors pertaining to those decisions (Step 6, Figure 2-2). Both
of these steps are critical to the DQO process, and the success of a study, because they explicitly
define whether a particular result qualifies as an effect of interest, and when and where something
might need to be done to mitigate or address a given observed effect. Also, these steps are critical
factors in designing a tiered or phased sampling program. Thresholds, for example, can be initially
set to identify problem areas with high accuracy (low decision error). This would be followed by a
second sampling, with a lower threshold, to identify emerging or more subtle problems in a cost-
effective manner.
The information used to help derive meaningful threshold or decision rules, and the tolerable errors
associated with those rules, is collectively referred to as Measurement Quality Objectives (MQOs).
MQOs are qualitative or quantitative statements that describe the type of data quality needed to
support or refute a given decision. These statements explicitly define acceptable precision, bias, and
sensitivity required of all analyses in the study and therefore, should be consistent with the expected
performance of a given analysis or test method (ITFM 1995). Thus, if a particular whole sediment
toxicity test is expected to yield 80% survival among control replicates, the MQO for control survival
should be > 80% for that test. Further, if one intends to compare sediment toxicity results between a
reference station and test stations, it is important to set the number of replicates and the decision rule
appropriately so that the study can determine with reasonable power and confidence whether a given
sediment sample is toxic to the test organisms. The number of replicates performed will depend on
the expected variability of a given test endpoint and the sensitivity desired in the study.
The following summarizes four different examples of sediment quality studies or programs, each
with a different study purpose, and the types of MQOs they used. These examples are for illustrative
purposes and are not meant to imply that these are the only acceptable ways in which MQOs can be
derived. The examples provided are:
• Shoreline ecology program following the Exxon Valdez oil spill in Alaska
• Great Lakes Assessment and Remediation of Contaminated Sediment (ARCS) Program
• An example of an EMAP study design in the St. Louis River, Minnesota/Wisconsin
• A focused assessment in Burlington Harbor, VT in Lake Champlain
• Excerpts from Washington Department of Ecology's Sampling and Analysis Plan Guidance
(WDE, 1995).
This latter guidance demonstrates how a particular program addresses sampling and analysis needs
depending on the monitoring objective. The guidance also provides an interesting comparison of
overall sampling procedures and sampling design considerations for two programs: WDE's Sediment
Management Standards Program and the Puget Sound Dredged Disposal Analysis Program, both of
which have some common monitoring objectives.
Appendix B: Examples of Measurement Quality Objectives B-3
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Example 1: Shoreline Ecology Program for Prince William Sound, Alaska, Following
the Exxon ValdezO\\ Spill
Background
A comprehensive shoreline ecology program was designed to assess recovery in Prince William
Sound following the Exxon Valdez oil spill on March 24, 1989 (Page et al., 1995a; b; Boehm et al.,
1995; Gilfillan et al., 1995; Gillfillan et al., 1999). The spill resulted in the release of about 258,000
barrels of Alaska North Slope crude oil into the marine environment. Nearly 500 miles of shorelines
in the sound were oiled to some degree.
Project Objectives
The shoreline ecology program was designed to assess the recovery of hundreds of miles of oiled
shorelines in Prince William Sound by using a limited number of sampling stations. The number of
sampling stations had to be small enough for a survey to be accomplished in the summer weather
window, but large enough to detect important spill effects. The study design consisted of two field
components: fixed sampling locations and stratified random sampling locations. The 12 fixed
locations provided information on the changes in amount and composition of petroleum residues over
the period 1989-1991 to assess the rate of shoreline recovery and oil loss. Stations chosen
represented worst-case oiling conditions and reference sites. Data gathered from these sites were
used to assess oil loss, oil weathering, and bioavailability of oil residues to mussel communities.
The stratified random sampling (SRS) of 64 sample locations permitted results to be generalized to
the affected area of the sound. The SRS survey of the spill area shoreline was divided into four
habitats which characterized over 99% of the shoreline of interest, and four oiling levels which
produced information on all shoreline spill levels. The matrix of four habitats by four oiling levels,
with each cell containing four replicates, constituted a reasonable compromise between project cost,
the need to complete sampling within the short Alaskan summer, and the need for statistical power.
The principal objective was to compare means within strata (habitat/oiling level) and not to obtain
overall estimates (see Table B-l).
Specific natural variables, including wave exposure, percentage sand, percentage silt/clay, and total
organic carbon (TOC) were also quantified, and served as covariates in statistical analyses of oil
effects.
Precautions were taken to minimize the possibilities for petroleum hydrocarbon contamination of
field samples by:
• positioning the ship's stern into the wind to prevent stack gases from blowing onto the sampling
equipment during deployment, recovery, and subsampling
• cleaning equipment just prior to arriving on station
• ensuring that the sampling equipment was never deployed or recovered through oil slicks or
sheens
• closing the top access doors to the sampler when it was not being deployed or cleaned
• field blanks were collected from each piece of equipment at regular intervals
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• potential sources of hydrocarbon contaminants were also collected to enable their identification
later
Sample documentation included station logs and chain-of-custody forms. All sediment samples were
logged in on the chain-of-custody forms along with other important information (station, date, time,
sampling equipment and method, subsampling method, and type of sample.) Any additional
information was also noted. This form accompanied each sample during shipping to the analytical
lab and each sample cooler was sealed with a custody seal which was initialed and dated by the
packer.
Several analytical laboratories were needed to process and analyze the large numbers of samples
collected. A laboratory standard oil was analyzed with each analytical batch to monitor analytical
precision and to provide data for interlaboratory comparisons. Duplicate precision for both subtidal
sediment studies and 1991 deep subtidal studies was ±30%. Other MQOs are listed in the Table B-l.
Table B-1. Measurement quality objectives for subtidal sediment studies in Prince William Sound oil
spill study (Gilfillan et al. 1995).
Parameter
Units
Practical Quantification Limit
(PQL)
Estimated Method Detection
Limit (MDL)
Procedural Blank
Field Blank
Matrix Spike Recovery
Surrogate Recovery
Duplicate Precision
EVC Control Oil Standard
Precision
Katalla Control Oil Standard
Precision
NIST SRM 1941 Precision
NIST SRM 1291 Accuracy
Subtidal Sediment Studies
|-ig/kg dry weight
10
1.0
5xMDL
5xMDL
40 - 120%a
40 - 120%b
±30%
±20%
NA
NA
NA
1991 Deep Subtidal Studies
|-ig/kg dry weight
1.0
0.1
5xMDL
5xMDL
40 - 120%a
40 - 120%b
±30%
±20%
±20%
±25%
± 15%
The average percentage recoveries for all 16 compounds must fall between 40 and 120%. Only one
compound can be below its minimum percentage recovery. This allowed a deviation for a single analyte
of not less than 10% for chrysene and benzo(a) pyrene and not less than 20% for the others.
Surrogate recoveries must fall between 40 and 120%. The upper control limit may be exceeded by one
compound.
The average percentage difference for the target compounds should not exceed 20% of the mean of all
previous values, and no single compound/isomer grouping should deviate by more than 30% of its mean
value of all previous determinations.
SRM = Standard reference material.
Appendix B: Examples of Measurement Quality Objectives
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Example 2: Measurement Quality Objectives used in the Great Lakes Assessment
and Remediation of Contaminated Sediment (ARCS) Program
Background
Although toxic discharges into the Great Lakes and elsewhere have been reduced in the last 20 years,
persistent contaminants in sediments continue to pose a potential risk to human health and the
environment (GLNPO 1994). Elevated concentrations of contaminants in bottom sediments and
associated adverse effects have been found throughout the Great Lakes and connecting channels.
The extent of sediment contamination and its associated adverse effects have been the subject of
considerable concern and study in the Great Lakes community.
To address these concerns, Annex 14 of the Great Lakes Water Quality Agreement between the
United States and Canada (as amended by the 1987 Protocol) stipulates that the cooperating parties
will identify the nature and extent of sediment contamination in the Great Lakes, develop methods to
assess impacts, and evaluate the technological capability of programs to remedy such contamination.
The 1987 amendments to the Clean Water Act, authorized GLNPO to coordinate and conduct a 5-
year study and demonstration projects relating to the appropriate treatment of toxic contaminants in
bottom sediments. To fulfill the requirements of the Act, GLNPO initiated the Assessment and
Remediation of Contaminated Sediments (ARCS) Program. ARCS is an integrated program for the
development and testing of assessment techniques and remedial action alternatives for contaminated
sediments. Information from ARCS Program activities will help address contaminated sediment
concerns in the development of Remedial Action Plans (RAPs) for all 43 Great Lakes Areas of
Concern (AOCs, as identified by the United States and Canadian governments), as well as similar
concerns in the development of Lakewide Management Plans.
Program Objectives
Sediments are associated with impairment of beneficial uses at 42 of the 43 Great Lakes AOCs.
Prior to addressing the potential need for remediation of those sediments, the following questions are
addressed:
• Are the sediments sufficiently "contaminated" to warrant consideration for remediation? In
this context, "contaminated" refers to the presence of chemicals in the sediments that have
the potential to cause adverse effects in humans or ecological receptors.
• Is there evidence indicating that existing concentrations of sediment contaminants are
adversely affecting ecological receptors? In other words, can it be shown that the presence
of contaminants in the sediments is causing adverse effects in organisms, either organisms
naturally occurring in the environment, or those exposed to sediments in controlled,
laboratory toxicity tests?
• Are ecological receptors exposed to the sediments bioaccumulating chemical contaminants to
the extent that the resultant body burdens are adversely affecting the organisms themselves
or other organisms higher in the food chain, including humans?
• If the sediments are judged to be sufficiently contaminated to be causing such effects, what is
the spatial extent (i.e., both horizontal and vertical) of the contamination, and what are the
implications of the distribution of contaminants on possible remedial alternatives?
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Early in the ARCS Program, it was recognized that the current state of sediment assessment methods
was rapidly evolving. The sediment assessment methods currently available consider a wide variety
of endpoints and effects, which differ in their suitability and sensitivity for investigating sediment
contamination. Therefore, assessment methods selected in the ARCS Program, reflect site- and
program-specific objectives of the study being conducted.
The ARCS Program developed several measurement quality objectives (MQOs) that it uses in the
design and conduct of studies at AOCs. Table B-2 summarizes these MQOs.
Table B-2. Examples of the measurement quality objectives for inorganic and organic chemistry
analyses of sediment used by the ARCS program in the Great Lakes (GLNPO, 1994).
Parameter
Total organic carbon
Oil and grease
pH
Acid-volatile sulfides
Organohalogens6
Total sulfur
Total solids
Volatile solids
Particle sizef
Solvent extractable
residue
Moisture content
PAHs
Pesticides
PCB/congener
PCB/Aroclor®
PCDDs/PCDFs
Methylmercury
Tributyltin
Metals8
MDLa
(Hg/kg)
0.03%
10,000
N/A
1,000
0.03
10,000
1,000
2,000
1,000
1,000
1,000
200
10
0.5
20
0.002
10
10
2,000
Accuracy11
± 20 percent
± 20 percent
±0.1 unit
N/A
± 20 percent
± 20 percent
N/A
N/A
windows
± 20 percent
N/A
± 20 percent
± 20 percent
± 20 percent
± 20 percent
± 20 percent
± 20 percent
± 20 percent
± 20 percent
Frequency
l/batchd
I/batch
I/batch
N/A
I/batch
I/batch
N/A
N/A
I/batch
I/batch
N/A
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
Precision'
< 20 percent
< 20 percent
±0.1 unit
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
< 20 percent
Frequency d
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
I/batch
Appendix B: Examples of Measurement Quality Objectives
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Table B-2 (continued). Examples of the measurement quality objectives for inorganic and organic
chemistry analyses of sediment used by the ARCS program (GLNPO, 1994).
Parameter
Except:
Arsenic
Cadmium
Mercury
MDLa
(US/kg)
100
100
100
Accuracy11
± 20 percent
± 20 percent
± 20 percent
Frequency
I/batch
I/batch
I/batch
Precision'
< 20 percent
< 20 percent
< 20 percent
Frequency
I/batch
I/batch
I/batch
Note: ARCS
MDL
N/A
PAH
PCB
PCDDs/PCDFs
Assessment and Remediation of Contaminated Sediments
method detection limit
not applicable
polynuclear aromatic hydrocarbon
polychlorinated biphenyl
polychlorinated dibenzo-p-dioxins/polychlorinated dibenzofurans
a Units presented in the subheading are applicable to all parameters unless otherwise noted.
b Accuracy is determined from a certified reference material, standard reference material, or
standard and is measured from the known concentration.
0 Precision is calculated as percent relative standard deviation. Precision requirements listed here
are for analytical replicates only; field duplicates are required to have a relative percent difference
< 30 percent.
d A batch is a sample group (usually 10-20 samples) that is carried through the analytical scheme
simultaneously.
e The MDL for chlorine and bromine is 30 ng, while the MDL for iodine is 10 ng.
f A soil sample with acceptance windows per size fraction was provided for use as an accuracy
standard.
E Metals include arsenic, cadmium, chromium, copper, iron, lead, manganese, mercury, nickel,
selenium, silver, and zinc. Exceptions are noted where different methodologies are used during
the metals quantification.
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Example 3: Sediment Toxicity, Contaminant Concentrations and Benthic Community
Structure as Indicators of Sediment Quality in the St. Louis River: A Test of EMAP
Concepts Applied to a Great Lakes Area of Concern
Background
The International Joint Commission (IJC) has designated 43 areas of concern (AOCs) throughout the
Great Lakes as threatened by conventional pollutants, heavy metals, toxic organic compounds,
habitat alterations, and introduction of undesirable species. Results of these disturbances have been
biological impacts (e.g., benthic macroinvertebrate and fish community degradation), human health
effects (fish consumption advisories), and beach closings. The geographic areas associated with the
AOCs contain a majority of the population residing in the Great Lakes basin, and comprise
approximately 50% of all Canadian citizens.
The St. Louis River AOC, which drains a watershed of 3,634 square miles in northern Minnesota and
Wisconsin, forms a large freshwater estuary that represents the second largest tributary to Lake
Superior. The 12,000-acre estuary is characterized by a diversity of habitat types. The AOC is
unique among the Great Lakes AOCs in that the range of habitat types and contamination status is
extreme: for example, the lower estuary contains two federal Superfund sites located across the river
from large, undisturbed tracts of forested land currently providing excellent habitat quality for a large
variety of species. The outer harbor contains actively dredged shipping channels and a number of
current or former municipal and industrial effluent discharges, as well as the world's largest
freshwater sand bar, which is home to numerous endangered or threatened plants and animals.
This project has a two-fold purpose: (1) determine if the EMAP intensified grid provides a sampling
framework that can be used, with structural modification, to assess AOCs; and (2) develop a set of
generic environmental indicators based on biological and chemical measures for long-term
assessment of AOCs using the EMAP-Great Lakes and Surface Water EMAP indicators.
In order to achieve these stated purposes, the project has four goals:
1. To test the application of the Great Lakes-EMAP design features in the Harbors and
Embayments resource class.
2. To identify percentage areas within the St. Louis River AOC having acceptable and
subnominal quality with respect to sediment contamination, toxicity and benthic community
structure, and to associate statistically certain sediment contaminants with observed ecological
effects.
3. To serve as a baseline status-and-trends monitoring survey of the St. Louis River ecosystem
health.
4. To determine the sampling intensity required to survey a complex Great Lakes AOC in order
to apply this knowledge to other AOCs within Region V.
The project will sample 120 sites within three habitat classes in the St. Louis River AOC for
sediment toxicity, chemical contaminant concentrations, and benthic community structure. The three
habitat classes are: (1) ship channels and areas in the lower estuary greater than 18 ft in depth, (2)
areas of the estuary less than 18 ft in depth, and (3) Thomson, Forbay and Fond du Lac reservoirs in
the lower St. Louis River.
Appendix B: Examples of Measurement Quality Objectives B-9
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
The distribution of sampling points in the three habitat classes is as follows: 30 sites in ship channels
and deep water areas, 30 sites in the reservoirs, and 60 sites in the shallow-water estuarine areas.
Sampling locations were selected based on the Great Lakes-EMAP grid for habitat classes 1 and 2,
and a 75-fold enhancement for habitat class 3. These numbers were determined through consultation
with EMAP statisticians at ERL-Corvallis. Each site will be sampled twice during the two-year
project period in order to estimate the short-term temporal variability for all three assessment
metrics. Split-sample, surface sediments will be used for toxicity, chemistry and benthic assessment.
Project Objectives
The questions to be answered by and/or objectives for this project are the following:
1. What percentage of the sediments in the St. Louis River AOC have unacceptable levels of
sediment contamination, toxicity, and benthic community disturbance?
2. Make statistical associations on an AOC-wide basis between contaminant levels and sediment
toxicity or sub-nominal benthic community status.
3. How many sampling sites and time points are necessary to characterize sediment quality,
using the criteria determined in Objective 1, in each of the identified habitat classes (i.e., ship
channels and deep holes, shallow shoal or stream areas, and upstream reservoirs)?
4. Establish a relevant integrity index for benthic community assessment for the St. Louis River
using the EMAP sampling design.
The requirements for precision, accuracy, completeness, representativeness and comparability of the
data in order to attain the project objectives are described in Table B-3. Objective #1 has the least
strict data quality requirements for toxicity and chemistry because the large number of samples was
designed to provide an excessively-thorough site characterization. This was done in order to increase
the likelihood of obtaining a wide variety of sediment types with which to carry out Objectives #2
and #3. In other words, the number of sites and sampling points is most likely overly abundant to
address Objective 1. However, because this project is intended as a pilot to actually establish the
requisite number of samples on an areal basis for each habitat type, an overestimate was required in
the sample design. Thus, fewer sites should be required to answer Objective #1 than to satisfy
Objectives 2 and 3; therefore, the required data attributes for Objective #1 are slightly less strict than
for the other objectives. Objective #4 does not require data for toxicity and chemistry.
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Table B-3. Summary of measurement quality objectives for the St. Louis River area of concern
sediment quality assessment by sampling goal
Objective-
Metric
Goall
#l-Toxicity
Benthos
Chemistry
Goal 2
#2-Toxicity
Benthos
Chemistry
Goal 3
#3-Toxicity
Benthos
Chemistry
Goal 4
#4-Toxicity
Benthos
Chemistry
Precision
40% RPDa
40% RPDa
30% RPD
50% RPD
30% RPD
30% RPD
30% RPD
40% RPD
30% RPD
30% RPD
30% RPD
40% RPD
N/A
N/A
30%
N/A
Accuracy
N/A
N/A
N/A
50-125%
N/A
N/A
N/A
70-125%
N/A
N/A
N/A
70-120%
N/A
N/A
N/A
N/A
Completeness
80%
80%
80%
90%
90%
90%
90%
90%
90%
90%
90%
90%
N/A
N/A
90%
N/A
Representativeness
80%
80%
80%
90%
90%
90%
90%
90%
90%
90%
90%
90%
N/A
N/A
90%
N/A
1 Relative percent difference
Appendix B: Examples of Measurement Quality Objectives
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Example 4: Ecological Effects of Sediment-Associated Contaminants in Inner
Burlington Harbor, Lake Champlain
Background
Inner Burlington Harbor of Lake Champlain has received numerous toxicants from point and
nonpoint sources in its watershed. Previous sediment sampling and analyses (Watzin et al., 1997)
demonstrated relatively high concentrations of silver, lead, and PAHs in the harbor, especially in the
southern end, compared to sites outside the breakwater. Much of this area corresponds to an old
sewage outfall and oil dolphins but could also represent migration of inputs from the old rail yard
and nonpoint sources in and around Burlington. Because the surficial sediment (top 2-3 cm) at most
sites had lower pollutant concentrations than sediments at greater depths, inputs of pollutants in
recent history (past 30 years) may be declining. However, these studies also indicated substantial
temporal and spatial heterogeneity with respect to sediment contaminant concentrations and toxicity
(Watzin etal., 1997).
Biological assessments, using benthic macroinvertebrates, were used in conjunction with other field
and laboratory analyses to help determine the effects of sediment contamination and other stressors
on the biota of Burlington Harbor.
Project Objectives
The overall objective of this project was to assess the hazard resulting from toxic contaminants in the
sediments of Inner Burlington Harbor using a sediment quality triad approach. Because certain
potentially toxic contaminants are known to occur in Burlington Harbor, the objective of this project
was divided into three major component questions.
• Have toxic sediments altered benthic communities of Burlington Harbor?
• Could such changes affect other ecological components of Lake Champlain?
• Do the toxic contaminants in Burlington Harbor sediments accumulate up the food chain and
cause risks to higher terrestrial and aquatic trophic levels and human health?
Sampling Design
Sampling locations in the present study were identified by reanalyzing the 1993-94 data from the
harbor with a spatial statistical model known as kriging (Myers, 1988) to estimate contaminant
concentrations and uncertainties throughout the harbor. Kriging is a geostatistical estimation method
which incorporates a model of the spatial variability of data directly. For each chemical, a variogram
was calculated using USEPA's software Geo-EAS (version 1.2.1) and fitted by a non-linear least-
squared procedure.
The sampling sites selected for the present study were those with the greatest uncertainty (using
existing data), and the highest likelihood of contamination. Ten sites were sampled in the harbor and
10 replicate samples from two different sites (reference sites) with relatively low contaminant
concentrations and/or toxicity were sampled to help assess sediment quality in the harbor,
particularly with respect to biological and toxicological measures. Five replicate samples were
collected from one site inside the harbor and 5 reference samples were collected from one site. The
five replicate samples collected at each reference site were tested separately for all toxicity and
biological analyses, yielding five individual measures for toxicity and macroinvertebrate community
structure at these two sites. Subsamples from each of the five samples collected at both sites were
composited into one sample from each site for physicochemical analyses. Two other sites were
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replicated once as well to obtain a measure of the variability surrounding chemical measures
obtained in this study. A total of eight sites were sampled both in this study and in previous work.
Sediment Sampling and Analyses
Sites were identified using differential global positioning and checked frequently during sampling to
ensure proper sampling location. Each site was sampled using five-seven petite Ponar grabs,
depending on the amount of sediment collected in each grab sample. Contents of the Ponar samples
from the site were composited and homogenized in the field using Teflon or high density plastic
equipment to obtain a representative sample from each site for chemical, lexicological, and
biological analyses.
Table B-4 summarizes the analyses performed in this study and the measurement quality objectives
used. Sediment chemical analyses included PAHs, simultaneously extracted metals (SEM), total
organic carbon (% TOC), acid volatile sulfides (AVS), total organic nitrogen (TON), ammonia,
particle size, and pH. Five metals (those previously showing the highest levels: silver, nickel,
copper, lead, and zinc) were measured. Zebra mussels (Dreissena polymorpha) were collected from
several sites and analyzed for tissue PAHs and percent lipid content on a composite sample of
organisms collected at each site. A portion of the sample from three inner harbor sites were sieved
(stainless steel) to isolate the fine fraction less than 63|a and also analyzed for PAHs, total organic
carbon, and organic nitrogen.
Appendix B: Examples of Measurement Quality Objectives B-13
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Table B-4. Summary of measurement quality objectives for precision, accuracy, and completeness of
biological, lexicological, sediment, organism tissue, and field chemistry analyses conducted in
Burlington Harbor (Diamond et al., 1999). RPD = relative percent difference; C.V. = coefficient of
variation.
Measurement Parameter
Benthic macroinvertebrates
• Metric values
• Metric scores
• Bioassessment scores
Field Water Quality Measurements
• Conductivity
• Temperature
• Dissolved Oxygen
pH
Laboratory Sediment Analyses
PAH
• Ammonia
• Total organic nitrogen
• Total organic carbon
AVS/SEM
• Particle size
Sediment Toxicity Analyses
• Hyalella 10-day acute
• Hyalella 28-day chronic
• Pimephales 7-day chronic
• Lumbriculus 28-day bioaccumulation
Organism Tissue Analyses
PAH
• Lead
Protein Expression Analyses
Accuracy
(% Recovery)
N/A*
N/A
N/A
N/A
N/A
N/A
N/A
±25
±30
±20
±30
±30
N/A
N/A
N/A
N/A
N/A
±30
±30
N/A
Precision
• RPD< 20%
• RPD< 5%
• RPD< 5%
± 1 % of range
±0.15°C
± 0.2 mg/L
± 0.2 units
RPD < 40%
RPD < 40%
RPD < 40%
RPD < 40%
RPD < 40%
RPD < 20%
C.V. < 30%
C.V. < 40%
C.V. < 30%
C.V. < 40%
RPD < 40%
RPD < 40%
RPD < 20%
Completeness
(%)
100
100
100
85
85
85
85
85
* Not applicable except through use of routine standards and calibration.
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Example 5: Washington Department of Ecology Sampling and Analysis Plan
Guidance
Background
The Washington Department of Ecology (WDE) provides technical guidance for developing
sampling and analysis plans for sediment investigations to be conducted under the Washington
Sediment Management Standards (SMS) program (WDE, 1995). Technical guidance on various
aspects of sediment sampling and analysis procedures that need to be considered in the design and
implementation of sediment investigations is made available through the Puget Sound Estuary
Program [PSEP] protocols.
1. Sediment Source Control Program - Methods are described for controlling the effects of point
and nonpoint source discharges through the National Pollutant Discharge Elimination System
(NPDES) permit program, state water quality permit programs, issuance of administrative orders,
or other mans determined appropriate by WDE; and
2. Sediment Cleanup Program - Administrative procedures and criteria are established to identify,
screen, rank, and prioritize, and clean up contaminated surface sediment sites.
Project Objectives: Sediment Investigations Conducted under the Sediment Source
Control Program
Adverse effects of contaminated sediments on biological resources and threats to human health
generally will only occur when there is a pathway to ecological or human receptors. In most cases,
such a pathway will only exist when surface sediments (defined by the SMS as those within the
biologically active zone) are contaminated. Contaminated sediments existing at depths below the
biologically active zone are unlikely to result in such effects unless the overlying sediments are
removed by natural (e.g., erosion, scouring) or anthropogenic (e.g., dredging, propeller scour) means, or
there are other mechanisms for the release of sediment contaminants such that exposure may occur.
Additionally, the surface sediment will be most likely to exhibit impacts from recent discharges of
contaminants. Hence, the focus of sediment sampling in the sediment source control process is
generally on the sediments within the biologically active zone.
Table B-5 summarizes sediment management standards for biological effects criteria used by
Washington Department of Ecology for Puget Sound marine sediments (WDE, 1995). These
standards are, in effect, decision rules in a Data Quality Objectives context (Step 5, Figure 2-2, this
Manual); cases where these standards are not met represent locations that are impaired and in need of
some type of management action (e.g., remediation, follow-up sampling). WDE also has standards
for many chemical contaminants (WDE, 1995) as does the Puget Sound Dredged Disposal Analysis
Program (WDE, 1995).
Appendix B: Examples of Measurement Quality Objectives B-15
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Table B-5. Sediment Management Standards Biological Effects Criteria for Puget Sound Marine
Sediments
Biological Test Sediment Quality Standards8
Sediment Impact Zone Maximum Levels,
Cleanup Screening Levels, or
Minimum Cleanup Levels*
Amphipod The test sediment has a significantly higher
(t-test, P<0.05) mean mortality than the
reference sediment, and the test sediment mean
mortality exceeds 25 percent on an absolute
basis
Larval The test sediment has a mean survivorship of
normal larvae that is significantly less (t-test,
P<0.05) than the mean normal survivorship in
the reference sediment, and the combined
abnormality and mortality in the test sediment
is more than 15 percent greater, on an absolute
basis, than the reference sediment
Benthic The test sediment has less than 50 percent of
infauna the reference area sediment's mean abundance
of any one of the following major taxa:
Crustacea, Mollusca, or Polychaeta, and the test
sediment abundance is significantly different (t-
test, P<0.05) from the reference sediment
abundance
Juvenile The mean biomass of polychaetes in the test
polychaete sediment is less than 70 percent of the mean
biomass of the polychaetes in the reference
sediment, and the test sediment biomass is
significantly different (t-test, P<0.05) from the
reference sediment biomass
Microtox® The mean light output of the highest
concentration of the test sediment is less than
80 percent of the mean light output of the
reference sediment, and the two means are
significantly different (t-test, P<0.05)
The test sediment has a significantly higher
(t-test, P<0.05) mean mortality than the
reference sediment, and the test sediment mean
mortality is more than 30 percent greater, on an
absolute basis, than the reference sediment
mean mortality
The test sediment has a mean survivorship of
normal larvae that is significantly less (t-test,
P<0.05) than the mean normal survivorship in
the reference sediment, and the combined
abnormality and mortality in the test sediment is
more than 30 percent greater, on an absolute
basis, than that in the reference sediment
The test sediment has less than 50 percent of
the reference area sediment's mean abundance
of any two of the following major taxa:
Crustacea, Mollusca, or Polychaeta, and the test
sediment abundance is significantly different (t-
test, P<0.05) from the reference sediment
abundances
The mean biomass of polychaetes in the test
sediment is less than 50 percent of the mean
biomass of the polychaetes in the reference
sediment, and the test sediment biomass is
significantly different (t-test, P<0.05) from the
reference sediment biomass
Not applicable
Source: WDE(1995).
a The sediment quality standards are exceeded if one test fails the listed criteria [WAC 173-204-320(3)].
b The sediment impact zone maximum level, cleanup screening level, or minimum cleanup level is exceeded if
one test fails the listed sediment impact zone maximum level, cleanup screening level, or minimum cleanup level
criteria [WAC 173-204-520(3)] or if two tests fail the sediment quality standards criteria [WAC 173-204-320(3)].
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WDE describes four general types of sediment monitoring (all of which are the responsibility of the
discharger) that may be conducted in support of the sediment source control process:
(a) Baseline monitoring—Used to confirm the screening evaluation for determining potential of
a discharge to cause sediment impacts conducted prior to authorization of a sediment impact
zone (SIZ) to collect information that will be used in determining whether such an authorization
is likely to be necessary, and to establish the baseline conditions with which future conditions
can be compared
(b) SIZ application monitoring—Conducted to collect information to support application of the
SIZ models
(c) SIZ maintenance monitoring—Conducted during the term of a permit that includes an
authorized SIZ, with the intent to determine whether the SIZ should be renewed, reduced, or
eliminated; whether areas of special importance have been adversely impacted by the discharge;
and the conditions for SIZ reauthorization
(d) SIZ closure monitoring—Conducted following closure of an SIZ to demonstrate successful
restoration of sediment quality.
The monitoring objectives vary with the type of monitoring being conducted, and the design of the
monitoring program varies with both discharge- and site-specific characteristics.
Project Objectives: Sediment Investigations Conducted under the Sediment Cleanup
The Sediment Cleanup Standards set forth a decision process for identifying contaminated sediment
areas and determining appropriate cleanup responses (WDE, 1995). The sediment cleanup decision
process includes procedures for screening and ranking contaminated areas of sufficient concern to
warrant active cleanup, as well as procedures for selecting an appropriate cleanup alternative on a site-
specific basis.
Because cleanup of contaminated sediments may require their removal, sediment sampling and
analyses, conducted in support of sediment cleanup studies, need to assess the total spatial extent
(including both lateral and vertical) of the sediment contamination. In this respect, these sediment
investigations differ from those previously described under the sediment source control process, where
the focus there is generally only on sediments within the biologically active zone.
In addition to initial investigations and site characterization, which are described in by WDE (1995),
there are three general types of monitoring that may be conducted in support of the sediment cleanup
process:
(a) Source control monitoring—Conducted prior to and following sediment cleanup to determine
how ongoing sources at or near a site may affect the success of active cleanup and/or natural
recovery
(b) Compliance monitoring—Long-term monitoring conducted following cleanup actions that
include containment of contaminated sediments, or to assess the progress of natural recovery
and/or to evaluate recontamination of the area
Appendix B: Examples of Measurement Quality Objectives B-17
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(c) Closure monitoring—Conducted following completion of removal actions or compliance
monitoring to demonstrate successful cleanup of sediment contamination. Closure monitoring
must be performed before a site can be considered for delisting.
The primary objectives of sediment sampling and analyses conducted as part of a preliminary
investigation of a contaminated sediment site are to: (1) Identifying sediment station clusters of
potential concern, and (2) Ranking identified cleanup sites.
Such sampling and analyses must be sufficient to enable a determination of whether there are
exceedances of the numerical chemical criteria or biological effects criteria (Table B-5) at three or more
stations within a specific area of concern. Thus, the decision rules used by WDE in these studies (Step
5 of the DQO Process, Figure 2-2, this Manual) are defined by explicit criteria and the number of the
samples demonstrating exceedence of criteria. The spatial extent of such exceedances is not required to
be defined as part of a preliminary investigation (WDE, 1995).
Given the decision rules above, there are clear implications for how sampling is designed, as there need
to be several samples collected and analyzed from a specific area of concern and some assurance of
representative coverage of the area. At smaller sites of known or suspected sediment contamination, the
addition of a relatively small number of stations or samples in a preliminary investigation is suggested
by WDE (1995) to allow assessment of the spatial extent of contamination, gradients toward or away
from other sources, or other important details. Hence, a single study could suffice, thereby precluding
the need for a second, focused investigation.
Alternatively, if there are no plans to dredge or otherwise disturb the sediments, sampling and
analyses, conducted as part of a preliminary investigation, could focus only on surface sediments.
After the need for cleanup has been identified, a more focused sediment sampling and analysis
program would then be required by WDE to define the spatial extent of contamination (including its
vertical extent) and to evaluate cleanup alternatives.
Comparison of Data Requirements: Sediment Management Standards (Sms) and the
Dredged Material Management Program (DMMP)
In addition to WDE's Sediment Management Strategy (SMS), the other major framework for sediment
management activities in the Dredged Material Management Program (DMMP). The SMS and DMMP
programs are very similar in the suites of biological and chemical evaluations that are required, and in
the evaluation criteria that are applied. While the two programs have the same goal, protection of
sediment quality, the two programs have different applications and, as a result, some differences in data
requirements.
Sediment sampling and analysis is conducted under the SMS to determine whether, and to what extent,
surface sediments are contaminated, whether point or nonpoint source discharges have contributed or
may still be contributing to such contamination, and whether contaminated sediments should be
remediated. Sediment sampling and analysis is conducted under theDMMP program to determine
whether the sediment matrix (volume) proposed for dredging, when dredged and discharged at
unconfined, open-water disposal sites within Puget Sound, could cause or contribute to unacceptable
adverse effects on the aquatic environment. Because of these different purposes, sampling gear and
compositing techniques will differ. However, both theDMMP and SMS data requirements are based
upon "exposure potential" and a "sediment unit" concept. In dredging situations (DMMP), the
exposure potential of concern is with the entire mass of sediments released at the DMMP sites and the
sediment unit of concern is the minimum dredge unit that can be effectively managed. In SMS
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situations, the exposure potential and sediment unit of concern is generally the surface, specifically the
"biologically active zone" (often the top 10 cm).
DMMP sampling is designed to characterize the bulk properties of the sediments to be dredged,
transported, and discharged. Sediment core samples (e.g., vibracorer) are typically collected to
characterize the sediment matrix to the depth of proposed dredging for disposal determinations and to
assure that the quality of newly exposed surfaces do not result in degradation. Because dredging
removes the material in bulk, the cores are typically segmented on a 4-foot basis and composited across
that segment (rather than further subdivided) to define a "dredged material management unit."
Sediment sampling under the sediment source control process of the SMS is generally designed to
characterize conditions near the sediment surface. In cases where the goal is to characterize the
exposure potential, such sampling may target the biologically active zone of the sediments. In other
cases, where the goal is to sample only the most recently deposited sediment, such sampling may target
only the uppermost 0-2 cm of sediments. Sediment sampling designed to identify contaminated
sediment sites under the sediment cleanup process of the SMS is also targeted on the near-surface,
biologically active zone of the sediments. After a contaminated site is identified, however, collection of
sediment cores will also generally be required to assess the vertical extent of contamination and to
determine the sediment quality of any new surface to be exposed after cleanup.
The process of compositing samples from a range of depth intervals below the sediment surface may
dilute higher concentrations of contaminants as pointed out in Section 2.4.3 of this Manual and in
USEPA/ACOE (1998). Compositing over depth provides an assessment of the condition of the overall
sediment matrix, but does not provide an assessment of the sediments within the biologically active
zone. Compositing of samples from a range of depth intervals is therefore appropriate for DMMP
purposes, but is ordinarily not performed for SMS investigations. In addition, many more samples may
be needed for SMS purposes to establish patterns or gradients of contamination, to identify contaminant
sources, or to delimit the area of contamination.
Development of Sediment Sampling and Analysis Plans
Although the specific details of individual sampling and analysis plans may be very different, all such
plans submitted for review by WDE contain certain basic elements. Figure B-l provides a
recommended outline for sediment sampling and analysis plans that can also serve as a checklist for
those preparing or reviewing such plans.
Each sediment sampling and analysis plan, regardless of whether it is being prepared under the
sediment source control process or the sediment cleanup process, should include as part of the
introduction a brief summary of site background information. The following background information
should be provided:
• Site history
• Regulatory framework (e.g., NPDES; Model Toxics Control Act; SMS; Comprehensive
Environmental Response, Compensation, and Liability Act)
• Summary of results of previous investigations, if any, of the site
• Location and characteristics of any current and/or historical wastewater or stormwater discharge(s) at
the site
Appendix B: Examples of Measurement Quality Objectives B-19
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
• Location and characteristics of any current and/or historical wastewater or stormwater discharge(s) in
the local area
• Information on onsite waste disposal practices or chemical spills in the local area, if any
• Site location, including a location map showing the surrounding area and a site map.
The second section of a sampling and analysis plan should describe the objectives of the sediment
investigation in the context of the appropriate regulatory framework (e.g., sediment source control
process, sediment cleanup process). WDE (1995) provides guidance on appropriate field sampling
methods; sample handling procedures; laboratory analytical methods; quality assurance and quality
control requirements; data analysis, record keeping, and reporting requirements; health and safety plan;
schedule; and project team and responsibilities.
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Sediment Sampling and Analysis Plan Outline and Checklist
(FromWDE, 1995)
1. Introduction and Background Information
Q Site history
Q Regulatory framework (e.g., NPDES, MTCA, SMS, CERCLA)
Q Summary of previous investigations, if any, of the site
Q Location and characteristics of any current and/or historical wastewater or storm water
discharge(s at the site
Q Location and characteristics of any current and/or historical wastewater or storm water
discharge(s) in the local area
Q Information on on-site waste disposal practices or chemical spills in the local area, if
any
Q Site location map showing the surrounding area
Q Site map showing site features
2. Objectives and Design of the Sediment Investigation
Q Objectives of the sediment investigation
Q Overall design of the sediment investigation, including related investigations, if any
Q Chemical analytes (including description of their relevance to the objectives and the
regulatory framework)
Q Biological tests (including description of their relevance to the objectives and the
regulatory framework)
Q Sampling Station Locations
Q Rationale for station locations
Q Site map(s) showing sampling stations and other pertinent features (e.g.,
bathymetry and current regime; outfall(s)/diffuser(s); authorized mixing
zone(s), if any; sites of waste disposal, spills, or other activities that may have
affected the sediments, such as sandblasting, boat repair, etc.; historical
dredging activities)
Q Proposed reference stations
Q Table showing the water depth at each proposed station
Q Proposed depth(s) below the sediment surface where sediments will be
collected
Figure B-1. Sediment Sampling and Analysis Plan Outline and Checklist Developed by Washington
Department of Ecology (WDE, 1995).
Appendix B: Examples of Measurement Quality Objectives B-21
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
3. Field Sampling Methods
Q Station positioning methods
Q Sampling equipment
Q Decontamination procedures
Q Sample compositing strategy and methods
Q Sample containers and labels
Q Field documentation procedures
Q Procedures for disposal of contaminated sediments
4. Sample Handling Procedures
Q Sample storage requirements (e.g., conditions, maximum holding times) for each type
of sample
Q Chain-of-custody procedures
Q Delivery of samples to analytical laboratories
5. Laboratory Analytical Methods
Q Chemical analyses and target detection limits
Q Biological analyses
Q Corrective actions
6. Quality Assurance and Quality Control Requirements
Q QA/QC for chemical analyses
Q QA/QC for biological analysis
Q Data quality assurance review procedures
7. Data Analysis, Record Keeping, and Reporting Requirements
Q Analysis of sediment chemistry data
Q Analysis of biological test data
Q Data interpretation
Q Record keeping procedures
Q Reporting procedures
Figure B-1 (continued). Sediment Sampling and Analysis Plan Outline and Checklist Developed by
Washington Department of Ecology (WDE, 1995) (cont.).
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8. Health and Safety Plan (required for cleanup investigations)
Q Description of tasks
Q Key personnel and responsibilities
Q Chemical and physical hazards
Q Safety and health risk analysis for each task
Q Air monitoring plan
Q Personal protective equipment
Q Work zones
Q Decontamination procedures
Q Disposal procedures for contaminated media and equipment
Q Safe work procedures
Q Standard operating procedures
Q Contingency plan
Q Personnel training requirements
Q Medical surveillance program
Q Record keeping procedures
9. Schedule
Q Table or figure showing key project milestones
10. Project Team and Responsibilities
Q Description of sediment sampling program personnel
Q Table identifying the project team members and their responsibilities
11. References
Q List of references
Figure B-l (continued). Sediment Sampling and Analysis Plan Outline and Checklist Developed by
Washington Department of Ecology (WDE, 1995).
Appendix B: Examples of Measurement Quality Objectives B-23
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APPENDIX C
STATISTICAL CONSIDERATIONS IN
DETERMINING THE APPROPRIATE
NUMBER OF REPLICATE SAMPLES
NEEDED AT EACH SAMPLING STATION
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For certain programs or types of studies, it is desirable (or necessary) to determine if a particular
location is significantly affected as compared to known non-impacted or reference locations (e.g.,
presence of toxicity and/or high contaminant concentrations in sediments or interstitial waters). This
type of monitoring objective is used frequently in certain regulatory programs, such as the Dredged
Materials Management Program and Superfund (CERCLA), however, many non-regulatory programs
also have a similar objective (see for example the Burlington Harbor example in Appendix B).
If one is interested in determining statistical differences in certain measures (e.g., toxicity to Hyalella
azteca) among or between stations, then analysis of replicate field samples may be necessary. This
entails collecting multiple samples from the same station (or other spatial unit of interest), processing
each sample independently, and analyzing separately each sample. For example, if the purpose of a
study is to determine whether the sediment in a specific location is toxic to the estuarine amphipod
Rhepoxynius abronius as compared to sediment from a reference location, then it is desirable to
collect multiple samples from each location and perform a Rhepoxynius whole sediment toxicity test
(including standard replication within a test) for each sample collected. Clearly, this type of
replication could entail substantial laboratory effort, as compared to compositing samples from a
single location and performing a single analysis or test (see Section 2.4.3 for a discussion of
compositing versus replication of samples). However, compositing does not provide any information
on the true variability of a given location and is rather, a form of pseudoreplication. For some
programs or studies, true field replication is necessary.
The appropriate number of replicates needed for a given study depends on the statistical power and
level of confidence (i.e., measurement quality objectives; see Appendix B for examples) one needs to
support or refute a given decision (see Data Quality Objectives Process, Section 1.1 and Appendix
A). Power is represented as 1-P and is a measure of the Type II error rate: the probability of
accepting the hypothesis that the results from two different samples or stations are similar, when in
fact they are not. Confidence is represented as 1-a and is a measure of the Type I error rate: the
probability of rejecting the hypothesis that the results from two different samples or stations are
different when in fact they are really the same. For examples, if the question is whether a given
location should be dredged for remediation purposes, the study will need to have a certain statistical
power, to determine if the sediment sample from the target location is more toxic or contaminated
than the reference location sediment, with a certain degree of confidence that one is making the
correct decision. Both power and confidence are dependent on the expected variability in the
endpoint or parameters of interest, both within a given location and within a given test or analysis.
The appropriate replication, then, is required so that one has sufficient statistical power and
confidence to reliably make correct decisions about the status of a given location.
To determine the number of replicates required, the following questions should be answered
(Alldredge, 1987):
1. What is being compared (i.e., toxicity endpoint, parameter value)?
2. Is the significance criterion directional (is one only interested in whether a station is more toxic
than another, not less toxic as well; i.e., one-tailed test)?
3. What is the level of significance between the expected and actual value of the parameter being
measured?
4. How large a difference is acceptable between the expected and actual value of the criterion being
measured, and with what level of probability?
5. What variability is expected in the data?
Appendix C: Statistical Considerations in Determining the Appropriate Number of Replicate Samples C-3
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There are a number of approaches that can be used to determine the number of replicates required to
achieve a minimum detectable difference at a specific confidence level and power (see Environment
Canada, 1995). While many programs specify a fixed number of replicates per station (often 3-5
replicates), in other cases, this could represent too many or too few replicates for study data quality
objectives. Several factors need to be defined to establish the appropriate number of replicates (see
text box). U.S. EPA (2000c) presents a concise discussion of the relationships of these statistical
considerations. Traditionally, acceptable coefficients of variation vary from 10 to 35%, the power
from 80 to 95%, the confidence level from 80 to 99%, and the minimum detectable relative
difference from 5 to 40% (Earth and Starks, 1985).
Several books on sampling design (e.g., Keith 1993; USEPA 2000b) discuss methods to determine
the appropriate number of replicates needed for a given set of objectives. Table C-l summarizes
statistical approaches for determining the appropriate number of replicate samples needed per station
given different study objectives.
Table C-1. Statistical Formulae for Determining Number of Samples to be Collected for
Environmental Monitoring and Assessment
Study Objective
Formula
Reference
To determine the sample size
required to detect an effect in
an impacted area versus a
control area over time:
a) resampling same sites
before and after impact
and testing if the mean
change in the control area
is the same as that in the
impacted area
b) sampling different sites
before and after impact
and testing if there is no
interaction between area
effect and time effect
n = 2(ta + tp)2 (S/A)2
Green, 1989
n = 4(ta + tp)2 (S/A)2
Green, 1989
where:
n =
S =
A =
tp, =
number of samples for each of
the control and impact areas
standard deviation
magnitude of change required
to be a real effect with
specified power (1-P)
t statistic given a Type I1 error
probability
t statistic given a Type II2 error
probability
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Table C-1 (continued). Statistical Formulae for Determining Number of Samples to be Collected
for Environmental Monitoring and Assessment
Study Objective
Formula
Reference
To determine if the mean
value for an impacted area:
a) differs significantly from a
standard value (e.g.,
sediment guideline)
b) differs significantly from
the mean of a control site
Alldredge, 1987
n >_[ZaJiZp]l + 0. 5 Za2
d2
n >JZaJiZp)! + 0. 25 Za2
d2
where:
n = number of samples
Za = Z statistic for Type I error
probability (e.g., a=0.05)
Zp = Z statistic for Type II error
probability (e.g., p=0.90)
d = magnitude of the difference to
be detected (i.e., effect level)
To determine the number of
samples required to estimate a
mean value (representative of
the area) with a given
statistical certainty
Hakanson, 1984
where:
y = accepted error as a proportion
of the mean value(e.g., y =
0.10)
x = mean value of x; (i = l...n)
Sx = standard deviation
tc = confidence coefficient (e.g.,
90% or to.95
n = number of samples
To determine the number of
samples required to estimate a
mean
n = _(Za/2]V.
Milton et al, 1986
where:
n
Z
a2
a/2
number of samples
Z statistic (standard normal
curve)
variance
probability of a 95%
confidence level
distance between the
center of the lower
confidence and upper
confidence bound
Appendix C: Statistical Considerations in Determining the Appropriate Number of Replicate Samples
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Table C-1 (continued). Statistical Formulae for Determining Number of Samples to be Collected
for Environmental Monitoring and Assessment
Study Objective
To determine the number of
samples required for a
particular power for a normal
distribution (i.e., x > s2)
Formula
n = 104 (tV) K
(R2x2)
where:
n = number of samples
t = t statistic for a desired
confidence level
x = mean value from preliminary
sampling or historical data
s = standard deviation of mean
R2 = percentage coefficient of
variation
K = index of clumping
Reference
Kratochvil and
Taylor, 1981
1 Type I (a) error is the probability of rejecting the hypothesis being tested when it is true.
2 Type II (P) error is the probability of not rejecting the hypothesis being tested when it is false.
Optimizing Sampling
Having estimated the variability in a given parameter or endpoint, and the number of replicate
samples per station that might be necessary to address data quality objectives, one can evaluate the
cost/benefit of collecting and analyzing more or less samples in terms of the overall confidence in a
given decision and the information gained. This is referred to as optimizing the study design (Step 7,
Figure 2-1). Ferraro et al. (1994, 1989) present a method for quantitatively evaluating the optimum
macrobenthic sampling protocol, including the number of replicates («), which has relevance to other
sediment quality studies as well. Their approach helps answer fundamental questions concerning the
design of sediment quality studies such as:
• How large should the sampling unit be?
• How many replicate samples should be taken?
The procedure calculates the "power-cost efficiency" (PCE), which incorporates both the number of
samples («), the cost (field collection effort and lab effort combined) and the expected statistical
power for each alternative sampling scheme. The various sampling schemes consist of different
combinations of sampling gear, gear area, and number of replicates. The method allows determining
the optimum among a set of sampling schemes for detecting differences between reference and test
sites when the statistical model is a t-distribution for comparing two means. The optimum scheme
can be defined as the least costly one capable of reliably (e.g., a = 0.5, 1-P = 0.95) detecting a
desired difference in the means of particular measure between two sites. The approach can be
applied to each parameter of interest and the results aggregated to determine the optimum protocol.
C-6
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Technical Manual
There are four primary steps in assessing the PCE of a suite of alternative sampling schemes:
1. For each scheme, collect replicate samples at paired reference and test sites. The observed
difference in values between the sites is operationally assumed to be the magnitude of the
difference desired to be detected. Alternatively, a percentage of the median (e.g., 20%) for a
given measure calculated across reference stations could be set as the magnitude of the
difference to be detected. In either case, this difference, divided by the standard deviation, is the
"effect size" (ES) of interest.
2. Assess the "cost" (c,), in time or money, of each sampling scheme i at each station. The cost can
include labor hours for sampling, analysis, and recording results.
3. Conduct statistical power analysis to determine the minimum number of replicate samples (n,)
needed to detect the ES with an acceptable probability of Type I (a) and Type II (P) error (e.g.,
a = p = 0.05).
4. Calculate the power-cost efficiency (PCE) for each sampling scheme by:
PCE,. = (n x c)min/ (n; x c,)
where (n x c)min = minimum value of (n x c) among the i sampling schemes. The reciprocal of PCEi
is the factor by which the optimal sampling scheme is more efficient than alternative scheme i.
When PCE is determined for multiple metrics, the overall optimal sampling protocol may be defined
as that which ranks highest in PCE for most metrics in the test set.
Appendix C: Statistical Considerations in Determining the Appropriate Number of Replicate Samples C-7
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APPENDIX D
ADVANTAGES AND DISADVANTAGES OF
DIFFERENT STATION POSITIONING
TECHNIQUES
-------
Technical Manual
Documentation of sampling station location or position is an important aspect of field operations to
ensure that: (1) sampling occurs where intended and (2) someone else (or another sampling team)
could re-sample the same location at a later date. This is particularly critical for trend monitoring
such as that performed by NOAA's Status and Trends Program.
With current technology, a global positioning system (GPS) device is generally the positioning
method of choice because it is usually very accurate, reliable, easy to use, and affordable. However,
occasionally, other positioning methods may be desired or necessary. The following tables,
originally developed under the Puget Sound Estuary Program, summarize most of the positioning
methods that have been used in monitoring studies, including their advantages and disadvantages.
Appendix D: Advantages and Disadvantages of Different Station Positioning Techniques D-3
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Table D-1. Positioning methods appropriate for small water bodies (small embayment, small lakes,
rivers) (modified from PSEP 1997a).
Method
GPS or Navstar
Theodolite
Electronic
Distance
Measurement
instrument
(EDMI)
Total stations
Sextant
Accuracy
± 100m (0.1
to
1 m for
differential
GPS)
10 to 30 s
> ± 1 m
1.5 to 3.0 cm
5 to 7 cm
± 10s
± 3 to 5 m but
variable
Range
no limit
on the
range
200m
to 5 km
3km
without
multiple
prisms
< 5 km
200 m to
5km
Advantages
• Continuous position reports
available worldwide
• System s available comprising a
range of accuracy and cost
• Traditional method, measuring
horizontal angles between known
targets
• High accuracy when applied
successfully
• Inexpensive
• High accuracy
• Compact, portable, rugged
• Relatively inexpensive
• Useable for other surveying
projects
• Not logistically complex,
requiring single onshore site
• Compatible with other uses
• High accuracy when used
nearshore by experienced
operator
• Portable, involving handheld
device
• Rapid, easy to implement
• Easily obtainable
• No shore party necessary
• Inexpensive
Disadvantages
• Site-specific problems due
to military scrambling
• Requires triangulation
between two manned shore
sites or targets
• Requires simultaneous
measurements
• Requires good visibility
which limits area! coverage
• Requires stationary
sampling platform
• Introduces error and
limitations due to reflector
movement and directionality
as well as ground wave
reflection
• Requires good line-of-sight
visibility unless microwave
unit is available
• Requires two shore sites
• Introduces limitations due to
reflector movement and
directionality, prism costs,
and line- of- sight, optical or
infrared range limitations
• Requires simultaneous
measurement of two angles
• Requires good target
visibility
• Requires location and
maintenance of targets for
relocation of site
• Requires calm conditions for
best results
• Orientation of target affects
accuracy
• Has limitations on
acceptable angles
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US Environmental Protection Agency
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Technical Manual
Table D-1 (continued). Positioning methods appropriate for small water bodies (small embayment,
small lakes, rivers) (modified from PSEP 1997a).
Method
Pelorus
RADAR
Autotape
Accuracy
variable
variable
±0.5m
Range
<5 km
30 to
50km
limited
Advantages
• High accuracy when used
nearshore
• Rapid, easy to implement
• Easily obtainable
• No shore party necessary
• Inexpensive
• Standard equipment on ships
• Easily operated
• Yields range and relative bearing
to targets
• High accuracy and precision
• Portable
Disadvantages
• Requires simultaneous
measurement of two angles
• Requires good target
visibility
• Requires location and
maintenance of targets for
relocation of site
• Requires calm conditions for
best results
• Has limitations on
acceptable angles
• Restricts applications by not
being portable
• Requires a target that
reflects microwave signals
• High cost
Appendix D: Advantages and Disadvantages of Different Station Positioning Techniques
D-5
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Table D-2. Positioning methods appropriate for large water bodies (ocean, estuaries, large lakes)
(modified from PSEP 1997a).
Category
GPS or Navstar
Microwave
navigation
systems (e.g.,
Miniranger,
Trisponder,
Racal Microfi,
Del Norte)
Shoran
LORAN-C
Decca HIFIX/6
Variable range
Decca Minifix
Range-azimuth
Satellite
navigation
(SATNAV)
Accuracy
± 100m
(0.1 to
1 m for
differential
GPS)
± 1 to 3 m
± 10m
>± 15m
± 1 m
±0.5°
± 2m
0.02 ° and
0.5m
1 - 10m
Range
no limit on
the range
25 to 80 km
(depends on
height of
transceiver
units)
< 80km
(short range)
up to 300 km
(medium
range)
up to 300 km
(medium
range)
16 to 72 km
> 70km
<5 km
(optical)
30km
(elect)
no limit on
the range
Advantages
• Continuous position reports
available worldwide
• System s available comprising
a range of accuracy and cost
No visibility restrictions
Multiple users
High accuracy
Radio line of sight
Portable, easy system to
operate
• High accuracy
• No visibility or range
restrictions
• Requires no additional
personnel
• Existing equipment
• Relatively inexpensive
• High accuracy and precision
• No visibility restrictions
• Requires no additional
personnel
• Existing equipment
• Inexpensive
• High accuracy and precision
• Light weight equipment
• High accuracy
• Single station
• Circular coverage
• High accuracy
• Single site with minimal
logistics
• Use possible in restricted and
congested areas
• No requirement for shore sites
• Capability for integrating
satellite fixes with other data
sources to improve precision
Disadvantages
• Site- specific problems due to
military scrambling
• Moderately expensive system
• Requires multiple onshore sites
• Cost impacts due to logistics
and security of the necessary
shore units
• Potential source of error due to
signal reflective nulls
• Limited range due to low-
powered shore units
• Limited range
• Requires two shore transmitters
• Incurs interference in some
areas
• Universal coverage not
available
• Used only for repositioning
after employing a more
geodetically precise system to
identify location
• Requires multiple shore sites
• Expensive system
• Uses line-of-sight method
• Relies on map accuracies of
targets
• Decreased accuracy with range
scale
• Expensive system
• User- specific
• Uses line-of-sight method
• Potential source of error due to
signal reflective nulls
• Expensive system
• Continuous coverage
unavailable
• Introduction of error due to
local and atmospheric effects
• Distorted when signal path
crosses polar ice caps
• Requires high initial
development expenditures
D-6
US Environmental Protection Agency
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APPENDIX E
ADVANTAGES, DISADVANTAGES AND
ILLUSTRATIONS OF GRAB AND CORE
SAMPLING DEVICES USED IN SEDIMENT
MONITORING STUDIES
-------
Technical Manual
Table E-1. Advantages and Disadvantages of Commonly Used Grab Samplers
(modified from Klemm etal., 1990; Environment Canada, 1994; PSEP, 1997a; WDE, 1995).
Device
Orange Peel
Smith-Mclntyre
Birge-Ekman,
small
Birge-Ekman,
large
PONAR,
standard
PONAR, petite
Van Veen
Modified Van
Veen (e.g.,
'Ted-Young
grab")
Use
Marine
waters, deep
lakes
Deep lakes,
rivers and
estuaries
Lakes and
marine areas;
soft
sediments,
silt and sand
Lakes and
marine areas;
soft
sediments,
silt and sand
Deep lakes,
rivers and
estuaries;
useful on
sand, silt or
clay
Deep lakes,
rivers and
estuaries;
useful on
sand, silt or
clay
Deep lakes,
rivers and
estuaries;
useful on
sand, silt or
clay; effective
in marine
environments
in deep water
and strong
currents
Lakes and
marine areas
Sample
Depth
(cm)
Oto 18
0 to 4 (in
deep
sand)
Oto 10
Oto 30
Oto 10
Oto 10
Oto 30
Oto 15
Sample
Volume
(L3)
10 to 20
10 to 20
<3.4
< 13.3
7.25
1.0
18 to 75
<18.0
Advantages
• Comes in a range of
sizes
• Reasonable quantitative
samples
• The trigger plates
provide added leverage
essential to its
penetration of substrate
• Handles easily without
winch or crane
• Can be adapted for
shallow water use
• Good for soft sediments,
sand and silt
• Allows subsampling
• Can be adapted for
shallow water use
• Good for soft sediments,
sand and silt
• Allows subsampling
• Most universal grab
sampler
• Adequate on most
substrates
• Large sample obtained
intact, permitting
subsampling
• Good for coarse and firm
bottom sediments
• Adequate for most
substrates that are not
compacted
• Adequate on most
substrates that are not
compacted
• Large sample obtained
intact, permitting
subsampling
• Available in stainless
steel
• Fluorocarbon plastic
liner can help avoid
metal contamination
• Screened bucket cover
helps reduce bow wave
effects
Disadvantages
• Need large boat, powered
winch and calbe line
• Blocking of jaws may cause
sample losss
• Heavy, need boat and power
winch
• Spring loaded jaws,
hazardous
• Inadequate for deep
burrowing organisms
• Restricted to low current due
to light weight and messenger
activation
• May exceed target
penetration depth
• Subsampling may be
restricted by size of top flaps
• Restricted to low current
conditions
• Penetration depth can exceed
desired level due to weight of
sampler
• Heavy; requires winch
• May not close completely,
resulting in sample loss
• Metal frame may contaminate
sample
• Heavy; requires winch
• May not penetrate sediment
to desired depth, especially in
consolidated sediments.
• Susceptible to incomplete
closure and loss of sample.
• Requires more casts to obtain
sufficient sample if many
analyses needed.
• May not close completely,
resulting in sample loss
• May close prematurely in
rough waters
• Metal frame may contaminate
sample
• Heavy; requires winch
• Requires winch
• Relatively expensive
Appendix E: Illustrations of Grab and Core Sampling Devices
E-3
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Table E-1. Advantages and Disadvantages of Commonly Used Grab Samplers
(modified from Klemm etal., 1990; Environment Canada, 1994; PSEP, 1997a; WDE, 1995).
Device
Petersen
Shipek,
standard
Mini Shipek
Use
Deep lakes,
rivers and
estuaries;
useful on
most
substrates
Used
primarily in
marine
waters and
large inland
lakes and
reservoirs;
not useful for
compacted
sandy clay or
till substrates
Lakes, useful
for most
substrates
that are soft
Sample
Depth
(cm)
OtoSO
Oto 10
Oto3
Sample
Volume
(L3)
9.45
3.0
0.5
Advantages
• Provides large sample
• Penetrates most
substrates
• Sample bucket opens to
permit subsampling
• Retains fine-grained
sediments effectively
• Handles easily without
winch or crane from
most platforms
Disadvantages
• Shock wave from descent
may disturb fine-grained
sediment
• Lacks lid cover to permit
subsampling
• May not close completely,
resulting in sample loss
• Metal frame may contaminate
sample
• Restricted to low current
conditions
• May exceed target
penetration depth
• Metal frame may contaminate
sample
• Heavy; requires winch
• Can result in the loss of the
topmost 2-3 cm of very fine,
unconsolidated sediment
• Requires vertical penetration
• Samples small volume
• May lose fine-grained
sediment
• May close prematurely
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Technical Manual
Table E-2. Advantages and Disadvantages of Commonly Used Core Samplers
(modified from Klemm etal., 1990; Environment Canada, 1994; PSEP, 1997a; WDE, 1995; USEPA/ACOE, 1998)
Device/
Dimensions
Fluorocarbon
plastic or glass
tube (3.5 to 7.5
cm inner
diameter (I. D.);
< 120 cm long)
Hand corer
with removable
fluorocarbon
plastic or glass
liners (3.5 to
7.5cm I.D.; <
120 cm long
Box corer
Gravity Corer,
Phleger Corer
(3.5cm I.D., <
50 cm long)
Gravity Corer,
Kajak-
Brinkhurst
Corer (5 cm
I.D., < 70cm
long)
Benthos
Gravity Corer
(6.6, 7.1 cm
I.D. < 3 m
long)
Alpine Gravity
Corer (3.5 cm
I.D.)
Piston Corers
Use
Shallow
wadeable
waters or deep
waters if
SCUBA
available; soft
or semi-
consolidated
deposits
Same as
above except
more
consolidated
sediments can
be obtained
Same as
above but the
depth of the
uncon-
solidated
sediment must
be at least 1 m
Deep lakes
and rivers;
semi-
consolidated
sediments
Deep lakes
and rivers;
Soft fine-
grained
sediments
Soft, fine-
grained
sediments
Soft, fine-
grained, semi-
consolidated
substrates
Ocean floor
and large
deep lakes;
Most
substrates
Depth
Sample
(cm)
Oto 10
Oto 10
Oto 70
Oto 50
Oto 70
0 to 3 m
< 2 m
3 to 20 m
Volume
Sample
(L3)
0.096-
0.44
0.96-0.44
<30.0
<0.48
<1.37
< 10.26
< 1.92
5-40
Advantages
• Preserves layering and
permits historical study of
sediment deposition
• Minimal risk of
contamination
• Rapid; samples
immediately ready for
laboratory shipment
• Same advantages as
fluorocarbon plastic or
glass tube
• Penetrates substrate with
greater ease through use
of handles
• Collects large, undisturbed
sample; optimal for
obtaining intact
subsamples
• Reduces risk of sample
contamination
• Maintains sediment
integrity relatively well
• Penetrates with sharp
cutting edge
• Collects greater volume
than the Phleger Corer.
• Retains complete sample
from tube because the
core valve is fitted to the
core liner
• Fins promote vertical
penetration
• Allows different
penetration depths due to
interchangeable steel
barrel
• Typically recovers a
relatively undisturbed
sediment core in deep
waters
Disadvantages
• Small sample size
necessitates repetitive
sampling
• Small sample size
necessitates repetitive
sampling
• Requires careful handling
to prevent spillage
• Requires removal of liners
before repetitive sampling
• Barrel and core cutter
metal may contaminate
sample
• Difficult to handle
• Relatively heavy; requiring
larger vessel and power
winch to deploy.
• Requires careful handling
to avoid sediment spillage
• Requires repetitive and
time-consuming operation
and removal of liners due
to small sample size
• Same as Phleger Corer
• Requires weights for deep
penetration so the required
lifting capacity is 750 to
1 ,000 kg
• Requires vertical
penetration
• Compacts sediment
sample
• Lacks stabilizing fins for
vertical penetration
• May penetrate non-
vertically and incompletely
• Requires a lifting capacity
of 2,000 kg
• Disturbs sediment stratas
and integrity
• Compacts sediment
sample
• Requires lifting capacity of
>2,000 kg
• Piston and piston
positioning at penetration
may fail
• Disturbs surface (0 to
0.5m) layer
Appendix E: Illustrations of Grab and Core Sampling Devices
E-5
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Table E-2. Advantages and Disadvantages of Commonly Used Core Samplers
(modified from Klemm etal., 1990; Environment Canada, 1994; PSEP, 1997a; WDE, 1995; USEPA/ACOE, 1998)
Device/
Dimensions
BMH-53 Piston
Corer
Boomerang
Corer (6.7 cm
I.D.)
Vibracorer (5.0
to 7.5 cm I.D.)
Use
Waters < 2 m
deep with
extension rod;
soft deposits
Ocean floor
(up to 9,000 m
deep)
Continental
shelf of
oceans, large
lakes; sand,
silty sand,
gravelly sand
substrates
Depth
Sample
(cm)
<2m
1 m
3 to 6 m
Volume
Sample
(L3)
<2
3.52
5.89 to
13.25
Advantages
• Piston provides for greater
sample retention
• Requries minimal
shipboard equipment so
small vessels can be used
• For deep profiles it
effectively samples most
substrates with minimum
disturbance
• Can be used in over 20 m
of water depth
• Portable models can be
operated from small
vessels (e.g. 10 m long)
Disadvantages
• Cores must be extruded
onsite to other containers
• Metal barrels introduce
risk of metal contamination
• Only penetrates 1.2 m
• Requires calm water for
recovery
• Loses 10 to 20% of
sample
• Labor intensive
• Assembly and
disassembly might require
divers
• Disturbs surface (0 to
0.5 m) layer
• Special generator may be
needed
• Heavier models require
larger boat and power
winch to deploy
E-6
US Environmental Protection Agency
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Technical Manual
SOFT BOTTOM
SURFICIAL SAMPLE
HARD BOTTOM
Figure E-1. Some recommended devices for collecting surficial sediments (drawings from
Murdoch and Azcue 1995 and Fredette et al. 1990).
Appendix E: Illustrations of Grab and Core Sampling Devices
E-7
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
PROFILE SAMPLE
SOFT BOTTOM
HARD BOTTOM
Figure E-2. Some recommended devices for obtaining sediment profiles (drawings from Murdoch
and Azcue 1995 and Fredette et al. 1990).
E-E
US Environmental Protection Agency
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APPENDIX F
EXAMPLES OF FIELD FORMS USED TO
DOCUMENT STATION AND SAMPLE
CHARACTERISTICS AND SAMPLE
TRACKING
-------
Technical Manual
NOAA - Chesapeake Bay and Adjacent Tribs. Sediment Toxicity Study - Field Form
Strata
Site Number
Alternate
Date
Time (local)
ANALYSIS
Metals
Organics
AVS
Chem-Grain Size
P450/microtox
Amphipod
Porewater
Benthos-comp.
Benthos-Biomass
Grain Sz&TOC
STATION LOCATION
STATION COORDINATES
GPS
Latitude:
N
.ongitude:
W
SEDIMENT DESCRIPTION
tolor:
Texture:
Odor/sheens:
3enthic Organisms:
WATER QUALITY
Top
Bottom
Temperature
Celsius Temperature
Celsius
Salinity
ppt
Salinity
ppt
Dissolved Oxygen
mg/l
Dissolved Oxygen
mg/l
)onductivity
umhos Conductivity
umhos
Water Depth
m or ft|Secchi Depth
m or ft
Sample Team
OTHER COMMENTS
Appendix F: Examples of Field Forms
F-3
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Example of field form used by the Great Lakes National Program Office:
Field Sampling Log Sheet
Location and Core Information
Station Number:
Date
Time
Primary GPS
Secondary GPS
Latitude
Longitude
Latitude
Longitude
Water Surface Elevation
Water Depth
Core tube Length
Depth of Penetration
Length of Retrieved Core
Loggers Initials
Samplers Initials
Sample Intervals
Sample Number Sample Interval Physical Descritpion of Sample
F-4
US Environmental Protection Agency
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Technical Manual
Example of field form used for site remediation sampling at Naval bases:
TS- Tetra Tech NUS, Inc.
SOIL & SEDIMENT SAMPLE LO€ SHEET
Page , of
Proj«et Site Name: Sample ID Ho.:
Project No.: Sample Location:
Q Surface Soil
0 Subsurface Soil
P Sediment
Q Other;
[] QA Sample Type:
Sampled By:
C.O.C. No,:
Type of Sample;
0 Low Concentration
D High Concentration
GRAB SAMPLE DATA:
Date: j Depth Interval
Time
Method:
MorMtor Reading (ppm):
Cwiw
DMcrlptton {SMid, S-IH, Cl»y, Uteintaw, «c-)
eoniposrre SAMPLE DATA:
Dale: Time Depth Interval
Mtthod:
Monitor Readings
; Range in ppm):
SAMPLE COLLECTION INFORMATION:
Analysis
Color
Doscrlpllon (S»nti, Slit, Clay, Moisture, etc.)
Container Requirements Collected Other
OBSERVATIONS ,' NOTES:
Circle If Applicabfe:
MS.'MSD Duplicate 10 No,:
MAP:
S!gnabira(i}:
Appendix F: Examples of Field Forms
F-5
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APPENDIX G
PHYSICO-CHEMICAL SEDIMENT
CHARACTERIZATION
-------
Technical Manual
1. General Information
It is often necessary or desirable to determine certain physico-chemical characteristics of sediments
in the laboratory, in conjunction with toxicity testing or chemical analysis for inorganic or organic
contaminants. This characterization should include measurement of certain parameters known to
mediate the availability of contaminants in sediment (ASTM, 2000f). Bulk chemical concentrations
alone should not be used to evaluate bioavailability (USEPA, 1998). The following parameters are
generally measured:
• pH (pore water)
• ammonia (pore water)
• total organic carbon
• particle size distribution (e.g., percent sand, silt and clay)
• percent water content
• salinity and hardness of pore water
• conductivity of pore water
Depending on the experimental design and/or study objectives, more extensive characterization may
be necessary. Several additional characteristics which may assist in study implementation, data
interpretation or QA/QC (i.e., assessing sediment integrity, artifact production, optimal extraction
and test procedures) include: sediment biochemical oxygen demand (BOD), sediment chemical
oxygen demand (COD), sediment oxygen demand (SOD), cation exchange capacity (CEC), Redox
(Eh) or oxidation-reduction potential (ORP), total inorganic carbon, total volatile solids, acid volatile
sulfides (AVS), simultaneously extracted metals (SEM), metals, synthetic organic compounds
(pesticides, PCBs, PAHs, and TCDD-dioxin), oil and grease, petroleum hydrocarbons, dissolved
organic carbon (DOC) in the pore water. Measurements of many sediment physicochemical
characteristics use analytical techniques originally developed for soils and waters, and the literature
should be consulted for details regarding recommended methodology (Black, 1965; USGS, 1969;
Plumb, 1981; Page et al., 1982). The following sections provide rationale for making each type of
sediment physicochemical measurement, along with brief descriptions of measurement techniques,
and references for further information and specific procedures.
2. pH
Sediment pH is often one of the single most important factors controlling speciation and equilibria
for many chemicals including sulfides, ammonia, cyanide, and metals, all of which ionize under the
influence of pH. The USEPA ammonia water-quality criterion, for example, is dependent in part on
pH because ammonia toxicity is largely governed by the unionized ammonia fraction which is pH-
dependent (USEPA, 1999). Metal (Cd, Cu, Ni, Pb, and Zn) speciation and bioavailability are also
known to be affected by pH (Schubauer-Berigan and Ankley, et al., 1991; Ho et al. 1999).
Generally, pH is measured using a pH meter consisting of a potentiometer, a glass electrode, a
reference electrode, and a temperature compensating device. A circuit is completed through the
potentiometer when the electrodes are submersed. General purpose process pH electrodes are
available in a wide variety of configurations for in-line and submersion applications. Generally,
electrodes with gel-filled references require less maintenance than electrodes with liquid-filled
references. The latest instruments have microprocessors that automatically calculate and display the
slope. Some older instruments have a percent-slope readout or (and) millivolt readout. For
instruments with a millivolt readout, the measured electrode potential is calculated as the difference
between millivolts measured at the known pH of two buffers.
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
Plumb (1981) and Gonzalez (1995) described a method for measuring pH in sediment using a pH
probe and meter. The probe was inserted into the sediment and pH directly measured after at least a
5 minute equibration time. Electrodes have also been used for direct measurements of pH in
sediment pore water, or in a 1 to 1 mixture of sediment to water (Jackson, 1958). Direct
measurement of sediment pH is also possible using electrodes with "spear tip" designs allowing for
greater penetration into the sample (Burgess, personal communication). Detailed methods for
measuring pH in water and sediment are also described by USEPA (1983;1986b;1987), in USEPA
(1979), and in USEPA (1987), respectively.
3. Ammonia in Pore Water
Nitrogen, a nutrient associated with over-enrichment of aquatic environments, exists in several
forms, including ammonia. Ammonia is highly soluble in water where it is found in an un-ionized
form (NH3) and in an ionized form as NH4+. The extent of ionization is dependent on pH
temperature, and salinity (in seawater). Ammonia in sediments and pore water is generally the result
of microbial degradation of nitrogenous organic material such as amino acids (Ankely et al., 1990).
Pore water concentrations of ammonia as high as 50 mg/L have been measured in otherwise
uncontaminated sediments (Murray et al., 1978; Kristensen and Blackburn, 1987), while ammonia in
pore waters from contaminated sediments can range from 50 to more than 200 mg/L (Ankley et al.,
1990; Schubauer-Berigan and Ankley, 1991).
The toxic effects of ammonia are generally considered to be associated with the un-ionized fraction
(NH3) rather than the ionic components (NH4+ and NH4SO4"), which co-exist in equilibria. This
equilibrium is highly dependent on pH, temperature, pressure, salinity, and ionic concentrations of
ammonia. The toxic un-ionized ammonia fraction can be calculated using known total ammonia
values and measurements of pH, pressure, salinity, and temperature as described by Whitfield (1978)
and Thurstonetal (1981).
USEPA (1983), and APHA (1995) describe five methods available to measure ammonia in the pore
water:
• the titrimetric method
• the ammonia-selective electrode method
• the ammonia-selective electrode method using known addition
• the phenate method
• the automated phenate method.
A preliminary distillation step may be required if interferences are present (APHA, 1995).
Interferences, e.g., sample constituents that interact with procedural reagents, are described in detail
in the APHA (1995) and ASTM (2000g) methods. Once distilled, the sample can be analyzed using
any of the methods listed above.
The distillation and titration methods are frequently used when ammonia concentrations are greater
than 5.0 mg/L. The ammonia-selective electrode method is appropriate when concentrations range
between 0.03 and 1400 mg NH3-N/L. Ammonia readings are calibrated against ammonia standards.
To verify meter readings, confirmatory subsamples can be preserved and analyzed for ammonia using
the standard Nessler technique described in APHA (1995). For the phenate method, APHA (1995)
recommends distillation with sulfuric acid when interferences are present (Bower and Holm-Hansen,
1980). The automated phenate method is suitable for pore waters with ammonia concentrations in
the range of 0.02 and 2.0 mg NH3-N/L.
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Hach Company, Inc. (Loveland, CO) describes the USEPA approved Nessler/distillation method
adapted from APHA (1995). This is a photometric procedure and has been modified for use with
Hach photometers.
4. Total Organic Carbon Content (TOC)
The total organic carbon (TOC) content of sediment is a measure of the total amount of oxidizable
organic material. TOC is the sum of dissolved organic carbon (DOC), particulate organic carbon
(POC) or suspended organic carbon (SOC), and colloids. TOC is an important parameter to measure
in sediments because it is a major determinant of nonionic organic chemical bioavailability (DiToro
et al., 1991). Metal bioavailability is also affected by the amount of TOC present in sediments. TOC
is usually expressed as a percentage of the bulk sediment and is used to normalize the dry-weight
sediment concentration of a chemical to the organic carbon content of the sediment. USEPA
Equilibrium Partitioning Guidelines estimate bioavailability as a function of contaminant
concentration sorbed to sediment organic carbon and contaminant concentration in the pore water
under equilibrium conditions (USEPA, 1998). Recently, the presence of soot carbon from the
combustion of organic carbon (e.g., fossil fuels) has been recognized as a fraction of the TOC in
sediment. Soot carbon may alter the geochemistry and bioavailability of some organic contaminants
(Gustuffsonetal., 1997).
The organic carbon content of sediments has been measured using several methods including: wet
oxidation titration, modified titration, and combustion after removal of carbonate by the addition of
HC1 and subsequent drying. USEPA methods(1986b; 1987), including SW-846 and 430/9-86-004,
are often used to measure TOC. Plumb (1981) recommends one of two methods to separate organic
from inorganic carbon before analyzing for TOC: (a) ignition and using HC1 as the acid for pre-
treating sediment, or (b) differential combustion, which uses thermal combustion to separate the two
forms of carbon.
EPA/ACOE guidance (1998) recommends that TOC analyses be based on high-temperature
combustion rather than on chemical oxidation, because some classes of organic compounds are not
fully degraded by combined chemical and ultraviolet oxidation techniques. Inorganic carbon (e.g.,
carbonates and bicarbonates) can be a significant proportion of the total carbon in some sediments.
Therefore, samples should be treated with acid to remove the inorganic carbon prior to TOC analysis.
The procedure described by the Puget Sound Estuary Program (PSEP, 1997a) is recommended for
TOC analysis because this method uses high-temperature combustion using an induction furnace.
USEPA recommends a similar method using catalytic combustion and non-dispersive infrared
detection (Leonard, 1991) for quantifying TOC.
U.S. EPA acknowledges that several methods for measuring the total organic carbon (TOC) content
of sediments exist (See Nelson and Sommers 1996 for a review). However, acceptable methods must
at a minimum include the following steps:
Sample Collection
• Sediment samples are collected and stored in non-organic containers
Sample Preparation
• Each sediment sample must have macroscopic pieces of shells (e.g., > 1 mm)
removed and then be pulverized and homogenized
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
• Each sediment sample must be treated by direct addition with a strong non-oxidizing
acid (e.g., HCL) for -18 hours to remove inorganic carbon; sample pH should be <2
after acidification (Yamamuro and Kayanne, 1995)
• Each sediment sample must be oven dried following acid treatment (60 - 70° C)
(Weliky et al., 1983; Yamamuro and Kayanne, 1995)
• Each sediment sample must be stored in a desiccator until analysis
• As noted, desiccation is highly recommended, however if not possible a pre- and
post-acidification sample weight should be performed to correct for water uptake
(Hedges and Stern, 1984).
Sample Analysis
• Each post-acidification sediment sample must be analyzed using acceptable
instrumentation
• Instrumentation should have a detection limit of approximately 100 mg/Kg
• Quantification of organic carbon should be based on a sample's weight, measured
before acidification.
Sample QA
A rigorous QA program should be in place to insure acceptable data quality, this may include:
• Performance of duplicate analysis on a subset of samples with the relative percent
difference (RPD) between replicates below 30%
• Performance of analyses on certified standard reference materials (SRM) (e.g.,
NIST)
5. Particle Size Distribution (Percent Sand, Silt, and Clay)
Particle size is used to characterize the physical characteristics of sediments. Because particle size
influences both chemical and biological characteristics, it can be used to normalize chemical
concentrations and account for some of the variability found in biological assemblages (USEPA
1998) or in laboratory toxicity testing (USEPA, 2000d; Hoss et al., 1999). Particle size can be
characterized in varying detail. The broadest divisions that generally are considered useful for
characterizing particle size distributions are percentages of gravel, sand, silt, and clay. However,
each of these size fractions can be subdivided further so that additional characteristics of the size
distribution are determined (PSEP, 1996).
Particle size determinations can either include or exclude organic material. If organic material is
removed prior to analysis, the "true" (i.e., primarily inorganic) particle size distribution is
determined. If organic material is included in the analysis, the "apparent" (i.e., organic plus
inorganic) particle size distribution is determined. Because true and apparent distributions may
differ, detailed comparisons between samples analyzed by these different methods are questionable.
Therefore, if comparisons among samples between studies is desired, sediment particle size should
be measured using consistent methods (PSEP, 1996).
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Sediment particle size can be measured by a number of different methods (Allen, 1975; Plumb, 1981;
PSEP, 1996; ASTM, 2000a). The best method will depend on the particle properties of the sample
(Singer et al., 1988). Particle size distribution is often determined by either wet sieving the sample
(USEPA, 1979; Plumb, 1981; PSEP, 1996; Singer et al., 1988), the hydrometer method (Day, 1965;
Patrick, 1958), the pipet method (USGS, 1969; Rukavina and Duncan, 1970), settling techniques
(Sandford and Swift, 1971), and X-ray absorption (Duncan and Lattaie, 1979; Rukavina and Duncan,
1970). The pipet method may be superior to the hydrometer method (Sternberg and Creager, 1961).
Combinations of multiple methods may provide refined measurements of particle size distribution.
Gee and Bauder (1986) used sieving and pipetting after soluble salts were removed. Gonzalez (1995)
used a combination of sieve and hydrometer methods. Folk (1968) and Buchanan (1984) discuss
additional methods to measure particle size.
Recommended methods for measuring sediment particle size distribution are those of PSEP (1996)
and USEPA (1995). Percent gravel, sand, silt, and clay are determined as apparent distribution using
a minimum sediment sample size of 100 g taken from a homogenized sediment sample (see Section
4.4). Organic matter should be removed prior to analysis by oxidation using hydrogen peroxide.
Wet-sieving followed by dry sieving (mechanical shaking) separates the two coarse particle size
groups. The silt-clay fraction is subdivided using a pipet technique that depends upon the differential
settling rates of the two different particle size fractions. All fractions are dried to a constant weight.
Cooled samples are stored in a desiccator and weighed.
To obtain an accurate determination of particle sizes for the fine fraction, the Coulter (particle size)
counter method may be employed (McCave and Jarvis, 1973; Vanderpleog, 1981). This method
gives the fraction of particles with an apparent spherical diameter. In a review of the available
methods, Swift et al. (1972) found the Coulter counter method to be the most versatile method
overall; however, it does not provide settling information. Another potential method for determining
the particle size distribution of a very fine fraction is through the use of electron microscopy
(Leppard et al., 1988). Collection techniques for very fine material can result in aggregation of
larger colloidal structures (Leppard, 1986; Leppard et al., 1988). In general, particle settling methods
are preferred to sediment sizing methods.
6. Percent Water or Moisture Content
Water content is a measurement of sediment moisture usually expressed as a percentage of the whole
sediment weight. It is known to influence toxicity and is used to aid in the interpretation of sediment
quality investigations. Sediment moisture content is measured as the difference between wet weight
of the sediment and dry weight following oven drying at 50 to 105°C to a constant weight. Percent
water is used to convert sediment concentrations of substances from wet-weight to a dry-weight.
Methods for determining moisture content are described by Plumb (1981) and Vecchi (1999).
Additional methods are provided in USEPA (1987).
7. Salinity of the Pore Water (Marine Sediments)
Salinity is a measure of the mass of dissolved salt in a given mass of solution. The most reliable
method to determine the true or absolute salinity is by complete chemical analysis. However, this is
time consuming and costly. Therefore, indirect methods are more suitable. Indirect methods include
conductivity, density, sound speed, or refractive index (APHA, 1995). Salinity is then calculated
from the empirical relationship between salinity and the indirect measurement. Conductivity
measurements have the greatest precision, but respond only to ionic solutes (APHA, 1995). Density
measurements respond to all solutes. APHA (1995) recommends the electrical conductivity method,
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
because it is sensitive and easily performed. APHA (1995) also recommends the density method,
using a vibrating flow densitometer. USEPA (1986) methods should also be consulted.
A salinity refractometer can be used for quick readings of salt density in solutions such as sea water.
These refractometers are easy to read, non-corrosive and lightweight. They have dual scales and an
adjustable focus. Temperature and non-temperature compensating refractometers are available. Most
refractometers are accurate to 1 ppt and read specific gravity (1.000 to 1.070 in .001 divisions) and
parts per thousand (0-100 in 1 ppt divisions).
8. Conductivity of the Pore Water (Fresh Water Sediments)
Conductivity is a measure of the ability of an aqueous solution to carry an electric current. This
ability is dependent on the presence of ions in the solution, the concentration of the ions, their
mobility and valence, and temperature. Solutions of inorganic compounds are usually good
conductors while those of organic compounds are usually poor conductors. Conductivity is enhanced
by calcium, potassium, sodium, and magnesium chlorides and sulfides.
Meters can be used to measure the degree to which electrical current can travel through water. The
unit of measure is 1 mS/m = 1 millisiemens/meter or 1 |_iS/cm = 1 microsiemens/cm. The reading
indicates the amount of ions in the water. While traditional chemical tests for hardness measure
calcium and magnesium, they fail to provide an indication of other ions (e.g., sodium). The
conductivity meter provides a much better measure of ionic strength.
9. Acid Volatile Sulfide (AVS)
Measurement of acid volatile sulfides (AVS) and simultaneously extracted divalent metal (SEM)
concentrations associated with AVS extraction can provide insight into the bioavailability of metals
in anaerobic (anoxic) sediments (DiToro et al., 1990; Ankley et al., 1996). AVS is the reactive solid-
phase sulfide fraction that is extracted by cold hydrochloric acid. AVS appears to affect the
bioavailability of most divalent metal ions as the sulfide ions have a high affinity for divalent metals.
This affinity results in the formation of insoluble metal sulfides with greatly reduced bioavailability.
AVS concentrations in freshwater and marine sediments can range between < 0.1 and > 50 (imol
AVS/g of sediment (DiToro et al., 1990).
The bioavailability of metals in sediments has been predicted by comparing the molar concentration
of AVS to the molar concentration of SEM (methods described below). If AVS is greater than SEM,
the metals are bound in sulfide complexes with greatly limited bioavailability. However, if AVS <
SEM, metals may or may not be toxic due to other controlling factors (e.g., TOC).
The easily extractable sulfide fraction can be measured using the acid purge and trap technique. The
sample sulfide is solubilized in cold hydrochloric acid. The analytical method involves conversion of
sulfides to aqueous H2S. This may be measured with a sulfide probe or by following a wet chemistry
method. In the latter method, silver sulfide is precipitated in a gas-tight assembly and flushed with
nitrogen to eliminate oxidation. The precipitate is filtered, dried, and weighed. The weight is
compared with the weight obtained from a non-acidified sample, and the difference is attributed to
the AVS fraction (DiToro et al., 1990).
10. Simultaneously Extracted Metals
A model for predicting toxicity from divalent trace metals (DiToro et al., 1990) is based on the
binding of these metals to AVS. Where the sum of the moles of the SEM, including Ag, Cd, Cu, Ni,
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Pb, and Zn is exceeded by the molar concentration of AVS, the metals are insoluble and largely
unavailable to biota. The extraction of AVS and metals should be achieved using a single
methodology to ensure that recoveries associated with each measure are consistent. Simultaneous
extraction improves the efficiency of the methodology.
SEM can be measured in filtered aliquots by atomic absorption methods (DiToro et al., 1990).
Recent SEM analysis methods use inductively coupled plasma atomic emission spectrometry (ICP-
AES; Berry et al., 1999). Other methods for analysis of metals are described in Section 11 below.
11. Metals
Low levels of trace metals occur naturally in the environment but highly elevated levels in sediment
are generally associated with anthropogenic contaminant loads. Metals are partitioned in sediments
as soluble free ions, soluble organic and inorganic complexes, easily exchangeable ions, precipitates
of metal hydroxides, precipitates with colloidal ferric and manganic oxyhydroxides, insoluble
organic complexes, insoluble sulfides, and residual forms (Gambrell et al., 1976).
Current instrument methods available for the analysis of trace metals include electrochemistry (e.g.,
differential pulse polarography), spectrophotometry (e.g., silver diethyldithiocarbamate), atomic
absorption spectrophotometry, atomic emission spectrophotometry, x-ray fluorescence (XRF), and
neutron activation (PSEP 1997c). The most commonly used instrumental method to analyze
sediments for metals is atomic absorption spectrophotometry (PSEP, 1991 c). Inductively coupled
plasma mass spectrometry (ICP-MS) or ICP-AES allow for simultaneous determination of many
metals at sub-ppb levels with little pretreatment (Crecelius et al., 1987; Berry et al., 1999).
The concentration of salt in marine or estuarine samples may interfere with metals analyses
(USEPA/ACOE, 1998). Therefore, acid digestion and atomic absorption spectroscopy should be
coupled with an appropriate technique to control for this interference. Methods in USEPA (1986b)
are recommended for the analysis of mercury in sediments and EPRI (1986) methods are
recommended for the analysis of selenium and arsenic. EPA methods for cadmium, hexavalent
chromium, copper, lead, mercury, nickel, selenium, silver, and zinc are described by USEPA
(1986b). PSEP (1991 c) suggests that mercury can be extracted using vacuum distillation and
analyzed by gas chromatography/mass spectrophotometry.
12. Synthetic Organic Compounds (Pesticides, PCBs, TCDD-Dioxin)
Analytical techniques for measuring organic compounds require five general steps: drying the
sample, extraction, drying the extract, clean up of the extract, and analysis of the extract. PSEP
(1997b) recommends centrifugation or sodium sulfate to dry the sample and a solvent extraction,
with application of shaker/roller, or sonication. Sample drying with sodium sulfate is recommended
for samples weighing approximately 10 grams (after overlying water is decanted). The sediment and
sulfate mixture is extracted and the extract is processed (MacLeod et al., 1985).
Soxhlet® extraction (USEPA, 1986b) involves distillation with a solvent such as acetone,
dichloromethane/methanol (2:1), dichloromethane/methanol (9:1), and benzene/methanol (3:2).
USEPA (1983) recommends sonication with solvent mixtures and a 30-gram subsample of sediment.
Drying the extract can be accomplished through separatory funnel partitioning as needed to remove
water and sodium sulfate or by using a Kuderna-Danish apparatus and rotary evaporation with
purified nitrogen gas for concentration to smaller volumes (PSEP, 1997c). Using the separatory
funnel partitioning method, the wet sample is mixed with methanol and centrifuged. The supernatant
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
is decanted and extracted later. Extraction of the sample is continued using less polar solvents and
the water/methanol and solvent extracts are combined and dried.
According to PSEP (1997c) elemental sulfur can be removed from the sediment sample with
vigorous mechanical agitation using a Vortex or Genie® or using activated copper. Organic
interferences can be removed with gel permeation chromatography (GPC) described in USEPA
(1983), bonded octadecyl columns (PSEP, 1991 c), high performance liquid chromatography (HPLC)
described by Metro (1981), silica gel (PSEP, 1991 c), or alumina (USEPA, 1983). Instrumental
analyses for volatiles and semivolatiles and pesticides/PCBs are performed using gas
chromatography/mass spectrophotometry (GC/MS) and gas chromatography/electron capture
detection (GC/ECD), respectively (PSEP, 1997b; Burgess and McKinney, 1997).
13. Oil and Grease
Oil and grease tests for sediments measure material recovered that is soluble in a nonpolar solvent
under acidic conditions. Oil and grease compounds are substances such as hydrocarbons, vegetable
oils, animal fats, waxes, soaps, and greases. Many solvents can dissolve other substances (e.g. sulfur
compounds, organic dyes, and chlorophyll). Therefore, oil and grease is operationally defined by the
solvent used and the analytical method used to perform the analysis. There are two basic methods
used to analyze oil and grease: the gravimetric technique and the IR (infrared spectrophotometer)
technique. Both are described by PSEP (1996).
14. Petroleum Hydrocarbons and Polycyclic Aromatic Hydrocarbons
Petroleum hydrocarbons are oil and grease constituents which remain in solution after contact with
silica gel. Petroleum distillates, also called hydrocarbons or petrochemicals, refer to a broad range of
compounds which are extracted by distillation during the refining of crude oil. During the fractional
distillation of petroleum, crude oil is heated to allow various compounds to turn from liquid into gas
and then captured as they rise, cool, and condense. Lighter, more volatile compounds rise higher
before they condense and are collected on distillation trays. Heavier, less volatile compounds such as
diesel fuel and oil are collected on lower distillation trays. Waxes and asphalts are collected from the
bottom after the other products have volatilized.
Petroleum distillates contain both aromatic hydrocarbons (carbon rings) and aliphatic hydrocarbons
(straight carbon chains). The chemical structure of the hydrocarbon largely defines the nature and
behavior of these compounds. Aromatic hydrocarbons are the most toxic compounds found in
petroleum products. Most aromatic hydrocarbons are chronic toxins and known carcinogens.
Aromatic compounds are found in all crude oils and most petroleum products. Many aromatic
hydrocarbons have a pleasant odor and include such substances as naphthalene, xylene, toluene, and
benzene. Aliphatic hydrocarbons are flammable and may be explosively flammable. Aliphatic
hydrocarbons include methane, propane, and kerosene.
Aromatic and aliphatic hydrocarbons were analyzed in sediments by Page et al. (1995 a, b). Sediment
samples were spiked with the appropriate surrogates, mixed with equal amounts of sodium sulfate to
dry the samples, and extracted with a methylene chloride acetone mixture (Method 3550, USEPA,
1986b). The concentrated extracts were partitioned on an alumina column into saturated and
unsaturated hydrocarbon fractions (Method 3611, USEPA, 1986b). The fractions were concentrated
using the appropriate pre-injection volume, spiked with the appropriate internal standards, and
analyzed by gas chromatography with flame ionization detection (GC/FID) and gas chromatography
with mass spectrometry detection (GC/MS) operating in the selected ion monitoring (SIM) mode.
The method of internal standards (Method 8000, USEPA, 1986b) using the average relative response
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factors generated from the linear initial calibration was used to quantify the target compounds. All
data were corrected for the recovery of the appropriate surrogate compound. Their relative
abundances could then be used for identification and quantification purposes.
TPH (total petroleum hydrocarbons) and PAH (polycyclic aromatic hydrocarbons) have also been
analyzed by first acidifying the sample with concentrated hydrochloric acid and then extracting
hydrocarbons with a mixture of methanol and hexane. The hexane extracts were then spiked with an
internal standard and analyzed by GC-FID for TPH content and by GC/mass spectrometry (MS) for
PAH analysis.
Kaplan et al. (1996) extracted hydrocarbons using anhydrous Na2SO4 with methylene chloride and
sonication. The total solvent extract was then concentrated with Kuderna-Danish equipment. The
concentrate was further concentrated using a gentle stream of dry nitrogen. An aliquot was then
injected directly into the gas chromatography.
15. Total Sulfides
Total sulfides represent the combined amount of acid-soluble H2S, HS", and S2" in a sample. Sulfides
are often measured because they are common in some sediments, particularly those that are anoxic,
and they can be toxic to aquatic organisms. PSEP (1996) describes a method to measure total
sulfides in sediments. Oxygen is removed from the sample using nitrogen gas, methyl orange and
hydrochloric acid is added, and the mixture is heated. Amine solution and iron chloride are added to
develop a colorimetric reaction product and sample absorbance is measured spectrophotometrically.
Methods for measuring sulfides in aqueous samples include: potentiometric methods described by
ASTM (2000e) and APHA (Method 4500, 1995). Sulfide ions are measured using a sulfide ion-
selective electrode in conjunction with a double-junction, sleeve type reference electrode (Phillips et
al., 1997). Potentials are read using a pH meter or a specific ion meter having a direct concentration
scale for the sulfide ion. Samples are treated with sulfide anti-oxidant buffer which fixes the solution
pH at a high alkaline level and retards air oxidation of sulfide ion in solution. This ensures that the
sulfide measured represents total sulfides as S= ion and rather than the HS" or H2S found at lower pH
values (see pH, Section 2 in this Appendix).
APHA (Method 4500, 1995) provides qualitative as well as quantitative methods to determine
aqueous sulfide concentrations. Qualitative methods include the antimony test, the silver-silver
sulfide electrode test, the lead acetate paper test, and the silver foil test. Quantitative methods
include the photometric method, the automated photometric methylene blue colorimetric methods,
and the iodometric titration method for standardizing stock solutions.
16. Sediment Oxygen Demand (SOD)
Sediment can exhibit significant rates of oxygen uptake attributable to either: (1) a benthic ecosystem
supported by soluble organic substances in the water column, (2) naturally occurring sediments
derived from aquatic plants and animals, and (3) detritus discharged into the water body by natural
runoff. When numerical modeling is required to predict dissolved oxygen concentrations, the rate of
dissolved oxygen consumed by the benthic ecosystem is defined as the sediment (benthic) oxygen
demand (SOD) in g O2/m2-day.
Two approaches for measuring SOD were reviewed by Truax et al. (1995) including in-situ
respirometry and laboratory respirometry methods. Numerous techniques have been developed for
each approach. Generally, in-situ methods are considered more credible than laboratory
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measurements although both apply the same technique. A given amount of sediment is enclosed in a
chamber with a known water volume and oxygen uptake is measured over time. The SOD rate is
then calculated based on the area of the enclosed sediment, the volume of water in the chamber, and
the rate of uptake.
In situ sediment oxygen demand measurement method were described by Uchrin and Ahlert (1985).
A cylindrical respirometer, a dissolved oxygen probe with stirring mechanism, and a dissolved
oxygen meter were used. Ambient dissolved oxygen was measured using the probe/meter as well as
by using the Winkler method (APHA, 1995) in the laboratory to determine the effect of respiration
on total dissolved oxygen uptake. The respirometer was deployed in a level area at the bottom of the
water body. Dissolved oxygen were recorded initially and at 15-minute intervals thereafter to
determine the SOD rate.
17. Sediment Biochemical Oxygen Demand (BOD)
Biochemical oxygen demand (BOD) is a measure of the dissolved oxygen consumed by microbial
organisms while assimilating and oxidizing the organic matter in a sample (PSEP, 1996). The test is
an empirical methodology in which standardized laboratory procedures are used to determine the
relative oxygen uptake of environmental samples. The test measures the amount of molecular
oxygen used during a specified incubation period to biochemically degrade organic material and to
oxidize reduced forms of nitrogen (APHA, 1995).
Plumb (1981) described a method to analyze BOD in sediments using freshwater bacteria as a "seed"
and buffered distilled water as dilution water. PSEP (1996) described an alternative procedure to
analyze BOD in marine sediments using marine bacteria as the "seed" and filtered, oxygenated
seawater as the dilution water. USEPA (1987) methods should also be consulted.
18. Sediment Chemical Oxygen Demand (COD)
Chemical oxygen demand (COD) is a measure of the oxygen equivalent of organic matter content in
a sample that is susceptible to oxidation by a strong chemical oxidant at elevated temperature and
reduced pH. The test was devised to augment the biochemical oxygen demand test. Chemical
oxygen demand can be related empirically to biochemical oxygen demand, organic carbon, or total
volatile solids (PSEP, 1996).
PSEP (1996) described a method for analyzing sediment COD using a closed reflux/colorimetric
method. DiChromate (Cr2O7) ions are used to oxidize organic matter to carbon dioxide and water
and to provide oxygen. The dichromate ions remaining after the reaction are measured by titration
and the amount of oxygen consumed is then calculated.
Four standards procedures for measuring COD in water are available in APHA (1995): the open
reflux method, the closed reflux method, the titrimetric method, and the closed reflux/colorimetric
method. USEPA (1983) methods for the colorimetric and titrimetric method are described in USEPA
(1979). Semi-automated methods are described in USEPA (1993).
Hach (Loveland, CO) has modified the EPA approved dichromate reflux method and the reactor
digestion method. The methods are photometric and are adapted for use with Hach photometers.
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19. Cation Exchange Capacity of Sediments
Cation exchange capacity (CEC) is a parameter that provides information relevant to metal
bioavailability studies (Black, 1965). Cations or positively charged elements (such as calcium,
magnesium, hydrogen, and potassium), are attracted to negatively charged surfaces of clay and
organic matter. There is a continuous exchange of cations between sediment and water. CEC is a
measure of the sediment's ability to retain cationic elements. It is also a measure of clay activity and
mineralogy, which is used to calculate mineralization rates, leaching rates, and to predict interactions
with contaminants. The degree of CEC is dependent on the kind and amount of suitable surfaces
such as organic matter and clay. High cation exchange capacities are associated with high clay
contents and high organic matter and changes in CEC are typically associated with changes in
organic carbon content and pH of the sediment. Organic matter generally supplies a greater number
of exchange sites than clay particles.
Various methods have been recommended to determine bioavailable fractions of metals in sediments
(Chao and Zhou, 1983; Crecelius et al., 1987; Kersten and Forstner, 1987; DiToro et al., 1990).
CEC can be measured by treating samples with ammonium acetate so that all exchangeable sites are
occupied by NH4+ ion, digesting the samples with sodium hydroxide during distillation, and titrating
to determine the ammonium ion concentration. The amount of exchangeable cations are expressed as
milliequivalents of ammonium ion exchanged (meq) per 100 g of dried sample. More detailed
methods are provided in Bascomb (1964), Black (1965), Klute (1986), and USEPA (1986b).
20. Redox Potential (Eh) of Sediments
Redox (Eh) is a measure of the oxidation-reduction potential (ORP) of sediments. Measurements of
Eh are particularly important for metal speciation and for determining the extent of sediment
oxidation. Eh values below approximately -100 millivolts would indicate biologically important
sulfide concentrations. Some trace metals form insoluble complexes with sulfides. These metal-
sulfide complexes bind the metals in a form that is not bioavailable. Since free ionic metals are
generally thought to possess the greatest toxicity potential, it is important to understand conditions
which control binding dynamics, such as pH and Eh.
Potentiometric measurements of Eh using a millivolt reader can be obtained with a platinum
electrode relative to a standard hydrogen electrode (Plumb, 1981). APHA (1995) does not
recommend the standard hydrogen electrode as it is fragile and impractical. Instead, their method
uses a silver-silver-chloride or calomel reference electrode. APHA (1995) recommends a graphite
rather than platinum electrode for sediments. Once the Eh equilibrium is reached, the difference
between the platinum or graphite electrode and the reference electrode is equal to the redox potential
of the system. For a more detailed explanation on how to calculate the Eh potential see APHA
(1995). Gonzalez (1995) also describes a detailed method that can be used to measure sediment Eh.
There are a number of problems associated with the accurate measurement and interpretation of Eh
in sediments, particularly in marine sediments. Therefore, considerable attention should be paid to
the use of proper equipment and techniques. Some of the problems identified by Whitfield (1969)
and Mudroch and Azcue (1995) include measurement inaccuracy due to disturbance of the sediment
sample during insertion of the electrode, instability and poor reproducibility of the measurements and
differential responses of platinum electrodes under different environmental conditions. A
comprehensive description of the limitations of sediment Eh measurement is beyond the scope of this
document. Rather, it is recommended that published studies on the problems associated with
measuring and interpreting sediment Eh be consulted before any attempt is made to measure these
parameters in sediment samples (Berner,1963; Morris and Stumm, 1967; Whitfield, 1969; Tinsley,
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
1979; Bates, 1981). The recommended procedure for measuring pH and Eh in the field are described
in detail in the table below:
Table G-1. General procedures for measurement of Eh in bottom sediments (from Murdoch and
Azcue 1995).
Equipment and solutions used in the measurements:
• A portable, battery-operated pH/Eh meter, batteries, and a power cord for recharging the meter.
• Combination glass and platinum electrodes or other electrodes suitable for the measurements.
• Plastic test-tube-shaped containers or other containers for storing the electrodes in solutions during transport in the
field.
• Commercially-available or laboratory-prepared pH buffer solutions (pH 4 and 7) in plastic bottles with lids.
• Freshly-prepared solution for calibration of Eh electrode in a plastic bottle with a tight lid.
• Freshly-prepared solution of saturated potassium chloride for storage of the electrodes.
• Other solutions necessary for proper functioning of electrodes as outlined by manufacturers.
• Distilled water and wash bottle for storing and rinsing the electrodes between measurements.
• Several small and larger plastic beakers for holding solutions, rinsing electrodes, etc..
• Support stands, rods, clamps to secure electrodes in solutions and during measurements.
• Large plastic containers for storage and transport of used buffers and Eh-calibration solutions.
• Notebook and pens, soft paper tissue.
Preparation of equipment before the field trip:
• Check batteries of the portable pH/Eh meter and replace/recharge them, if necessary.
• Prepare calibration solutions.
• Check and test the pH and Eh electrodes.
• Mark the electrodes vertically at desired intervals for insertion into the sediment samples.
• Store the electrodes according the manufacturers instructions.
• Pack all equipment for transport to the field and take spare electrodes if available.
Measurements in the field:
• Allocate a space where measurements will be carried out. Within this space, all equipment should be assembled,
checked for proper functioning, and prepared for measurement of the first sample.
• Place grab sampler and sediment cores with recovered sediment in such a way that they will remain steady without
disturbing the sediment samples during the measurements.
• Insert electrodes carefully into the undisturbed sediment samples to avoid any air. contamination, particularly around
the Eh electrode. Care must be taken not to generate any open space between the electrode and the sediment. Proper
insertion of the electrode without disturbing the sediment is the most important step in measuring the Eh.
• Insert electrodes into the sediment to the depth marked. Switch the pH/Eh meter to the pH scale and the value
recorded within 1 minute after inserting the electrode into the sample. Switch the meter to the mV scale for
recording the Eh value. The potential usually drifts considerably over the first 10 to 15 minutes, and then stabilizes.
After stabilization, record the mV value. In measuring Eh of sediments from waters with low ionic strength, such as
most freshwater bodies, it is recommended to "acclimatize" the electrodes in the water prior to measurement,
particularly the electrodes that were stored in saturated potassium chloride solution. This will reduce the drifting of
the potential after inserting the electrode into the sediment.
• Remove both electrodes, wash them with distilled water to remove all adhering sediment particles, and dry them
gently with a soft paper tissue.
• Calibrate the electrodes after each five measurements. The electrodes may need less frequent calibration if pH and
Eh are being measured in a sediment core.
21. Total Inorganic Carbon
Inorganic carbon has been measured as a complement to microbial activity (Bregnard et al., 1996), to
determine the fate of an organic contaminant in biodegradation studies (West and Gonsior, 1996),
and to determine the % carbon unaccounted for in fate transport predictions of hydrophobic
contaminants (Tye, et al., 1996). Often the total inorganic carbon (TIC) fraction in samples is many
times greater than the TOC fraction and presents an interference in the measurement of TOC. There
are several options to eliminate TIC interferences when trying to measure TOC. One option is to
compensate for the 1C interference by measuring total carbon (TC) and total inorganic carbon (see
Section 4 in this Appendix). The difference between the two is the TOC.
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TIC is determined by acidifying the sample to convert the inorganic carbon (i.e., carbonates,
bicarbonates, and dissolved CO2) to carbon dioxide. Carbon dioxide is purged from the sample and
then detected by a non-dispersive infrared detector (NDIR) calibrated to directly display the mass of
carbon dioxide measured. This mass is proportional to the mass of TIC. Other instrumentation for
the analysis of TIC is described in West and Gonsior (1996) and Tye et al. (1996).
22. Total Volatile Solids (TVS)
Total volatile solids represent the fraction of total solids that are lost on ignition at a higher
temperature than that used to determine total solids. Total volatile solids are used as a crude estimate
of the amount of organic matter in total solids (PSEP, 1996). In this regard, total volatile solids are
often measured instead of, or in addition to, organic carbon content.
Total volatile solids are operationally defined by ignition temperature. Total volatile solids content
does not always represent the organic content of a sample because some organic material may be lost
at the drying temperature and some inorganic material (e.g, carbonates, chlorides) may be lost at the
ignition temperature. Because of the temperature dependence of total volatile solids, valid interstudy
comparisons require the use of standardized drying and ignition temperatures (PSEP, 1996).
Total volatile solids measurements are generally made by igniting the sediments at 550 ± 10°C until a
constant weight is achieved and reporting the percent ash-free dry weight (McLeese et al., 1980;
APHA, 1995; Keilty et al., 1988a). Plumb (1981) and PSEP (1996) describe standard methods for
determining the total volatile solid content of sediments. Additional methods are provided in USEPA
(1987).
23. Dissolved Organic Carbon in Pore Water
Dissolved organic carbon (DOC) often consists of humic substances and is the fraction of the organic
carbon pool that is dissolved in water and passed through a 0.45 |im glass fiber filter. DOC is an
indicator of the chemically reactive organic fraction and accurately measures the dissolved organic
load. Sediment pore waters can be rich in humic acids. Fifty to 90% of the pore water DOC can be
colloidal which is a significant factor because organic chemicals will preferentially partition to pore
water DOC (Resendes et al., 1992; Burgess et al., 1996).
Hermann (1996) and Gilek et al. (1996) measured DOC using a TOC apparatus and infrared
detection of CO2. Borga et al. (1996) measured DOC using atomic emission spectrometry (ECP-
AES). The APHA (Method 5310, 1995) methods for total organic carbon which can be applied to
the measurement of DOC are (a) the combustion-infrared method; (b) the persulfate-ultraviolet
oxidation method; and (c) the wet-oxidation method. Adjustments for inorganic carbon interference
may be required (see Section 21 in this Appendix).
24. Alkalinity and Hardness of the Pore Water (Fresh Water Sediments)
Alkalinity is defined as the acid-neutralizing (i.e., proton-accepting) capacity of water. It is the sum
of all the titratable bases and a measure of the quality and quantity of constituents in the pore water
that result in a shift in the pH toward the alkaline side of neutrality. The measured value may vary
significantly with the pH end-point used. Studies have shown that effects of certain contaminants
such as metals are influenced by alkalinity as it alters speciation and bioavailability.
APHA (1995) recommends a color-change titration method to measure alkalinity which is also
described by ASTM (2000h). The sample is titrated with standard alkali or acid to a designated pH
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Methods for Collection, Storage, and Manipulation of Sediments for Chemical and Toxicological Analyses
and the endpoint is determined electrometrically or by the color change of an internal standard. In
addition, ASTM (2000h) describes two additional methods: (1) a titration curve is developed to
identify inflection points, a standard acid of alkali is added to the sample by small increments and pH
is recorded after each addition, and the total volume of acid or alkali is plotted against the observed
pH values; and (2) pH is determined, standard acid is added to lower the pH to 4.0 or less, the
solution is boiled with hydrogen peroxide, and titrated while hot to the phenolphthalein endpoint or
when cooled electrometrically with standard alkali to pH 8.2, the desired endpoint. The color-change
titration method is most commonly used. Hach (Method 8202) has developed a portable water
chemistry kit based on the APHA (1995) color-change titration method and an additional method
using sulfuric acid with a digital titrator (Hach, Method 8203).
Hardness is the concentration of metallic cations, with the exception of alkali metals, present in water
samples. Generally, hardness is a measure of the concentration of calcium and magnesium ions in
water. Hardness is usually expressed as a calcium carbonate equivalent in mg/L. Like alkalinity,
hardness alters speciation and bioavailability of certain contaminants particularly many metals.
AHPA (Method 2340, 1995) describes two methods to measure hardness: (1) the calculation method
and (2) the EDTA titrimetric method. ASTM (2000i) describes the APHA (1995) EDTA titrimetric
method. Calcium and magnesium ions in water are sequestered by the addition of EDTA. The
endpoint of the reaction is measured by means of Chrome Black T3, which is red in the presence of
calcium and magnesium and blue when both are sequestered. APHA recommends the calculation
method because it is more accurate. The method uses direct determinations of calcium and
magnesium to determine hardness. Hach has developed portable water chemistry kits (Methods
8222, 8204, 8030, 8226, 8213, 8338, 8329) for a variety of hardness determinations using a
spectrophotometer or titration methods with a decision tree for selecting the appropriate procedure.
Three of the Hach methods (1992) were adapted from APHA (Method 2340, 1995): the buret and
0.020 N titrant method (8222); the ManVer 2 buret and 0.020 N titrant method (8226); and the buret
titration method (8338). The APHA EDTA titration method is most often used.
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