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EPA Methods for Measuring the 

Toxicity and 

Bioaccumulation of 

Sediment-associated 

Contaminants with 

Freshwater Invertebrates 
EPA 600/R-94/024 

June 1994 

Methods for Measuring the Toxicity and 

Bioaccumulation of Sediment-associated 

Contaminants with Freshwater 

Invertebrates 

Office of Research and Development 

U.S. Environmental Protection Agency 

Duluth, Minnesota 55804 
Disclaimer 

This document has been reviewed in accordance with U.S. Environmental Protection Agency Policy and approved for publication. Mention of trade names or 

commercial products does not constitute endorsement or recommendation for 

use. 
Foreword 

Sediment contamination is a widespread environmental problem that can potentially pose a threat to a variety of aquatic ecosystems. Sediment functions as a 

reservoir for common contaminants such as pesticides, herbicides, polychlorinated biphenyls (PCBs), polycyclic aromatic hydrocarbons, and metals such as 

lead, mercury, and arsenic. In-place contaminated sediment can result in 

depauparate benthic communities, while disposal of contaminated dredge material 

can potentially exert adverse effects on both pelagic and benthic systems. 

Historically, assessment of sediment quality has been limited to chemical characterizations. The United States Environmental Protection Agency (USEPA) is 

developing methodologies to calculate chemical-specific sediment quality criteria 

for use in the Agency’s regulatory programs. However, quantifying contaminant 

concentrations alone cannot always provide enough information to adequately 

evaluate potential adverse effects that arise from interactions among chemicals, or 

that result from time-dependent availability of sediment-associated contaminants 

to aquatic organisms. Because relationships between concentrations of contaminants in sediment and bioavailability are not fully understood, determination of 

contaminated sediment effects on aquatic organisms may require the use of 

controlled toxicity and bioaccumulation tests. 

As part of USEPA’s Contaminated Sediment Management Strategy, all Agency 

programs have agreed to use the same methods to determine whether sediments 

have the potential to affect aquatic ecosystems. More than ten federal statutes 

provide authority to many USEPA program offices to address the problem of 

contaminated sediment. The sediment test methods in this manual will be used by 

USEPA to make decisions under a range of statutory authorities concerning such 

issues as: dredged material disposal, registration of pesticides and toxic substances, Superfund site assessment, and assessment and cleanup of hazardous 

waste treatment, storage, and disposal facilities. The use of uniform sediment 

testing procedures by USEPA programs is expected to increase data accuracy 

and precision, facilitate test replication, increase the comparative value of test 

results, and, ultimately, increase the efficiency of regulatory processes requiring 

sediment tests. 

For additional guidance on the technical considerations in the manual, please 

contact Teresa Norberg-King, USEPA, Duluth, MN. 

iii 
Abstract 

Procedures are described for testing freshwater organisms in the laboratory to 

evaluate the toxicity or bioaccumulation of contaminants associated with whole 

sediments. Sediments may be collected from the field or spiked with compounds in 

the laboratory. Toxicity methods are outlined for two organisms, the amphipod 

Hyalella azteca and the midge Chironomus tentans. The toxicity tests are conducted for 10d in 300-mL chambers containing 100 mL of sediment and 175 mL of 

overlying water. Overlying water is renewed daily and test organisms are fed 

during the toxicity tests. The endpoint in the toxicity test with H. azteca is survival 

and the endpoints in the toxicity test with C. tentans are survival and growth. 

Procedures are primarily described for testing freshwater sediments; however, 

estuarine sediments (up to 15% salinity) can also be tested with H. azteca. 

Guidance for conducting 28-d bioaccumulation tests with the oligochaete 

Lumbriculus variegatus is provided in this manual. Overlying water is renewed 

daily and test organisms are not fed during bioaccumulation tests. Methods are 

also described for determining bioaccumulation kinetics of different classes of 

compounds during 28-d exposures with L. variegatus. 

iv 
Contents 

Foreword ........................................................................................................ ............................. iii 

Abstract ......................................................................... .............................. ............................. iv 

Figures ........................................................................................................... .. ........................... viii 

Tables .............................................................................................................................................. x 

Acknowledgments .......... ...................................................... ....................................................... xii 

Text Introduction .............................................................................................................................. 1 

1.1 Significance of Use ........................................................................................................ 1 

1.2 Program Applicability.. .................................................................................................... 2 

1.3 Scope and Application ................................................................................................... 3 

1.4 Performance-based Criteria ........................................................................................... 8 

Text Summary of Method.. ............................................................................................................. 10 

2.1 Method Description and Experimental Design ............................................................. 10 

2.2 Types of Tests.. ............................................................................................................ 11 

2.3 Test Endpoints ............................................................................................................. 11 

Text Definitions .............................................................................................................................. 12 

Text Interferences .......................................................................................................................... 14 

4.1 General Introduction ..................................................................................................... 14 

4.2 Non-Contaminant Factors ............................................................................................ 15 

4.3 Changes in Bioavailability ............................................................................................ 15 

4.4 Presence of Indigenous Organisms ............................................................................. 16 

Text Health, Safety, and Waste Management ............................................................................... 17 

5.1 General Precautions.. ................................................................................................... 17 

5.2 Safety Equipment ......................................................................................................... 17 

5.3 General Laboratory and Field Operations .................................................................... 17 

5.4 Disease Prevention ...................................................................................................... 18 

5.5 Safety Manuals.. ........................................................................................................... 18 

5.6 Pollution Prevention, Waste Management, and Sample Disposal ............................... 18 

Text Facilities, Equipment, and Supplies ....................................................................................... 19 

6.1 General.. ....................................................................................................................... 19 

6.2 Facilities ....................................................................................................................... 19 

6.3 Equipment and Supplies .............................................................................................. 19 

Text Water, Formulated Sediment, Reagents, and Standards ..................................................... .22 

7.1 Water ............................................................................................................................ 22 

7.2 Formulated Sediment ................................................................................................... 23 

7.3 Reagents ...................................................................................................................... 25 

7.4 Standards ..................................................................................................................... 25 

v 
Contents (continued) 

8 Sample Collection, Storage, Manipulation, and Characterization ......................................... .26 

8.1 Collection.. .................................................................................................................... 26 

8.2 Storage ......................................................................................................................... 26 

8.3 Manipulation ................................................................................................................. 27 

8.4 Characterization ........................................................................................................... 28 

9 Quality Assurance and Quality Control .................................................................................. 30 

9.1 

9.2 

9.3 

9.4 

9.5 

9.6 

9.7 

9.8 

9.9 

9.10 

9.11 

9.12 

9.13 

9.14 

9.15 

9.16 

9.17 

Introduction.. ................................................................................................................. 30 

Performance-based Criteria ......................................................................................... 30 

Facilities, Equipment, and Test Chambers .................................................................. .30 

Test Organisms ............................................................................................................ 31 

Water.. .......................................................................................................................... 31 

Sample Collection and Storage .................................................................................... 31 

Test Conditions ............................................................................................................ 31 

Quality of Test Organisms.. .......................................................................................... 31 

Quality of Food.. ........................................................................................................... 31 

Test Acceptability ......................................................................................................... 31 

Analytical Methods ....................................................................................................... 31 

Calibration and Standardization ................................................................................... 31 

Replication and Test Sensitivity ................................................................................... 32 

Demonstrating Acceptable Performance ...................................................................... 32 

Documenting Ongoing Laboratory Performance .......................................................... 32 

Reference Toxicants .................................................................................................... 32 

Record Keeping.. .......................................................................................................... 33 

10 Collection, Culturing, and Maintaining Test Organisms ........................................................35 

10.1 Life Histories ................................................................................................................ 35 

10.2 General Culturing Procedures.. .................................................................................... 37 

10.3 Culturing Procedures for Hyalella azteca .................................................................... .38 

10.4 Culturing Procedures for Chironomus tentans ............................................................ .39 

10.5 Culturing Procedures for Lumbriculus variegarus ........................................................ 43 

11 Test Method 100.1: Hyalella azteca 10-d Survival Test for Sediments ................................44 

11.1 Introduction.. ................................................................................................................. 44 

11.2 Recommended Test Method for Conducting a 10-d Sediment Toxicity Test 

with Hyalella azteca ..................................................................................................... 44 

11.3 General Procedures ..................................................................................................... 44 

11.4 interpretation of Results ............................................................................................... 48 

12 Test Method 100.2: Chironomus tentans 10-d Survival and Growth Test for Sediments.. ... 51 

12.1 Introduction.. ................................................................................................................. 51 

12.2 Recommended Test Method for Conducting a 10-d Sediment Toxicity Test 

with Chimnomus tentans.. ............................................................................................ 51 

12.3 General Procedures ..................................................................................................... 51 

12.4 Interpretation of Results ............................................................................................... 55 

vi 
Contents (continued) 

13 Test Method 100.3: Lumbriculus variegatus Bioaccumulation Test for Sediments .............. 57 

13.1 Introduction.. ................................................................................................................. 57 

13.2 Procedure for Conducting Sediment Bioaccumulation Tests with 

Lumbriculus variegatus ................................................................................................ 57 

13.3 General Procedures ..................................................................................................... 58 

13.4 Interpretation of Results ............................................................................................... 63 

14 Data Recording, Data Analysis and Calculations, and Reporting .......................................... 64 

14.1 Data Recording ............................................................................................................ 64 

14.2 Data Analysis ............................................................................................................... 64 

14.3 Data Interpretation.. ...................................................................................................... 80 

14.4 Reporting.. .................................................................................................................... 81 

15 Precision and Accuracy ......................................................................................................... 82 

15.1 Determining Precision and Accuracy .......................................................................... .82 

15.2 Accuracy.. ..................................................................................................................... 82 

15.3 Replication and Test Sensitivity .................................................................................. .83 

15.4 Demonstrating Acceptable Laboratory Performance.. .................................................. 83 

15.5 Precision of Sediment Toxicity Test Methods ............................................................. .84 

References ................................................................................................................................... 90 

Appendices 

A. Summary of USEPA Workshop on Development of Standard 

Sediment Test Methods ............................................................................................. 101 

B. Exposure Systems ..................................................................................................... 109 

C. Food Preparation.. ...................................................................................................... 120 

D. Sample Data Sheets .................................................................................................. 125 

vii 
Figures 

Figure 10.1 

Figure 10.2 

Figure 11.1 

Figure 11.2 

Figure 12.1 

Figure 14.1 

Figure 14.2 

Figure 14.3 

Figure 14.4 

Figure 14.5 

Figure 14.6 

Figure 14.7 

Figure 14.8 

Figure 15.1 

Figure B.1 

Figure B.2 

Figure B.3 

Figure B.4 

Figure D.1 

Figure D.2 

Figure D.3 

Length and relative age of Hyalella azteca collected by sieving in comparison 

with length of known-age organisms. .................................................................... 40 

Aspirator chamber (A) and reproduction and oviposit chamber (B) 

for adult midges. .................................................................................................... 42 

Lifestage sensitivity of Hyalella azteca in 96-h water-only exposures. ................. 49 

Average recovery of different age Hyalella azteca from sediment 

by 7 individuals. ..................................................................................................... 50 

Lifestage sensitivity of chironomids. ...................................................................... 56 

Treatment response for a Type I and Type II error.. ............................................. .66 

Power of the test vs. percent reduction in treatment response 

relative to the control mean at various CVs 

(8 replicates, alpha = 0.05 (one-tailed)). ................................................................ 67 

Power of the test vs. percent reduction in treatment response 

relative to the control mean at various CVs 

(5 replicates, alpha = 0.05 (one-tailed)). ................................................................ 68 

Power of the test vs. percent reduction in treatment response 

relative to the control mean at various CVs 

(8 replicates, alpha = 0.10 (one-tailed)). ................................................................ 69 

Effect of CV and number of replicates on the power to detect a 

20% decrease in treatment response relative to the control mean 

(alpha = 0.05 (one-tailed)). .................................................................................... 70 

Effect of alpha and beta on the number of replicates at various CVs 

(assuming combined alpha + beta = 0.25). ........................................................... 71 

Decision tree for analysis of survival and growth data 

subjected to hypothesis testing. ............................................................................. 72 

Decision tree for analysis of point estimate data. .................................................75 

Control (cusum) charts: (A) hypothesis testing and 

(B) point estimates (LC, EC, or 1C). ....................................................................... 83 

Portable table top STIR system described in Benoit et al. (1993). ..................... 110 

Portable table top STIR system with several additional options 

as described in Benoit et al. (1993). .................................................................... 111 

Tanks for the STIR system in Benoit et al. (1993). .............................................. 112 

Water splitting chamber described in Zumwalt et al. (1994). ............................... 117 

Data sheet for the evaluation of a Chironomus tentans culture. .......................... 126 

Data sheet for performing reference toxicant tests. ............................................. 127 

Data sheet for temperature and overlying water chemistry measurements. ...... 128 

viii 
Figures (continued) 

Figure D.4 Data sheet for daily checklist for sediment tests. ................................................ 129 

Figure D.5 Data sheet for water quality parameters. ............................................................. 130 

Figure D.6 Chemistry data sheet. .......................................................................................... 131 

Figure D.7 Daily comment data sheet. .................................................................................. 132 

Figure D.8 Weight data sheet. ............................................................................................... 133 

ix 
Tables 

Table 1.1 

Table 1.2 

Table 1.3 

Table 1.4 

Table 4.1 

Table 6.1 

Table 7.1 

Table 7.2 

Table 7.3 

Table 9.1 

Table 9.2 

Table 10.1 

Table 10.2 

Table 11.1 

Table 11.2 

Table 11.3 

Table 12.1 

Table 12.2 

Table 12.3 

Table 13.1 

Table 13.2 

Table 13.3 

Table 13.4 

Sediment Quality Assessment Procedures.. ...........................................................3 

Statutory Needs for Sediment Quality Assessment ................................................. 4 

Rating of Selection Criteria for Freshwater Sediment 

Toxicity Testing Organisms.. .................................................................................... 5 

Water-only, 10-d LC50 (μg/L) Values for Hyalella azteca, 

Chironomus tentans, and Lumbriculus variegatus .................................................. .6 

Advantages and Disadvantages for Use of Sediment Tests ................................. 14 

Equipment and Supplies for Culturing and Testing Specific Test Organisms ...... 21 

Characteristics of Three Sources of Clays and Silts 

Used in Formulated Sediments ............................................................................. 24 

Carbon, Nitrogen, Phosphorus Levels for Various Sources of Organic Carbon.. . 24 

Sources of Components Used in Formulated Sediments ..................................... .25 

Recommended Test Conditions for Conducting Reference-Toxicity Tests 

with One Organism/Chamber ............................................................................... .33 

Recommended Test Conditions for Conducting Reference-Toxicity Tests 

with More Than One Organism/Chamber .............................................................34 

Sources of Test Organisms ................................................................................... 37 

Chironomus tentans Instar and Head Capsule Widths ......................................... .40 

Test Conditions for Conducting a 10-d Sediment Toxicity Test 

with Hyalella azteca ............................................................................................... 45 

General Activity Schedule for Conducting a Sediment Toxicity Test 

with Hyalella azteca ............................................................................................... 45 

Test Acceptability Requirements for a 10-d Sediment Toxicity Test 

with Hyalella azteca ............................................................................................... 46 

Recommended Test Conditions for Conducting a 

10-d Sediment Toxicity Test with Chironomus tentans.. ....................................... .52 

General Activity Schedule for Conducting a Sediment Toxicity Test 

with Chironomus tentans ....................................................................................... 53 

Test Acceptability Requirements for a 10-d Sediment Toxicity Test 

with Chironomus tentans ....................................................................................... 54 

Recommended Test Conditions for Conducting a 

28-d Sediment Bioaccumulation Test with Lumbriculus variegatus.. .................... 58 

Recommended Test Conditions for Conducting a Preliminary 4-d Sediment 

Toxicity Screening Test with Lumbriculus variegatus ........................................... .59 

General Activity Schedule for Conducting a 28-d Sediment Bioaccumulation 

Test with Lumbriculus variegatus.. ......................................................................... 60 

Test Acceptability Requirements for a 28-d Sediment Bioaccumulation Test 

with Lumbriculus variegatus.. ................................................................................. 61 

X 
Tables (continued) 

Table 13.5 

Table 13.6 

Table 14.1 

Table 14.2 

Table 15.1 

Table 15.2 

Table 15.3 

Table 15.4 

Table 15.5 

Table 15.6 

Table 15.7 

Table 15.8 

Table A.1 

Table A.2 

Table A.3 

Table A.4 

Table B.1 

Table B.2 

Table B.3 

Table C.1 

Table C.2 

Grams of Lumbriculus variegatus Tissue (Wet Weight) Required for 

Various Analytes at Selected Lower Limits of Detection ....................................... 62 

Detection Limits (ng) of Individual PAHs by HPLC-FD .......................................... 62 

Suggested a Levels to Use for Tests of Assumptions.. ......................................... 72 

Estimated Time to Obtain 95 Percent of Steady-State Tissue Residue ............... 79 

lntralaboratory Precision for Survival of Hyalella azteca and 

Chironomus tentans in 10-d Whole-Sediment Toxicity Tests, June 1993 ............ 84 

Participants in Round Robin Studies .................................................................... .85 

Interlaboratory Precision for Hyalella azteca 96-h LC50s from Water-only 

Static Acute Toxicity Tests Using a Reference Toxicant (KCI) (October 1992). ... 85 

Interlaboratory Precision for Survival of Hyalella azteca in 10-d 

Whole-Sediment Toxicity Tests Using Four Sediments (March 1993) ................. 86 

Interlaboratory Precision for Chironomus tentans 96-h LC50s from 

Water-only Static Acute Toxicity Tests Using a Reference Toxicant 

(KCI) (December 1992) .......................................................................................... 87 

Interlaboratory Precision for Chironomus tentans 96-h LC50s from 

Water-only Static Acute Toxicity Tests Using a Reference Toxicant 

(KCI) (May 1993) ................................................................................................... 87 

Interlaboratory Precision for Survival of Chironomus tentans in 10-d 

Whole-Sediment Toxicity Tests Using Three Sediments (May 1993). .................. 88 

Interlaboratory Precision for Growth of Chironomus tentans in 10-d 

Whole-Sediment Toxicity Tests Using Three Sediments (May 1993). .................. 89 

List of Laboratories Responding to the Survey.. .................................................. 101 

Summary of Testing Procedures Used to Evaluate the Toxicity of 

Whole Sediments with Hyalella azteca.. .............................................................. 102 

Summary of Testing Procedures Used to Evaluate the Toxicity of 

Whole Sediments with Chironomus tentans ........................................................ 104 

Summary of Testing Procedures Used to Conduct Whole-Sediment 

Bioaccumulation Tests with Lumbriculus variegatus ........................................... 106 

Sediment Copper Concentrations and Organism Survival and Growth 

at the End of a 10-d Test.. ................................................................................... 115 

Sediment Dieldrin Concentrations and Organism Survival and Growth 

at the End of a 10-d Test.. ................................................................................... 116 

Materials Needed for Constructing a Zumwalt et al. (1994) Delivery System ... 118 

Nutrient Stock Solutions for Maintaining Algal Stock Cultures ............................ 121 

Final Concentration of Macronutrients and Micronutrients in the 

Algal Culture Medium .......................................................................................... 121 

xi 
Acknowledgments 

This document is a general purpose testing manual for freshwater sediments. The 

approaches have also been described in ASTM (1994a) ASTM (1994b), Ankley et 

al. (1993), Phipps et al. (1993), Brooke et al. (1993), Call et al. (1993a), Call et al. 

(1993b), Lee et al. (1994) and USEPA (1994a). 

This manual reflects the consensus of the Freshwater Sediment Toxicity Assessment Committee and the U.S. Environmental Protection Agency (USEPA) Program Offices. Members of the Freshwater Sediment Toxicity Assessment Committee are G.T. Ankley, USEPA, Duluth, MN; D.A. Benoit, USEPA, Duluth, MN; G.A. 

Burton, Wright State University, Dayton, OH; F.J. Dwyer, National Biological 

Survey (NBS; formerly U.S. Fish and Wildlife Service), Columbia, MO; I.E. Greer, 

NBS, Columbia, MO; R.A. Hoke, SAIC, Hackensack, NJ; C.G. Ingersoll, NBS, 

Columbia, MO; P. Kosian, USEPA, Duluth, MN; P.F. Landrum, NOAA, Ann Arbor, 

MI; J.M. Lazorchak, USEPA, Cincinnati, OH; T.J. Norberg King, USEPA, Duluth, 

MN; and P.V. Winger, NBS, Athens, GA. 

The principal authors of this document are C.G. Ingersoll, G.T. Ankley, G.A. 

Burton, F.J. Dwyer, R.A. Hoke, T.J. Norberg-King, and P.V. Winger. Contributors 

to specific sections of this manual are listed below. 

1. Sections 1-9; General Guidelines 

G.T. Ankley, USEPA, Duluth, MN 

G.A. Burton, Wright State University, Dayton, OH 

F.J. Dwyer, NBS, Columbia, MO 

R.A. Hoke, SAIC, Hackensack, NJ 

C.G. Ingersoll, NBS, Columbia, MO 

T.J. Norberg-King, USEPA, Duluth, MN 

C.E. Schlekat, SAIC, Narragansett, RI 

K.J. Scott, SAIC, Narragansett, RI 

2. Sections 10-13; Culture and Test Methods 

G.T. Ankley, USEPA, Duluth, MN 

D.A. Benoit, USEPA, Duluth, MN 

E.L. Brunson, NBS, Columbia, MO 

F.J. Dwyer, NBS, Columbia, MO 

I.E. Greer, NBS, Columbia, MO 

R.A. Hoke, SAIC, Hackensack, NJ 

C.G. Ingersoll, NBS, Columbia, MO 

P.F. Landrum, NOAA, Ann Arbor, Ml 

H. Lee, USEPA, Newport, OR 

T.J. Norberg-King, USEPA, Duluth, MN 

P.V. Winger, NBS, Athens, GA 

xii 
Acknowledgments (continued) 

3. Section 14; Statistical Analysis 

J. Heltshe, SAIC, Narragansett, RI 

R.A. Hoke, SAIC, Hackensack, NJ 

H. Lee, USEPA, Newport, OR 

T.J. Norberg-King, USEPA, Duluth, MN 

C. Schlekat, SAIC, Narragansett, RI 

4. Section 15; Precision and Accuracy 

G.T. Ankley, USEPA, Duluth, MN 

G.A. Burton, Wright State University, Dayton, OH 

C.G. Ingersoll, NBS, Columbia, MO 

T.J. Norberg-King, USEPA, Duluth, MN 

Review comments from the following individuals are gratefully acknowledged: C. 

Philbrick Barr and P. Nolan, Region 1, Lexington, MA; D. Reed, Permits Division, 

OWEC, Washington, D.C.; P. Crocker, Technical Section and S. McKinney, 

Marine and Estuarine Section, Region 6, Dallas, TX: F. Schmidt, Monitoring 

Branch, OWOW, Washington, D.C.; T. Armitage, Standards and Applied Science 

Division, OST, Washington, D.C., T. Bailey, Environmental Effects Branch and J. 

Smrchek, OPPT, Washington, D.C.; D. Klemm, EMSL and L. Gast, TAI, Newtown, 

OH; G. Hanson, OSW, Washington, DC., S. Ferraro and R. Swartz, ERL-N, 

Newport, OR; J. Arthur, R. Spehar, and C. Stephan, ERL-D, Duluth, MN; J. 

Thompson, T. Dawson, J. Jenson, S. Collyard, and J. Juenemann, ILS, Duluth, 

MN; J. Scott and C. Scheklat, SAIC, Narragansett, RI. 

Participation by the following laboratories in the round-robin testing is greatly 

appreciated: ABC Laboratories, Columbia, MO; Environment Canada, Burlington, 

Ontario; EVS Consultants, Vancouver, BC; Michigan State University, East Lansing, MI; National Fisheries Contaminant Research Center, Athens, GA; Midwest 

Science Center, Columbia, MO; Center University of Mississippi, University, MS; 

University of Wisconsin-Superior, Superior, WS; USEPA, Cincinnati, OH; USEPA, 

Duluth, MN; Washington Department of Ecology, Manchester, WA; Wright State 

University, Dayton, OH. Culturing support was supplied for USEPA Duluth by S. 

Collyard, J. Juenenmann, J. Jenson, and J. Denny. 

USEPA’s Office of Science and Technology provided support for the development 

of this manual. 

xiii 
Section 1 

Introduction 

1.1 Significance of Use 

1.1.1 Sediment provides habitat for many aquatic organisms and is a major repository for many of the more 

persistent chemicals that are introduced into surface 

waters. In the aquatic environment, most anthropogenic 

chemicals and waste materials including toxic organic 

and inorganic chemicals eventually accumulate in sediment. Mounting evidence exists of environmental degradation in areas where USEPA Water Quality Criteria 

(WQC; Stephan et al., 1985) are not exceeded, yet 

organisms in or near sediments are adversely affected 

(Chapman, 1989). The WQC were developed to protect 

organisms in the water column and were not intended to 

protect organisms in sediment. Concentrations of contaminants in sediment may be several orders of magnitude higher than in the overlying water; however, bulk 

sediment concentrations have not been strongly correlated to bioavailability (Burton, 1991). Partitioning or 

sorption of a compound between water and sediment 

may depend on many factors including: aqueous solubility, pH, redox, affinity for sediment organic carbon and 

dissolved organic carbon, grain size of the sediment, 

sediment mineral constituents (oxides of iron, manganese, and aluminum), and the quantity of acid volatile 

sulfides in sediment (Di Toro et al., 1990, 1991). Although certain chemicals are highly sorbed to sediment, 

these compounds may still be available to the biota. 

Contaminated sediments may be directly toxic to aquatic 

life or can be a source of contaminants for bioaccumulation in the food chain. 

1.1.2 Assessments of sediment quality have commonly 

included sediment chemical analyses and surveys of 

benthic community structure. Determination of sediment 

contaminant concentrations on a dry weight basis alone 

offers little insight into predicting adverse biological effects because bioavailability may be limited by the intricate partitioning factors mentioned above. Likewise, 

benthic community surveys may be inadequate because 

they sometimes fail to discriminate between effects of 

contaminants and those that result from unrelated 

non-contaminant factors, including water-quality fluctuations, physical parameters, and biotic interactions. In 

order to obtain a direct measure of sediment toxicity or 

bioaccumulation, laboratory tests have been developed 

in which surrogate organisms are exposed to sediments 

under controlled conditions. Sediment toxicity tests have 

evolved into effective tools providing direct, quantifiable 

evidence of biological consequences of sediment contamination that can only be inferred from chemical or 

benthic community analyses. The USEPA is developing 

a national inventory of contaminated sediment sites. 

This inventory will be used 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 testing methods will provide high quality data needed for the national 

inventory and for regulatory programs to prevent, remediate, and manage contaminated sediment (Southerland 

et al., 1991). 

1.1.3 The objective of a sediment test is to determine 

whether contaminants in sediment are harmful to or are 

bioaccumulated by benthic organisms. The tests can be 

used to measure interactive toxic effects of complex 

contaminant mixtures in sediment. Furthermore, knowledge of specific pathways of interactions among sediments and test organisms is not necessary in order to 

conduct the tests (Kemp and Swartz, 1988). Sediment 

tests can be used to (1) determine the relationship 

between toxic effects and bioavailability, (2) investigate 

interactions among contaminants, (3) compare the sensitivities of different organisms, (4) determine spatial 

and temporal distribution of contamination, (5) evaluate 

hazards of dredged material, (6) for measuring toxicity 

as part of product licensing or safety testing or chemical 

approval, (7) rank areas for clean up, and (8) set cleanup 

goals and estimate the effectiveness of remediation or 

management practices. 

1.1.4 Results of toxicity tests on sediments spiked at 

different concentrations of contaminants can be used to 

establish cause and effect relationships between chemicals and biological responses. Results of toxicity tests 

with test materials spiked into sediments at different 

concentrations may be reported in terms of an LC50 

(median lethal concentration), an EC50 (median effect 

concentration), an IC50 (inhibition concentration), or as 

a NOEC (no observed effect concentration) or LOEC 

(lowest observed effect concentration). in some cases, 

results of bioaccumulation tests may also be reported in 

terms of a Biota-sediment Accumulation Factor (BSAF) 

(Ankley et al., 1992a; Ankley et al., 1992b). 

1.1.5 Evaluating effect concentrations for chemicals in 

sediment requires knowledge of factors controlling their 

bioavailability. Similar concentrations of a chemical in 

1 
units of mass of chemical per mass of sediment dry 

weight often exhibit a range in toxicity in different sediments (Di Toro et al., 1990: Di Toro et al., 1991). Effect 

concentrations of chemicals in sediment have been 

correlated to interstitial water concentrations, and effect 

concentrations in interstitial water are often similar to 

effect concentrations in water-only exposures. The bioavailability of nonionic organic compounds in sediment 

is often inversely correlated with the organic carbon 

concentration. Whatever the route of exposure, these 

correlations of effect concentrations to interstitial water 

concentrations indicate that predicted or measured concentrations in interstitial water can be used to quantify 

the exposure concentration to an organism. Therefore, 

information on partitioning of chemicals between solid 

and liquid phases of sediment is useful for establishing 

effect concentrations (Di Toro et al., 1991). 

1.1.6 Field surveys can be designed to provide either a 

qualitative reconnaissance of the distribution of sediment contamination or a quantitative statistical comparison of contamination among sites. Surveys of sediment 

toxicity or bioaccumulation are usually part of more 

comprehensive analyses of biological, chemical, geological, and hydrographic data. Statistical correlations 

may be improved and sampling costs may be reduced if 

subsamples are taken simultaneously for sediment tests, 

chemical analyses, and benthic community structure. 

1.1.7 Table 1.1 lists several approaches the USEPA 

has considered for the assessment of sediment quality 

(USEPA, 1992c). These approaches include (1) equilibrium partitioning, (2) tissue residues, (3) interstitial water 

toxicity, (4) whole-sediment toxicity and sediment-spiking 

tests, (5) benthic community structure, and (6) Sediment 

Quality Triad and Effects Range Median (see Chapman, 

1989 and USEPA, 1989a; USEPA, 1990a; USEPA, 

1990b; USEPA, 1992b for a critique of these methods). 

The sediment assessment approaches listed in Table 

1.1 can be classified as numeric (e.g., equilibrium partitioning), descriptive (e.g., whole-sediment toxicity tests), 

or a combination of numeric and descriptive approaches 

(e.g., Effects Range Median: USEPA, 1992c). Numeric 

methods can be used to derive chemical-specific sediment quality criteria (SQC). Descriptive methods such 

as toxicity tests with fieldcollected sediment cannot be 

used alone to develop numerical SQC for individual 

chemicals. Although each approach can be used to 

make site-specific decisions, no one single approach 

can adequately address sediment quality. Overall, an 

integration of several methods using the weight of evidence is the most desirable approach for assessing the 

effects of contaminants associated with sediment (Long 

and Morgan, 1991). Hazard evaluations integrating data 

from laboratory exposures, chemical analyses, and 

benthic community assessments provide strong complementary evidence of the degree of pollution-induced 

degradation in aquatic communities (Chapman et al., 

1992; Burton, 1991). 

1.2 Program Applicability 

1.2.1 The USEPA has authority under a variety of 

statutes to manage contaminated sediment. Until recently, the USEPA has not addressed sediment quality 

except in relation to disposal of material removed during 

navigational dredging (Table 1.2). Southerland et al. 

(1992) outlined four goals of an USEPA management 

strategy for contaminated sediments: (1) in-place sediment should be protected from contamination to ensure 

beneficial uses of surface waters, (2) protection of in-place 

sediment should be achieved through pollution prevention and source control, (3) in-place remediation should 

be limited to locations where natural recovery will not 

occur in an acceptable period of time, and (4) consistent 

methods should be used to trigger regulatory decisions. 

1.2.2 The Clean Water Act (CWA) is the single most 

important law dealing with environmental quality of surface waters in the United States. The goal of the CWA is 

to restore and maintain physical, chemical, and biological integrity of the nation’s waters (Southerland et al., 

1992). Federal and state monitoring programs traditionally have focused on evaluating water column problems 

caused by point-source dischargers. The USEPA is 

developing a national inventory of contaminated sediment sites. This inventory will be used 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 

testing methods will provide high quality data needed for 

the national inventory and for regulatory program to 

prevent, remediate, and manage contaminated sediment (Southerland et al., 1992). 

1.2.3 The Office of Water (OW), the Office of Pesticide 

Programs (OPP), the Office of Pollution Prevention and 

Toxics (OPPT), the Office of Solid Waste (OSW), and 

the Office of Emergency and Remedial Response 

(OERR) are all committed to the principle of consistent 

tiered testing outlined in the Agency-wide Contaminated 

Sediment Strategy (Southerland et al., 1992). 

Agency-wide consistent testing is desirable because all 

USEPA programs will use similar methods to evaluate 

whether a sediment poses an ecological or human 

health risk, and comparable data would be produced. It 

will also provide the basis for uniform cross-program 

decision-making within the USEPA. Each program will, 

however retain the flexibility of deciding whether identified risks would trigger regulatory actions. 

1.2.4 Tiered testing should include a hierarchy of tests 

with the tests in each successive tier becoming progressively more rigorous, complex, and costly (Southerland 

et al, 1992). Guidance needs to be developed to explain 

how information within each tier would trigger regulatory 

action. The guidance could be program specific, describing decisions based on a weight of evidence approach, a pass-fail approach, or comparison to a reference site depending on statutory and regulatory requirements. There are now two approaches for sediment 

2 
Table 1.1 Sediment Quality Assessment Procedures’ 

Type 

Method Numeric Descriptive Combination 

Equilibrium Partitioning * 

Approach 

A sediment quality value for a given contaminant IS determined by 

calculating the sediment concentration of the contaminant that 

corresponds toan interstitial water concentration equivalent to the 

USEPA water-quality criterion for the contaminant. 

Tissue Residues *** Safe sediment concentrations of specific chemicals are established 

by determining the sediment chemical concentration that results 

acceptable tissue residues. 

Interstitial Water Toxicity * ** Toxicity of interstitial water IS quantified and identification 

evaluation procedures are applied to identify and quantity chemical 

components responsible for sediment toxicity. 

Benthic Community 

Structure 

Environmental degradation IS measured by evaluating alterations 

in benthic community structure. 

Whole-sediment Toxicity * * * 

and Sediment Spiking 

Test organisms are exposed to sedimentshat may contain known 

or unknown quantities of potentially toxic chemicals. At the end of a 

specified time period, the response of the test organisms IS examined 

in relation to a specified endpoint. Dose-response relationships 

can be established by exposing test organisms to sediments that 

have been spiked with known amounts of chemicals or mixtures of 

chemicals. 

Sediment Quality Triad * * * Sediment chemical contamination, sediment toxicity, and benthic 

community structure are measured on the same sediment sample 

Correspondence between sediment chemistry, toxicity, and field 

effects is used to determine sediment concentrations that 

discriminate conditions of minimal, uncertain, and major biological 

effects. 

Apparent Effects Threshold * * * 

(AET) 

The sediment concentration of a contaminant above which 

statistically significant biological effects (e.g., sediment toxicity) are 

always expected. AET values are empirically derived from paired 

field data for sediment chemistry and a range of biological effects 

indicators. 

1 Modified from USEPA (1992c) 

testing used by USEPA: (1) the Office of Water-U.S. 

Army Corps of Engineers dredged material testing framework and (2) the OPPT ecological risk assessment 

tiered testing framework. Tier 1 of the dredged material 

testing framework consists of a review of existing chemical and biological data or an inventory of nearby sources. 

In Tier 2, chemical data are compared to water and 

sediment quality criteria. Tier 3 evaluations consist of 

acute toxicity and bioaccumulation testing. and a comparison of the results to a reference area. The OPP 

testing framework consists of acute toxicity testing in 

Tier 1, followed by chronic (early life stage) toxicity 

testing in Tier 2 and further chronic toxicity testing (full 

life cycle) in Tier 3. Tier 4 consists of field or mesocosm 

testing. A tiered testing framework has not yet been 

chosen for Agency-wide use, but some of the components have been identified to be standardized. These 

components are toxicity tests. bioaccumulation tests, 

chemical criteria, and other measurements that may 

have ecological significance including benthic community structure evaluation, colonization rate, and in situ 

sediment testing within a mesocosm (USEPA, 1992a). 

1.3 Scope and Application 

1.3.1 Procedures are described for testing freshwater 

organisms in the laboratory to evaluate the toxicity or 

bioaccumulation of contaminants associated with whole 

sediments. Sediments may be collected from the field or 

spiked with compounds in the laboratory. Toxicity methods are outlined for two organisms, the amphipod Hyalella 

azteca and the midge Chironomus tentans. The toxicity 

tests are conducted for 10 d in 300-mL chambers containing 100 mL of sediment and 175 mL of overlying 

water. Overlying water is renewed daily and organisms 

are fed during the toxicity tests. The endpoint in the 

toxicity test with H. azteca is survival and the endpoints 

in the toxicity test with C. tentans are survival and 

growth. Procedures are primarily described for testing 

freshwater sediments; however, estuarine sediments 

(up to 15% salinity) can also be tested with H. azteca. 

Guidance for conducting 28-d bioaccumulation tests 

with the oligochaete Lumbriculus variegatus is provided 

in this manual. Overlying water is renewed daily and 

organisms are not fed during bioaccumulation tests. 

Methods are also described for determining bioaccumu3 
Table 1.2 Statutory Needs for Sediment Quality Assessment* 

Law- Area of Need 

CERCLA * Assess need for remedral actlon wrth contaminated sedrments; assess degree of cleanup required, dtsposttlon of 

sedrments 

C’WA 

. 

. 

. 

NPDES permttmg. especraily under Best AvarIable Technology (BAT) In water-qualrty-lrmrted water 

Section 403(c) cntena for ocean discharges; mandatory addItional requrrements to protect marine envtronment 

Sectlon 301(g) waivers for publtcly owned treatment works (POT%) discharging to manne waters 

Sectron 404 permrts for dredge and fill activities (admmrstered by the Corps of Engineers) 

FIFRA . 

. 

Revrew uses of new and exrstrng chemicals 

Pestrcrde labeling and reglstratron 

MPRSA * Permrts for ocean dumprng 

NEPA * Preparation of envrronmental Impact statements for projects with surface water discharges 

TSCA * 

. 

Section 5: Premanufacture notrce reviews for new chemicals 

Sectrons 4. 5. and 6: Reviews for existing chemrcals 

RCAA * Assess surtabrlrty (and permit) on-land disposal or beneftcral use of contamrnated sedtments considered “hazardous” 

Modlfred from Drckson et al. 1964, and Southerland et al. 1992. 

CERCLA Comprehensive Envrronmental Resoonse. Compensatton and Lrability Act (Superfund). 

CWA 

FIFRA 

MPRSA 

NEPA 

TSCA 

RCRA 

Clean Water Act. 

Federal Insectlade, Fungerde, and Rodenttcrde Act. 

Manne Protectron. Resources and Sanctuary Act. 

National Envrronmental Pokey Act. 

TOXIC Substances Control Act. 

Resource Conservatron and Recovery Act. 

lation kinetics of different classes of compounds during 

28-d exposures with L. wariegatus. 

1.3.2 Additional research and methods development 

are now in progress to (1) develop standard chronic 

sediment toxicity tests (e.g., 28-d exposures with H. 

azteca), (2) refine formulated sediment, (3) refine sediment dilution procedures, (4) refine sediment Toxicity 

Jdentification Evaluation (TIE) procedures (Ankley and 

Thomas, 1992), (5) refine sediment spiking procedures, 

and (6) produce additional data on confirmation of responses in laboratory tests with natural populations of 

benthic organisms. This information will be described in 

future editions of this manual. 

1.3.3 This methods manual serves as a companion to 

the marine sediment testing method manual (USEPA, 

1994a). 

1.3.4 Procedures described in this manual are based on 

the following documents: ASTM (1994a), Ankley et al. 

(1993), Phipps et al. (1993), Call et al. (1994), Lee et al. 

(1994), and USEPA (1993a). This manual outlines specific test methods for evaluating the toxicity of sediments 

with I-/. azteca and C. tentans. Because additional research is still needed on the standardization of bioaccumulation procedures with sediments, this manual outlines only general guidance on procedures for evaluating the bioaccumulation of contaminants in sediment 

with L. variegatus. Many of the critical issues necessary 

for interpretation of test results are the subject of continuing research including the influence of feeding on 

bioavaitability, nutritional requirements of the test organisms, additional performance criteria for organism health, 

and confirmation of responses in laboratory tests with 

natural benthos populations. See Section 4 for additional details. 

1.3.5 General procedures described in this manual 

might be useful for conducting tests with other aquatic 

organisms; however, modifications may be necessary. 

Altering the procedures described in this manual may 

alter bioavailability and produce results that are not 

directly comparable with results of acceptable procedures. Comparison of results obtained using modified 

versions of these procedures might provide useful information concerning new concepts and procedures for 

conducting sediment tests with aquatic organisms (e.g., 

Diporeia spp., Tub&x tubifex, Hexagenia spp.). If tests 

are conducted with procedures different from those described in this manual, additional tests are required to 

determine comparability of results. 

1.36 Methods have been described for culturing and 

testing indigenous species that may be as sensitive or 

more sensitive than the species recommended in this 

manual. However, the USEPA currently allows the use 

of indigenous species only where state regulations require their use or prohibit importation of the recommended species. Where state regulations prohibit im4 
portation or use of the recommended test species, permission should be requested from the appropriate regulatory agency before their using indigenous species. 

1.3.7 Where states have developed culturing and testing methods for indigenous species other than those 

recommended in this manual, data comparing the sensitivity of the substitute species and one or more of the 

recommended species must be obtained with sediments 

or reference toxicants, to ensure that the species selected are at least as sensitive and appropriate as the 

recommended species. 

1.3.8 Selection of lest Organisms 

1.3.8.1 The choice of a test organism has a major 

influence on the relevance, success, and interpretation 

of a test. Test organism selection should be based on 

both environmental relevance and practical concerns 

(Dewitt et al., 1989, Swartz 1989). Ideally, a test organism should (1) have a toxicological database demonstrating relative sensitivity and discrimination to a range 

of contaminants of interest in sediment; (2) have a 

database for interlaboratory comparisons of procedures 

(e.g., round-robin studies); (3) be in direct contact with 

sediment; (4) be readily available through culture or 

from field collection; (5) be easily maintained in the 

laboratory; (6) be easily identified; (7) be ecologically or 

economically important; (8) have a broad geographical 

distribution, be indigenous (either present or historical) 

to the site being evaluated, or have a niche similar to 

organisms of concern (e.g., similar feeding guild or 

behavior to the indigenous organisms); (9) be tolerant of 

a broad range of sediment phsyico-chemical characteristics (e.g., grain size); and (10) be compatible with 

selected exposure methods and endpoints (Table 1.3, 

ASTM, 1993a). The method should also be (11) peer 

reviewed (e.g., journal articles, ASTM guides) and (12) 

confirmed with responses with natural populations of 

benthic organisms (Sections 1.3.8.8 and 1.3.9.6). 

Table 1.3 Rating of Selection Criter’ta for Freshwater Sediment Toxicity Testing Organisms’ 

Criterion Hyalella Dlpofeia Chrronomus Chlronomus Lumbriculus Tubifex Hexagema Mollusks Daphma spp. and 

azteca VP. tentans ripafius vanegatus tubifex SPP. Ceriodaphnia spp. 

Relative 

sensitivity 

toxicity + 

database 

Round-robin 

studres + 

conducted 

Contact wrth + 

sediment 

Laboratory + 

culture 

Taxonomic +Iidentification 

Ecological + 

Importance 

Geographical + 

distribution 

Sediment 

physico- + 

chemical 

tolerance 

Response 

confirmed + 

with benthos 

populations 

Peer reviewed + 

+ 

+l- 

+ 

+I- 

+ 

+ 

+ 

Endpomts2 S, G, M S, 0, A 

momtored 

+ 

+ 

+ 

+ 

+l- 

+ 

+ 

+I- 

+ 

+ 

S, G, E 

+ 

+ 

+I- 

+ 

+ 

+ 

+ 

+ 

S, G, E 

+ 

+ 

+ 

+ 

+ 

+ 

+ 

+ 

+ 

6 s, R 

+ 

+ 

+ 

+ 

+ 

+ 

+ 

+ 

s, R 

+ + 

+ 

+ + + 

+ + + 

+ + +I- 

+ NA 

+ + 

+ +iS. G 0 S. G. R 

’ A “+” or “-” rating Indicates a positive or negative attribute 

2 S = Survival, G = Growth, El = Broaccumulation, A = Avoidance, R - Reproduction, M = Maturation. E = Emergence. NA = not apo1lcab.e 

5 
1.3.8.2 Of these criteria (Table 1.3). a database demonstrating relative sensitivity to contaminants, contact with 

sediment, ease of culture in the laboratory, interlaboratory compansons, tolerance to varying sediment phsyicochemical characteristics, and confirmation with responses 

of natural benthos populations were the primary criteria 

used for selecting i-l. azteca. C. fentans, and L. variegatus 

for the current edition of this manual. Many organisms 

that mtght be appropriate for sediment testing do not 

now meet these selection criteria because historically 

little emphasis has been placed on developing standardized teshng procedures for benthic organisms. A similar 

database must be developed in order for other organisms to be included in future editions of this manual 

(e.g.. mayfkes (Hexagenia spp.), other midges (C. 

npariusl. other amphipods (Diporeia spp.), cladocerans 

(Daphnia magna, Ceriodaphnia d&a), or molt usks). 

1.3.8.3 An important consideration in the selection of 

specific species for test method development is the 

existence of information concerning relative sensitivity 

of the organisms both to single chemicals and complex 

mrxtures. A number of studies have evaluated the sensitivity of H. azteca, C. tenfans and L. variegatus. relative 

to one another, as well as other commonly tested freshwater species. For example, Ankley et al. (1991 b) found 

H. azteca to be as, or slightly more, sensitive than 

Ceriodaphma dubia to a variety of sediment elutriate 

and pore-water samples. In that study, L. variegafus 

were less sensitive to the samples than either the amphipod or the cladoceran. West et al. (1993) found the 

rank sensitivity of the three species to the lethal effects 

of copper-contaminated sediments could be ranked (from 

greatest to least): H. azteca > C. tentans :, L. variegatus. 

In short-term (48 to 96 h) exposures, L. variegatus 

generally was less sensitive than H. azteca, C. dubia, or 

Pimephales promelas to cadmium, nickel, zinc, copper, 

and lead (Schubauer-Berigan et al., 1993). Of the latter 

Table 1.4 Water-only, 10-d LCSO @g/L) Values ior Hyalella 

azteca. Chironomus tentans, and Lumbriculus 

vafiegatus * 

Cnemlcal H. azfeca C. tenfans L. vanegatus 

Copper 35 54 

Zinc 73 1.1252 

Cadmiurr 2.83 NT’ 

N,ckel 780 NT 

Lead ~16 NT 

3.p -DDT 0.07 1.23 

P.P -DDD 0.17 0.18 

p.p -DDE 1.39 3.0 

Dleldrln 7.6 1.1 

Chlorpynfos 0.086 0.07 

35 

2.984 

‘158 

12,?60 

794 

NT 

NT 

B3.3 

NT 

NT 

Chemica!s tested at ERL-Duluth In soft water-hardness 45 mg/L 

as CaCO, a! pH 7.8 to 8.2 (Phlpps et al., 1994). 

. 53% mortality at highest concentration tested. 

73% mortality at lowest concentration rested. 

’ NT = rot tested. 

three species, no one was consistently the most sensitive to all five metals. 

1.3.8.3.1 In a study of contaminated Great Lakes sediment, H. azteca, C. tentans, and C. riparius were among 

the most sensitive and discriminatory of 24 organisms 

tested (Burton and Ingersoll, 1994; Ingersoll et al., 1993). 

Kemble et al. (1993) found the rank sensitivity of four 

species to metal-contaminated sediments to be (from 

greatest to least): H. azteca > C. riparius > Oncorhynchus 

mykiss (rainbow trout) > Daphnia magna. The relative 

sensitivity of the three endpoints evaluated in the H. 

azteca test with Clark Fork River sediments was (from 

greatest to least): length > sexual maturation > survival. 

1.3.8.3.2 In 1 O-d water-only and whole-sediment tests, 

H. azteca and C. tenfans were more sensitive than D. 

magna to fluoranthene (Suede1 et al. 1993). 

1.3.8.3.3 Ten-day, water-only tests also have been 

conducted with a number of chemicals using the three 

species described in this manual (Phipps et al., 1994; 

Table 1.4). All tests were flow-through exposures using 

a soft natural water (Lake Superior) with measured 

chemical concentrations that, other than the absence of 

sediment, were conducted under conditions (e.g., temperature, photoperiod, feeding) similar to those being 

described for the standard 10-d sediment lest. In general, H. azteca was more sensitive to copper, zinc, 

cadmium, nickel and lead than either C. tentans or L. 

variegatus. Conversely, the midge was usually comparable to or more sensitive than the amphipod to the 

pesticides tested. Lumbriculus variegatus was not tested 

with several of the pesticides; however, in other studies 

with whole sediments contaminated by DDT and associated metabolites, and in short-term (96-h) experiments 

with organophosphate insecticides (diazinon, 

chlorpyrifos), L. variegatus has proven to be far less 

sensitive than either H. azteca or C. tenfans. These 

results highlight two important points germane to the 

methods in this manual. First, neither of the two test 

species selected for estimating sediment toxicity (I-/. 

azteca, C. fentans) was consistently more sensitive to 

all chemicals, indicating the importance of using multiple 

test organisms when performing sediment assessments. 

Second, L. variegatus appears to be relatively insensitive to most of the test chemicals, which perhaps is a 

positive attribute for an organism used in bioaccumulation tests. 

1.3.8.3.4 Using the data from Table 1.4, sensitivity of H. 

azteca, C. fentans and L. variegatus can be evaluated 

retative to other freshwater species. For this analysis, 

acute and chronic toxicity data from water quality criteria 

(WQC) documents for copper, zinc, cadmium, nickel, 

lead, DDT, dieldrin and chlorpyrifos, and toxicity information from the AQUIRE database (AQUIRE, 1992) for 

DDD and DDE, were compared to assay results for the 

three species (Phipps et al., 1994). The sensitivity of H. 

azteca to metals and pesticides, and C. tentans to 

pesticides was comparable to chronic toxicity data generated for other test species. This was not completely 

6 
unexpected given that the 10-d exposures used for 

these two species are likely more similar to chronic 

partial life-cycle tests than the 48- to 96-h exposures 

traditionally defined as acute in WQC documents. lnterestingly, in some instances (e.g., dieldrin, chlorpyrifos), 

LC50 data generated for H. azteca or C. tentans were 

comparable to or lower than any reported for other 

freshwater species in the WQC documents. This observation likely is a function not only of the test species, but 

of the test conditions; many of the tests on which early 

WQC were based were static, rather than flow-through, 

and utilized unmeasured contaminant concentrations. 

1.3.8.4 Relative species sensitivity frequently varies 

among contaminants; consequently, a battery of tests 

including organisms representing different trophic levels 

may be needed to assess sediment quality (Craig, 1984; 

Williams et al., 1986a; Long et al., 1990; Ingersoll et al., 

1990; Burton and Ingersoll, 1994; USEPA, 1989b). For 

example, Reish (1988) reporled the relative toxicity of 

six metals (As, Cd, Cr, Cu, Hg, and Zn) to crustaceans, 

polychaetes, pelecypods, and fishes and concluded that 

no single species or group of test organisms was the 

most sensitive to all of the metals. 

1.3.8.5 The sensitivity of an organism to contaminants 

should be balanced with the concept of discrimination 

(Burton and Ingersoll, 1994). The response of a test 

organism should provide discrimination between different levels of contamination. 

1.3.8.6 The sensitivity of an organism is related to the 

route of exposure and biochemical response to contaminants. Sediment-dwelling organisms can receive exposure via three primary sources: interstitial water, sediment particles, and overlying water. Food type, feeding 

rate, assimilation efficiency, and clearance rate will control the dose of contaminants from sediment. Benthic 

invertebrates often selectively consume different particle sizes (Harkey et al., 1994) or particles with higher 

organic carbon concentrations that may have higher 

contaminant concentrations. Grazers and other 

collector-gatherers that feed on aufwuchs and detritus 

may receive most of their body burden directly from 

materials attached to sediment or from actual sediment 

ingestion. In amphipods (Landrum, 1989) and clams 

(Boese et al., 1990) uptake through the gut can exceed 

uptake across the gills of certain hydrophobic compounds. Organisms in direct contact with sediment may 

also accumulate contaminants by direct adsorption to 

the body wall or by absorption through the integument 

(Knezovich et al., 1987). 

1.3.8.7 Despite the potential complexities in estimating 

the dose that an animal receives from sediment, the 

toxicity and bioaccumulation of many contaminants in 

sediment such as Kepone@, fluoranthene, organochlorines, and metals have been correlated with either the 

concentration of these chemicals in interstitial water or 

in the case of nonionic organic chemicals, concentrations in sediment on an organic carbon normalized basis 

(Di Toro et al., 1990; Di Toro et al., 1991). The relative 

importance of whole sediment and interstitial water routes 

of exposure depends on the test organism and the 

specific contaminant (Knezovich et al., 1987). Because 

benthic communities contain a diversity of organisms. 

many combinations of exposure routes may be important. Therefore, behavior and feeding habits of a test 

organism can influence its ability to accumulate contaminants from sediment and should be considered 

when selecting test organisms for sediment testing. 

1.3.8.8 The use of H. azteca and C. tentans in laboratory toxicity studies has been confirmed with natural 

benthos populations. 

1.3.8.8.1 Chironomids were not found in sediment 

samples that decreased growth of C. tentans by 30% or 

more in 1 O-d laboratory toxicity tests (Giesy et al., 1988). 

Wentsel et al. (1977a. 1977b. 1978) reported a correlation between effects on C. tentans in laboratory tests 

and the abundance of C. tentans in metal-contaminated 

sediments. 

1.3.8.8.2 Benthic community evaluations and laboratory 

tests with H. azteca both provided evidence of 

metal-induced degradation of aquatic communities in 

the Clark Fork River (Canfield et al., 1994). Total abundance of benthic organisms did not follow a consistent 

pattern when compared to metals in sediment samples. 

The number of chironomid genera was higher at stations 

that showed reduced growth or sexual maturation of H. 

azteca in laboratory sediment tests and had higher 

concentrations of metals in sediment. 

1.3.8.8.3 The results from laboratory sediment toxicity 

tests were compared to colonization of artificial substrates exposed in situ to contaminated Great Lakes 

sediment (Burton and Ingersoll, 1994). Survival or growth 

of H. azteca and C. tentans in lo- to 28-d laboratory 

exposures were negatively correlated to percent chironomids and percent tolerant taxa colonizing artificial 

substrates in the field. Scklekat et al. (1994) reported 

generally good agreement between sediment tests with 

H. azfeca and benthic community responses in the 

Anacostia River, Washington, D.C. 

1.3.8.8.4 Sediment toxicity to amphipods 1r-1 1 O-d toxicity 

tests, field contamination, and field abundance of benthic 

amphipods were examined along a sediment contamination gradient of DDT (Swartz et al., 1994). Survival of 

Eohaustorius estuarius. Rhepoxynius abronius. and H. 

azteca in laboratory toxicity tests was positively correlated to abundance of amphipods in the field and along 

with the survival of H. azteca, was negatively correlated 

to DDT concentrations. The threshold for 1 O-d sediment 

toxicity in laboratory studies was about 300 pg DDT 

(+metabolites)/g organic carbon. The threshold for abundance of amphipods in the field was about 100 pg DDT 

(+metabolites)/g organic carbon. Therefore, correlations 

between toxicity, contamination, and field populations 

indicate that acute sediment toxicity tests can provide 

reliable evidence of biologically adverse sediment con7 
tamination in the field, but may be underprotective of 

chronic effects. 

1.3.9 Selection of Organisms for 

Se&men t Bioaccumula tion Testing 

1.3.9.1 Several studies have demonstrated that hydrophobic organic compounds are bioaccumulated from 

sediment by freshwater infaunal organisms including 

larval insects (C. tentans, Adams et al., 1985; Adams, 

1987; Hexagenla limbata, Gobas et al., 1989), oligochaetes (Tubifex tubifex and Limnodrilus hoffmeisteri; Oliver, 

1984, Oliver. 1987; Connell et al., 1988), and by marine 

organisms (polychaetes, Nephtys incisa; mollusks, 

Mercenaria mercenaria, Yoldia iimatula; Lake et al., 

1990). Consumers of these benthic organisms may 

broaccumulate or biomagnify contaminants. Therefore, 

in addition to sediment toxicity, it may be important to 

examine the uptake of chemicals by aquatic organisms 

from contaminated sediments. 

1.3.9.2 Various species of organisms have been suggested for use in studies of chemical bioaccumulation 

from aquatic sediments. Several criteria should be considered before a species is adopted for routine use in 

these types of studies (Ankley et al., 1992a; Call et al., 

1994). These criteria include (1) availability of organisms throughout the year, (2) known chemical exposure 

hrstory, (3) adequate tissue mass for chemical analyses, 

(4) ease of handling, (5) tolerance of a wide range of 

sedrment phsyico-chemical characteristics (e.g., particle 

size), (6) low sensitivity to contaminants associated with 

sediment (e.g., metals, organics), (7) amenability to 

long-term exposures without adding food, (8) and ability 

to accurately reflect concentrations of contaminants in 

field-exposed organisms (e.g., exposure is realistic). 

With these criteria in mind, the advantages and disadvantages of several potential freshwater taxa for bioaccumulation testing are discussed below. 

1.3.9.3 Freshwater clams provide an adequate tissue 

mass, are easily handled, and can be used in long-term 

exposures. However, few non-exotic freshwater species 

are available for testing. Exposure of clams is uncertain 

because of valve closure. Furthermore, clams are filter 

feeders and may accumulate lower concentrations of 

contaminants compared to detritivores (Lake et al,, 1990). 

Chironomids can be readily cultured, are easy to handle, 

and reflect appropriate routes of exposure. However, 

their rapid life-cycle makes it difficult to perform long-term 

exposures with hydrophobic compounds; also, chironomids can readily biotransform organic compounds such 

as benzo[a]pyrene (Harkey et al., 1994). Larval mayflies 

reflect appropriate routes of exposure, have adequate 

tissue mass for residue analysis, and can be used in 

long-term tests. However, mayflies cannot be continuously cultured in the laboratory and consequently are 

not always available for testing. Furthermore, the background concentrations of contaminants and health of 

field-collected individuals may be uncertain. Amphipods 

(e.g.. H. azteca) can be cultured in the laboratory, are 

easy to handle, and reflect appropriate routes of exposure. However, their size may be insufficient for residue 

analysis and H. azteca are sensitive to contaminants in 

sediment. Fish (e.g., fathead minnows) provide an adequate tissue mass, are readily available, are easy to 

handle, and can be used in long-term exposures. However, the route of exposure is not appropriate for evaluating the bioavailability of sediment-associated contaminants to benthic organisms. 

1.3.9.4 Oligochaetes are infaunal benthic organisms 

that meet many of the test criteria listed above. Certain 

oligochaete species are easily handled and cultured, 

provide reasonable biomass for residue analyses, and 

are tolerant of varying sediment physical and chemical 

characteristics. Oligochaetes are exposed to contaminants via all appropriate routes of exposure including 

pore water and ingestion of sediment particles. Oligochaetes need not be fed during long-term bioaccumulation exposures (Phipps et al., 1993). Various oligochaete species have been used in toxicity and bioaccumulation evaluations (Chapman et al., 1982a, Chapman et 

al., 1982b; Wiederholm, 1987; Kielty et al., 1988a, Kielty 

et al., 1988b; Phipps et al., 1993), and field populations 

have been used as indicators of the pollution of aquatic 

sediments (Brinkhurst, 1980; Spencer, 1980; Oliver, 

1984; Lauritsen, 1985; Robbins et al., 1989; Ankley et 

al., 1992b; E.L. Brunson, NBS, Columbia, MO, unpublished data). 

1.3.9.5 Lumbriculus variegatus does not biotransform 

PAHs (Harkey et al., 1994b). 

1.3.9.6 The response of L. variegafus in laboratory 

bioaccumulation studies has been confirmed with natural populations of oligochaetes. 

1.3.9.6.1 Total PCB concentrations in laboratoryexposed 

L. variegatus were similar to concentrations measured 

in fieldcollected otigochaetes from the same sites (Ankley 

et al., 1992b). PCB homologue patterns also were similar between laboratoryexposed and field-collected otigochaetes. The more highly chlorinated PCBs tended to 

have greater bioaccumulation in the field-collected organisms. In contrast, total PCBs in laboratory-exposed 

(Pimephales promelas) and field-collected (Ictalurus 

me/as) fish revealed poor agreement in bioaccumulation 

relative to the sediment concentrations at the same 

sites. 

1.3.9.6.2 Bioaccumulation of laboratory-exposed L. 

variegatus and field-collected oligochaetes from the same 

sites were also compared (E.L. Brunson, NBS, Columbia, MO, unpublished data). Select PAH and DDT peak 

concentrations were similar in field-collected oligochaetes and L. variegatus exposed for 28 d in the laboratory. 

1.4 Performance-based Criteria 

1.4.1 USEPA’s Environmental Monitoring Management Council (EMMC) recommended the use 

of performance-based methods in developing 
chemical analytical standards (Williams, 1993). 

Performance-based methods were defined by EMMC as 

a monitoring approach that permits the use of appropriate methods that meet pre-established demonstrated 

performance standards (Section 9.2). 

1.4.2 The USEPA Office of Water, Office of Science 

and Technology, and Office of Research and Development held a workshop on September 16-18, 1992 in 

Washington, DC to provide an opportunity for experts in 

the field of sediment toxicology and staff from USEPA’s 

Regional and Headquarters program offices to discuss 

the development of standard freshwater and marine 

sediment testing procedures (USEPA, 1992a and Appendix A). Workgroup participants reached a consensus 

on several culturing and testing methods. In developing 

guidance for culturing freshwater test organisms to be 

included in the USEPA methods manual for sediment 

tests, it was agreed that no single method should be 

required to culture organisms. However, the consensus 

at the workshop was that since the success of a test 

depends on the health of the cultures, having healthy 

test organisms of known quality and age for testing was 

the key consideration. A performance-based criteria approach was selected as the preferred method through 

which individual laboratones should evaluate culture 

methods rather than by control-based criteria. This 

method was chosen to allow each laboratory to optimize 

culture methods and minimize effects of test organism 

health on the reliability and comparability of test results. 

See Tables 11.3, 12.3, and 13.4 for a listing of performance criteria for culturing and testing. 

9 
Section 2 

Summary of Method 

2.1 Method Description and 

Experimental Design 

2.1.1 Method Description 

2.1.1.1 This manual describes procedures for testing 

freshwater organisms in the laboratory to evaluate the 

toxicity or bioaccumulation of contaminants associated 

with whole sediments. Sediments may be collected from 

the field or spiked with compounds in the laboratory. 

Toxicity methods are outlined for two organisms, the 

amphipod Hyalella azteca and the midge Chironomus 

tentans. The toxicity tests are conducted for 10 d in 

300-mL chambers containing 100 ml of sediment and 

175 mL of overlying water. Overlying water is renewed 

dally and test organisms are fed during the toxicity tests. 

The endpoint in the toxicity test with H. azteca is survival 

and the endpoints in the toxicity test with C. tentans are 

survival and growth. Procedures are primarily described 

for testing freshwater sediments; however, estuarine 

sediments (up to 15% salinity) can also be tested with 

H. azteca. Guidance for conducting 28-d bioaccumulation tests with the oligochaete Lumbriculus variegatus is 

provided in this manual. The overlying water is renewed 

daily and the test organisms are not fed during bioaccumulation tests. This guidance describes are also described for determining bioaccumulation kinetics of different classes of compounds during 28-d exposures 

with L. variegatus. 

2.1.2 Experimental Design 

The following section is a general summary of experimental design. See Section 14 for additional detail. 

2.1.2.1 Control and Reference Sediment 

2.1.2.1.1 Sediment tests include a control sediment 

(sometimes called a negative control). A control sediment is a sediment that is essentially free of contaminants and is used routinely to assess the acceptability of 

a test and is not necessarily collected near the site of 

concern. Any contaminants in control sediment are 

thought to originate from the global spread of pollutants 

and do not reflect any substantial input from local or 

nonpoint sources (Lee et al.. 1994). A control sediment 

provides a measure of test acceptability, evidence of 

test organism health. and a basis for interpreting data 

obtained from the test sediments. A reference sediment 

IS collected near an area of concern and is used to 

assess sediment conditions exclusive of material(s) of 

interest. Testing a reference sediment provides a 

site-specific basis for evaluating toxicity. 

2.1.2.1.2 Natural geomorphological and physicochemical characteristics such as sediment texture may influence the response of test organisms (Dewitt et al., 

1988). The physicochemical characteristics of test sediment must be within the tolerance limits of the test 

organism. Ideally, the limits of a test organism should be 

determined in advance; however, controls for factors 

including grain size and organic carbon can be evaluated if the limits are exceeded in a test sediment. See 

Section 10.1 for information on physicochemical requirements of test organisms. If the physicochemical 

characteristics of a test sediment exceed the tolerance 

limits of the test organism it may be desirable to include 

a control sediment that encompasses those characteristics. The effects of sediment characteristics on the results of sediment tests may be able to be addressed with 

regression equations (Dewitt et al., 1988; Ankley et al., 

1994a). The use of formulated sediment can also be 

used to evaluate physicochemical characteristics of sediment on test organisms (Walsh et al., 1991; Suedel and 

Rodgers, 1994). 

2.1.2.2 The experimental design depends on the purpose of the study. Variables that need to be considered 

include the number and type of control sediments, the 

number of treatments and replicates, and water-quality 

characteristics. For instance, the purpose of the study 

might be to determine a specific endpoint such as an 

LC50 and may include a control sediment, a positive 

control, a solvent control, and several concentrations of 

sediment spiked with a chemical. A useful summary of 

field sampling design is presented by Green (1979). See 

Section 14 for additional guidance on experimental design and statistics. 

2.1.2.3 If the purpose of the study is to conduct a 

reconnaissance field survey to identify contaminated 

sites for further investigation, the experimental design 

might include only one sample from each site to allow for 

maximum spatial coverage. The lack of replication at a 

site usually precludes statistical comparisons (e.g., 

ANOVA), but these surveys can be used to identify 

contaminated sites for further study or may be evaluated 

using regression techniques (Sokal and Rohlf, 1981; 

Steel and Torrie. 1980). 

10 
2.1.2.4 In other instances, the purpose of the study 

might be to conduct a quantitative sediment survey to 

determine statistically significant differences between 

effects among control and test sediments from several 

sites. The number of replicates/site should be based on 

the need for sensitivity or power (Section 14). In a 

quantitative survey, replicates (separate samples from 

different grabs collected at the same site) would need to 

be taken at each site. Chemical and physical characterizations of each of these grabs would be required for 

each of these replicates used in sediment testing. Separate subsamples might be used to determine 

within-sample variability or to compare test procedures 

(e.g., comparative sensitivity among test organisms), 

but these subsamples cannot be considered to be true 

field replicates for statistical comparisons among sites 

(ASTM, 1994a). 

2.1.2.5 Sediments often exhibit high spatial and temporal variability (Stemmer et al., 1990a). Therefore, replicate samples may need to be collected to determine 

variance in sediment characteristics. Sediment should 

be collected with as little disruption as possible; however, subsampling, cornpositing, or homogenization of 

sediment samples may be necessary for some experimental designs. 

2.1.2.6 Site locations might be distributed along a 

known pollution gradient, in relation to the boundary of a 

disposal site, or at sites identified as being contaminated 

in a reconnaissance survey. Comparisons can be made 

in both space and time. In pre-dredging studies, a 

sampling design can be prepared to assess the contamination of samples representative of the project area 

to be dredged. Such a design should include subsampling 

cores taken to the project depth. 

2.1.2.7 The primary focus of the physical and experimental test design, and statistical analysis of the data, is 

the experimental unit. The experimental unit is defined 

as the smallest physical entity to which treatments can 

be independently assigned (Steel and Torrie, 1980) and 

to which air and water exchange between test chambers 

are kept to a minimum. As the number of test chambers/ 

treatment increases, the number of degrees of freedom 

increases, and, therefore, the width of the confidence 

interval on a point estimate, such as an LC50, decreases, and the power of a significance test increases 

(Section 14). Because of factors that might affect results 

within test chambers and results of a test, all test chambers should be treated as similarly as possible. Treatments should be randomly assigned to individual test 

chamber locations. Assignment of test organisms to test 

chambers should be non-biased. 

2.2 Types of Tests 

2.2.1 Toxicity methods are outlined for two organisms, 

the amphipod H. azteca (Section 11) and the midge C. 

tentans (Section 12). This manual primarily describes 

methods for testing freshwater sediments; however, the 

methods described can also be used for testing H. 

azteca in estuarine sediments (up to 15 % salinity). 

2.2.2 Guidance for conducting 28-d bioaccumulation 

tests with the oligochaete L. variegates is described in 

Section 13. Methods are also described for determining 

bioaccumulation kinetics of different classes of compounds during 28-d exposures with L. variegatus. 

2.3 Test Endpoints 

2.3.1 The endpoints measured in the toxicity tests are 

survival or growth (the growth endpoint is optional in the 

H. azteca test). Endpoints measured in bioaccumulation 

tests are tissue concentrations of contaminants and for 

some types of studies, lipid content. Behavior of test 

organisms should be qualitatively observed daily in all 

tests (e.g., avoidance of sediment). 

11 
Section 3 

Definitions 

3.1 Terms 

The following terms were defined in Lee (1980), NRC 

(1989). USEPA (1989c), USEPA-USCOE (1991), 

USEPA-USCOE (1994), Lee et al. (1994), ASTM (1993b), 

or ASTM (1994a). 

3.1.1 Technical Terms 

3.1.1.1 Sediment. Particulate material that usually lies 

below water. Formulated particulate material that is 

intended to lie below water in a test. 

3.1.1.2 Contaminated sediment. Sediment containing 

chemical substances at concentrations that pose a known 

or suspected threat to environmental or human health. 

3.1.1.3 Whole sediment. Sediment and associated 

pore water that have had minimal manipulation. The 

term bulk sediment has been used synonymously with 

whole sediment. 

3.1.1.4 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 does not reflect any substantial input from 

local or nonpoint sources. Comparing test sediments to 

control sediments is a measure of the toxicity of a test 

sediment beyond inevitable background contamination. 

3.1.1.5 Reference sediment. A whole sediment near 

an area of concern used to assess sediment conditions 

exclusive of material(s) 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. This is the manner in which 

reference sediment is used in dredge material evaluations. 

3.1.1.6 lnterstitial water or pore water. Water occupying space between sediment or soil particles. 

3.1.1.7 Spiked sediment A sediment to which a material has been added for experimental purposes. 

3.1.1.8 Reference toxicity test. A test with a high-grade 

reference material conducted in conjunction with sediment tests to determine possible changes in condition of 

the test organisms. Deviations outside an established 

normal range indicate a change in the condition of the 

test organism population. Reference-toxicity tests are 

most often performed in the absence of sediment. 

3.1.1.9 Clean. Denotes a sediment or water that does 

not contain concentrations of test materials which cause 

apparent stress to the test organisms or reduce their 

survival. 

3.1.1.10 Overlying water. The water placed over sediment in a test chamber during a test. 

3.1.1.11 Concentration. The ratio of weight or volume 

of test material(s) to the weight or volume of sediment. 

3.1.1.12 No-observable-effect concentration (NOEC). 

The highest concentration of a toxicant to which organisms are exposed in a test that causes no observable 

adverse effect on the test organisms (i.e., the highest 

concentration of a toxicant in which the value for the 

observed response is not statistically significant different 

from the controls). 

3.1.1.13 Lowest-observable-effect concentration 

(LOEC). The lowest concentration of a toxicant to which 

organisms are exposed in a test that causes an adverse 

effect on the test organisms (i.e., where a significant 

difference exists between the value for the observed 

response and that for the controls). 

3.1.1.14 Lethal concentration (LC). The toxicant concentration that would cause death in a given percent of 

the test population. Identical to EC when the observable 

adverse effect is death. For example, the LC50 is the 

concentration of toxicant that would cause death in 50% 

of the test population. 

3.1.1.15 Effect concentration (EC). The toxicant concentration that would cause an effect in a given percent 

of the test population. Identical to LC when the observable adverse effect is death. For example, the EC50 is 

the concentration of toxicant that would cause death in 

50% of the test population. 

12 
3.1.1.16 lnhibition concentration (IC). The toxicant 

concentration that would cause a given percent reduction in a non-quantal measurement for the test population. For example, the lC25 is the concentration of 

toxicant that would cause a 25% reduction in growth for 

the test population, and the IC50 is the concentration of 

toxicant that would cause a 50% reduction. 

3.1-1.17 Biota-sediment accumulation factor (BSAF). 

The ratio of tissue residue to source concentration (e.g., 

sediment at steady state normalized to lipid and sediment organic carbon). 

3.1.1.18 Bioaccumulation. The net accumulation of a 

substance by an organism as a result of uptake from all 

environmental sources. 

3.1.1.19 Bioaccumulation factor. Ratio of tissue residue to contaminant source concentration at steady-state. 

3.1.1.20 Bioaccumulation potential. Qualitative assessment of whether a contaminant is bioavailable. 

3.1.1.21 Bioconcentration. The net assimilation of a 

substance by an aquatic organism as a result of uptake 

directly from aqueous solution. 

3.1.1.22 Bioconcentration factor (BCF). Ratio of tissue residue to water contaminant concentration at 

steady-state. 

3.1.1.23 Depuration. Loss of a substance from an 

organism as a result of any active (e.g., metabolic 

breakdown) or passive process when the organism is 

placed into an uncontaminated environment. Contrast 

with Elimination. 

3.1.1.24 Elimination. General term for the loss of a 

substance from an organism that occurs by any active or 

passive means. The term is applicable either in a contaminated environment (e.g., occurring simultaneously 

with uptake) or in a clean environment. Contrast with 

Depuration. 

3.1.1.25 k,. Uptake rate coefficient from the aqueous 

phase, with units of g-water x g-tissue-’ x time-‘. Contrast with ks. 

3.1.1.26 ks. Sediment uptake rate coefficient from the 

sediment phase, with units of g-sediment x g-tissue- x 

time-‘. Contrast with k,. 

3.1.1.27 k2, Elimination rate constant, with units of 

time-‘. 

3.1.1.28 Kinetic Bioaccumulation Model. Any model 

that uses uptake and/or elimination rates to predict 

tissue residues. 

3.1.1.29 Koc. Organic carbon-water partitioning coefficient. 

3.1.1.30 Kow, Octanol-water partitioning coefficient. 

3.1.1.31 Steady-stare. An equilibrium or “constant” 

tissue residue resulting from the balance of the flux of 

compound into and out of the organism. Operationally 

determined by no statistically significant difference in 

tissue residue concentrations from three consecutive 

sampling periods. 

3.1.2 Grammatical Terms 

3.1.2.1 The words “must,” “should,” “may.” “can.” and 

“might” have very specific meanings in this manual. 

3.1.2.2 “Must” is used to express an absolute requirement, that is, to state that a test ought to be designed to 

satisfy the specified conditions, unless the purpose of 

the test requires a different design. “Must” is only used 

in connection with the factors that directly relate to the 

acceptability of a test. 

3.1.2.3 “Should” is used to state that the specified 

condition is recommended and ought to be met if possible. Although a violation of one “should” is rarely a 

serious matter, violation of several will often render the 

results questionable. 

3.1.2.4 Terms such as “is desirable,” “is often desirable,” and “might be desirable” are used in connection 

with less important factors. 

3.1.2.5 “May” is used to mean “is (are) allowed to.” 

“can” is used to mean “is (are) able to,” and “might” is 

used to mean “could possibly. “Thus, the classic distinction between “may” and “can” is preserved, and “might” 

is never used as a synonym for either “may” or “can.” 

13 
Section 4 

Interferences 

4.1 General Introduction 

4.1.1 Interferences are characteristics of a sediment or 

sediment test system that can potentially affect test 

organism survival aside from those related to 

sediment-associated contaminants. These interferences 

can potentially confound interpretation of test results in 

two ways: (1) toxicity is observed in the test when 

contamination is not present, or there is more toxicity 

than expected; and (2) no toxicity or bioaccumulation is 

observed when contaminants are present at elevated 

concentrations, or there is less toxicity or bioaccumulation than expected. 

4.1.2 There are three categories of interfering factors: 

those characteristics of sediments affecting survival independent of chemical concentration (i.e., 

non-contaminant factors); changes in chemical bioavailability as a function of sediment manipulation or storage; 

and the presence of indigenous organisms. Although 

test procedures and test organism selection criteria 

were developed to minimize these interferences, this 

section describes the nature of these interferences. 

4.1.3 Because of the heterogeneity of natural sediments, extrapolation from laboratory studies to the field 

can sometimes be difficult (Table 4.1; Burton, 1991). 

Sediment collection, handling, and storage may alter 

bioavailability and concentration by changing the physical, chemical, or biological characteristics of the sediment. Maintaining the integrity of a field-collected sediment during removal, transport, mixing, storage, and 

testing is extremely difficult and may complicate the 

Interpretation of effects. Direct comparisons of organisms exposed in the laboratory and in the field would be 

useful to verify laboratory results. However, spiked sediment may not be representative of contaminated sediment in the field. Mixing time (Stemmer et al., 1990a) 

and aging (Word et al., 1987; Landrum, 1989; Landrum 

and Faust, 1992) of spiked sediment can affect responses of organisms. 

4.1.3.1 Laboratory sediment testing with field-collected 

sediments may be useful in estimating cumulative effects and Interactions of multiple contaminants in a 

sample. Tests with field samples usually cannot discriminate between effects of individual chemicals. Most 

sediment samples contain a complex matrix of inorganic 

Table 4.1 Advantages and Disadvantages for Use of Sediment 

Tests1 

Advantages 

Measure bioavailable fraction of contaminant(s). 

Provide a direct measure of benthic effects, assuming no field 

adaptation or amelioration of effects. 

Limited special equipment is required. 

Methods are rapid and Inexpensive. 

Legal and scientific precedence exists for use; ASTM standard 

guides are available. 

Measure unique information relative to chemical analyses or 

benthic community analyses. 

Tests with spiked chemicals provide data on cause-effect 

relationships. 

Sediment-toxicity tests can be applied to all chemicals of 

concern. 

Tests applied to field samples reflect cumulative effects of 

contaminants and contaminant interactions. 

Toxicity tests are amenable to confirmation with natural benthos 

populations. 

Disadvantages 

Sediment collection, handling, and storage may alter bioavailability. 

Spiked sediment may not be representative of field contaminated sediment. 

Natural geochemical characteristics of sediment may affect the 

response of test organisms. 

Indigenous animals may be present in field-collected sediments. 

Route of exposure may be uncertain and data generated in 

sediment toxicity tests may be difficult to Interpret if factors 

controlling the bioavailability of contaminants in sediment are 

unknown. 

Tests applied to field samples may not discriminate effects of 

individual chemicals. 

Few comparisons have been made of methods or species. 

Only a few chronic methods for measuring sublethal effects 

have been developed or extensively evaluated 

Laboratory tests have Inherent limitations in predicting ecological effects. 

1 Modified from Swartz (1989) 

and organic contaminants with many unidentified compounds. The use of Toxicity Identification Evaluations 

(TIE) in conjunction with sediment tests with spiked 

chemicals may provide evidence of causal relationships 

and can be applied to many chemicals of concern 

(Ankley and Thomas, 1992; Adams et al., 1985). Sedi14 
ment spiking can also be used to investigate additive, 

antagonistic, or synergistic effects of specific contaminant mixtures in a sediment sample (Swartz et al., 

1988). 

4.1.4 Methods that measure sublethal effects are either 

not available or have not been routinely used to evaluate 

sediment toxicity (Craig, 1984; Dillon and Gibson, 1986; 

Ingersoll and Nelson, 1990; Ingersoll. 1991; Button et 

al., 1992). Most assessments of contaminated sediment 

rely on short-term-lethality testing methods (e.g., [ 10 d; 

USEPA-USCOE, 1977; USEPA-USCOE, 1991). Shortterm-lethality tests are useful in identifying “hot spots” of 

sediment contamination but may not be sensitive enough 

to evaluate moderately contaminated areas. However, 

sediment quality assessments using sublethal responses 

of benthic organisms such as effects on growth and 

reproduction have been used to successfully evaluate 

moderately contaminated areas (Scott, 1989). Additional 

methods development of chronic sediment testing procedures and culturing of infaunal organisms with a variety of feeding habits including suspension and deposit 

feeders is needed. 

4.1.5 Despite the interferences discussed in this section, existing sediment testing methods can be used to 

provide a rapid and direct measure of effects of contaminants on benthic communities. Laboratory tests with 

field-collected sediment can also be used to determine 

temporal, horizontal, or vertical distribution of contaminants in sediment. Most tests can be completed within 

two to four weeks. Legal and scientific precedents exist 

for use of toxicity and bioaccumulation tests in regulatory decision-making (e.g., USEPA, 1986a). Furthermore, sediment tests with complex contaminant mixtures are important tools for making decisions about the 

extent of remedial action for contaminated aquatic sites 

and for evaluating the success of remediation activities. 

4.2 Non-Contaminant Factors 

4.2.1 Results of sediment tests can be used to predict 

effects that may occur with aquatic organisms in the field 

as a result of exposure under comparable conditions. 

Yet motile organisms might avoid exposure in the field. 

Photoinduced toxicity caused by ultraviolet (UV) light, 

may be important for some compounds associated with 

sediment (e.g., polycyclic aromatic hydrocarbons (PAHs); 

Davenport and Spacie, 1991; Ankley et al.. 1994b). 

Fluorescent light does not contain UV light, but natural 

sunlight does. Lighting can therefore affect toxicological 

responses and is an important experimental variable for 

photoactivated chemicals. However, lighting typically 

used to conduct laboratory tests does not include the 

appropriate spectrum of ultraviolet radiation to photoactivate compounds (Oris and Giesy, 1985), and thus 

laboratory tests may not account for toxicity expressed 

by this mode of action. 

4.2.2 Natural geomorphological and physicochemical 

characteristics such as sediment texture may influence 

the response of test organisms (Dewitt et al., 1988). 

The physicochemical characteristics of test sediment 

need to be within the tolerance limits of the test organism. Ideally, the limits of the test organism should be 

determined in advance: however, control samples reflecting differences in factors such as grain size and 

organic carbon can be evaluated if the limits are exceeded in the test sediment (Section 10.1). The effects 

of sediment characteristics can also be addressed with 

regression equations (Dewitt et al., 1988; Ankley et al.. 

1994a). The use of formulated sediment can also be 

used to evaluate physicochemical characteristics of sediment on test organisms (Walsh et al., 1991; Suedel and 

Rodgers, 1994). 

4.2.3 Interferences of tests with each specific species 

are described in Tables 11.3, 12.3, and 13.4. 

4.3 Changes in Bioavailability 

4.3.1 Sediment toxicity tests are meant to serve as an 

indicator of contaminant-related toxicity that might be 

expected under field or natural conditions. Although the 

tests are not designed to simulate natural conditions, in, 

some cases contaminant availability in laboratory toxicity test may be different from what it is representative of 

in-place sediments in the field. 

4.3.2 Sediment collection, handling, and storage may 

alter contaminant bioavailability and concentration by 

changing the physical, chemical, or biological characteristics of the sediment. These manipulation processes 

are generally thought to increase availability of organic 

compounds because of disruption of the equilibrium with 

organic carbon in the pore water/particle system. Similarly, oxidation of anaerobic sediments increases the 

availability of certain metals (Di Toro et al.. 1990). 

Because the availability of contaminants may be a function of the degree of manipulation, this manual recommends that handling, storage, and preparation of the 

sediment for actual testings be as consistent as possible. If sieving is performed, it is done primarily to 

remove predatory organisms and large debris. This 

manipulation most likely results in a worst-case condition of heightened bioavailability yet eliminates predation as a factor that might confound test results. When 

sediments are sieved, it may be desirable to take samples 

before and after sieving (e.g., pore-water metals or 

DOC, AVS, TOC) to document the influence of sieving 

on sediment chemistry. USEPA does not recommend 

sieving sediments on a routine basis. 

4.3.3 Testing sediments at temperatures different from 

the field might affect contaminant solubility. partitioning 

coefficients, or other physical and chemical characteristics. Interaction between sediment and overlying water 

and the ratio of sediment to overlying water may influence bloavailability (Stemmer et al., 1990b). 

15 
4.3.4 The addition of food, water, or solvents to the test 

chambers might obscure the bioavailability of contaminants in sediment or might provide a substrate for 

bacterial or fungal growth. Without addition of food, the 

test organisms may starve during exposures (Ankley et 

al., 1994). However, the addition of food may alter the 

availability of the contaminants in the sediment 

(Wiederholm et al., 1987, Harkey et al., 1994) depending on the amount of food added, its composition (e.g., 

TOC), and the chemical(s) of interest. 

4.3.5 Depletion of aqueous and sediment-sorbed contaminants resulting from uptake by an organism or test 

chamber may also influence availability. In most cases, 

the organism is a minor sink for contaminants relative to 

the sediment. However, within the burrow of an organism, sediment desorption kinetics may limit uptake rates. 

Within minutes to hours, a major portion of the total 

chemical may be inaccessible to the organisms because 

of depletion of available residues. The desorption of a 

particular compound from sediment may range from 

easily reversible (labile; within minutes) to irreversible 

(non-labile; within days or months; Karickhoff and Morris, 

1985). Interparticle diffusion or advection and the quality 

and quantity of sediment organic carbon can also affect 

sorption kinetics. 

4.3.6 The route of exposure may be uncertain, and data 

from sediment tests may be difficult to interpret if factors 

controlling the bioavailability of contaminants in sediment are unknown. Bulk-sediment chemical concentrations may be normalized to factors other than dry weight. 

For example, concentrations of nonionic organic compounds might be normalized to sediment organic-carbon 

content (USEPA, 1992c) and certain metals normalized 

to acid volatile sulfides (Di Toro et al., 1990). Even with 

the appropriate normalizing factors, determination of 

toxic effects from ingestion of sediment or from dissolved chemicals in the interstitial water can still be 

difficult (Lamberson and Swartz, 1988). 

4.4 Presence of Indigenous Organisms 

4.4.1 Indigenous organisms may be present in 

field-collected sediments. An abundance of the same 

organism or organisms taxonomically similar to the test 

organism in the sediment sample may make interpretation of treatment effects difficult. For example, growth of 

amphipods, midges, or mayflies may be reduced if high 

numbers of oligochaetes are in a sediment sample 

(Reynoldson et al., 1994). Previous investigators have 

inhibited the biological activity of sediment with sieving, 

heat, mercuric chloride, antibiotics, or gamma irradiation 

(ASTM, 1994b; K.E. Day Environment Canada, 

Burlington, Ontario, personal communication). However, 

further research is needed to determine effects on contaminant bioavailability or other modifications of sediments from treatments such as those used to remove or 

destroy indigenous organisms. 

16 
Section 5 

Health, Safety, and Waste Management 

5.1 General Precautions 

5.1.1 Development and maintenance of an effective 

health and safety program in the laboratory requires an 

ongoing commitment by laboratory management and 

includes (1) the appointment of a laboratory health and 

safety officer with the responsibility and authority to 

develop and maintain a safety program, (2) the preparation of a formal written health and safety plan, which is 

provided to each laboratory staff member, (3) an ongoing training program on laboratory safety, and (4) regular safety inspections. 

5.1.2 This manual addresses procedures that may 

involve hazardous materials, operations, and equipment, 

and it does not purport to address all of the safety 

problems associated with their use. It is the responsibility of the user to establish appropriate safety and health 

practices, and determine the applicability of regulatory 

limitations before use. While some safety considerations 

are included in this manual, it is beyond the scope of this 

manual to encompass all safety requirements necessary to conduct sediment tests. 

5.1.3 Collection and use of sediments may involve 

substantial risks to personal safety and health. Contaminants in field-collected sediment may include carcinogens, mutagens, and other potentially toxic compounds. 

Inasmuch as sediment testing is often begun before 

chemical analyses can be completed, worker contact 

with sediment needs to be minimized by (1) using gloves, 

laboratory coats, safety glasses, face shields, and respirators as appropriate, (2) manipulating sediments under 

a ventilated hood or in an enclosed glove box, and (3) 

enclosing and ventilating the exposure system. Personnel collecting sediment samples and conducting tests 

should take all safety precautions necessary for the 

prevention of bodily injury and illness that might result 

from ingestion or invasion of infectious agents. inhalation or absorption of corrosive or toxic substances through 

skin contact, and asphyxiation because of lack of oxygen or presence of noxious gases. 

5.1.4 Before sample collection and laboratory work, 

personnel should determine that all required safety equipment and materials have been obtained and are in good 

condition. 

5.2 Safety Equipment 

5.2.1 Personal Safety Gear 

5.2.1.1 Personnel should use safety equipment. such 

as rubber aprons, laboratory coats, respirators, gloves. 

safety glasses, face shields, hard hats, and safety shoes. 

5.2.2 Laboratory Safety Equipment 

5.2.2.1 Each laboratory should be provided with safety 

equipment such as first aid kits, fire extinguishers. fire 

blankets, emergency showers, and eye fountains. 

5.2.2.2 Mobile laboratories should be equipped with a 

telephone to enable personnel to summon help in case 

of emergency. 

5.3 General Laboratory and Field 

Operations 

5.3.1 Special handling and precautionary guidance in 

Material Safety Data Sheets should be followed for 

reagents and other chemicals purchased from supply 

houses. 

5.3.2 Work with some sediments may require compliance with rules pertaining to the handling of hazardous 

materials. Personnel collecting samples and performing 

tests should not work alone. 

5.3.3 It is advisable to wash exposed parts of the body 

with soap and water immediately after collecting or 

manipulating sediment samples. 

5.3.4 Strong acids and volatile organic solvents should 

be used in a fume hood or under an exhaust canopy 

over the work area. 

5.3.5 An acidic solution should not be mixed with a 

hypochlorite solution because hazardous vapors might 

be produced. 

5.3.6 To prepare dilute acid solutions, concentrated 

acid should be added to water, not vice versa. Opening 

a bottle of concentrated acid and adding concentrated 

acid to water should be performed only in a fume hood. 

17 
5.3.7 Use of ground-fault systems and leak detectors IS 

strongly recommended to help prevent electrical shocks. 

Electrical equipment or extension cords not bearing the 

approval of Underwriter Laboratories should not be used. 

Ground-fault interrupters should be installed in all “wet” 

laboratories where electrical equipment is used. 

5.3.8 All containers should be adequately labeled to 

Identify their contents. 

5.3.9 Good housekeeping contributes to safety and 

reliable results. 

5.4 Disease Prevention 

5.4.1 Personnel handling samples that are known or 

suspected to contain human wastes should be given the 

opportunity to be immunized against hepatitis B, tetanus, typhoid fever, and polio. 

5.5 Safety Manuals 

5.5.1 For further guidance on safe practices when 

handling sediment samples and conducting toxicity tests, 

check with the permittee and consult general industrial 

safety manuals including USEPA (1986b) and Walters 

and Jameson (1984). 

5.6 Pollution Prevention, Waste 

Management, and Sample Disposal 

5.6.1 It is the laboratory’s responsibility to comply with 

the federal, state, and local regulations governing the 

waste management, particularly hazardous waste identification rules and land disposal restrictions, and to 

protect the air, water and land by minimizing and controlling all releases from fume hoods and bench operations. Also, compliance is required with any sewage 

discharge permits and regulations. For further information on waste management, consult “The Waste Management Manual for Laboratory Personnel” available 

from the American Chemical Society’s Department of 

Government Relations and Science Policy, 1155 16th 

Street N.W., Washington, D.C. 20036. 

5.6.2 Guidelines for the handling and disposal of hazardous materials should be strictly followed. The federal 

government has published regulations for the management of hazardous waste and has given the states the 

option of either adopting those regulations or developing 

their own. If states develop their own regulations, they 

are required to be at least as stringent as the federal 

regulations. As a handler of hazardous materials, it is 

your responsibility to know and comply with the pertinent 

regulations applicable in the state in which you are 

operating. Refer to The Bureau of National Affairs Inc., 

(1986) for the citations of the federal requirements. 

18 
Section 6 

Facilities, Equipment, and Supplies 

6.1 General 

6.1.1 Before a sediment test is conducted in any test 

facility, it is desirable to conduct a “non-toxicant” test 

with each potential test species, in which all test chambers contain a control sediment (sometimes called the 

negative control), and clean overlying water for each 

organism to be tested. Survival, growth, or reproduction 

of the test organism will demonstrate whether facilities, 

water, control sediment, and handling techniques are 

adequate to result in acceptable species-specific control 

numbers. Evaluations may also be made on the magnitude of between-chamber variance in a test. 

6.2 Facilities 

6.2.1 The facility must include separate areas for culturing and testing to reduce the possibility of contamination 

by test materials and other substances, especially volatile compounds. Holding and culture chambers should 

not be in a room in which sediment tests are conducted, 

where stock solutions or sediments are prepared, or 

equipment is cleaned. Test chambers may be placed in 

a temperature-controlled recirculating water bath or a 

constant-temperature area. An enclosed test system is 

desirable to provide ventilation during tests to limit exposure of laboratory personnel to volatile substances. 

6.2.2 Light of the quality and luminance normally obtained in the laboratory is adequate (about 500 to 1000 

Iux using wide-spectrum fluorescent lights; e.g., 

cool-white or daylight) for culturing and testing. Lux is 

the unit selected for reporting luminance in this manual. 

Multiply units of lux by 0.093 to convert to units of foot 

candles. Multiply units of lux by 6.91 x 10-3 to convert to 

units of μElm2/s1 (assuming an average wavelength of 

550 nm (μmol-2 s-1 = W m x l(nm) x 8.36 x 10-3) (ASTM, 

1994c). Luminance should be measured at the surface 

of the water. A uniform photoperiod of 16L:8D can be 

achieved in the laboratory or in an environmental chamber using automatic timers. 

6.2.3 During phases of rearing, holding, and testing, 

test organisms should be shielded from external disturbances such as rapidly changing light or pedestrian 

traffic. 

6.2.4 The test facility should be well ventilated and free 

of fumes. Air used for aeration should be free of oil and 

fumes. Filters to remove oil, water, and bacteria are 

desirable. Oil-free air pumps should be used where 

possible. Particulates can be removed from the air using 

filters such as BALSTON® Grade BX or equivalent 

(Balston, Inc., Lexington, MA), and oil and other organic 

vapors can be removed using activated carbon filters 

(e.g., BALSTON@, C-l filter, or equivalent). Laboratory 

ventilation systems should be checked to ensure that 

return air from chemistry laboratories or sample handling areas is not circulated to culture or testing rooms, 

or that air from testing rooms does not contaminate 

culture rooms. Air pressure differentials between rooms 

should not result in a net flow of potentially contaminated air to sensitive areas through open or loosely 

fitting doors. 

6.3 Equipment and Supplies 

6.3.1 Equipment and supplies that contact stock solutions, sediments, or overlying water should not contain 

substances that can be leached or dissolved in amounts 

that adversely affect the test organisms. In addition, 

equipment and supplies that contact sediment or water 

should be chosen to minimize sorption of test materials 

from water. Glass, type 316 stainless steel, nylon, 

high-density polyethylene, polycarbonate, and fluorocarbon plastics should be used whenever possible to 

minimize leaching, dissolution, and sorption. Concrete 

and high-density plastic containers may be used for 

holding and culture chambers, and in the water-supply 

system. These materials should be washed in detergent, acid rinsed, and soaked in flowing water for a week 

or more before use. Cast-iron pipe should not be used in 

water-supply systems because colloidal iron will be added 

to the overlying water and strainers will be needed to 

remove rust particles. Copper, brass, lead. galvanized 

metal, and natural rubber should not contact overlying 

water or stock solutions before or during a test. Items 

made of neoprene rubber and other materials not mentioned above should not be used unless it has been 

shown that their use will not adversely affect survival, 

growth, or reproduction of the test organisms. 

6.3.2 New lots of plastic products should be tested for 

toxicity by exposing organisms to them under ordinary 

test conditions before general use. 

19 
6.3.3 General Equipment 

6.3.3.1 Environmental chamber or equivalent facility 

with photoperiod and temperature control (20 to 25°C). 

6.3.3.2 Water purification system capable of producing 

at least 1 mega-ohm water (USEPA, 1993a). 

6.3.3.3 Analytical balance capable of accurately weighing to 0.01 mg. 

6.3.3.4 Reference weights, Class S-for documenting 

the performance of the analytical balance(s). The 

balance(s) should be checked with reference weights 

that are at the upper and lower ends of the range of the 

weighings made when the balance is used. A balance 

should be checked at the beginning of each series of 

weighings, periodically (such as every tenth weight) 

during a long series of weighings, and after taking the 

last weight of a series. 

6.3.3.5 Volumetric flasks and graduated cylindersClass A, borosilicate glass or nontoxic plastic labware, 

10 to 1000 mL for making test solutions. 

6.3.3.6 Volumetric pipets-Class A, 1 to 100 mL. 

6.3.3.7 Serological pipets-1 to 10 mL, graduated. 

6.3.3.8 Pipet bulbs and fillers-PROPIPET® or equivalent. 

6.3.3.9 Droppers, and glass tubing with fire polished 

edges, 4 to 6 mm ID-for transferring test organisms. 

6.3.3.10 Wash bottles-for rinsing small glassware, 

instrument electrodes and probes. 

6.3.3.11 Glass or electronic thermometers-for measuring water temperature. 

6.3.3.12 National Bureau of Standards Certified thermometer (see USEPA Method 170.1; USEPA, 1979b). 

6.3.3.13 Dissolved oxygen (DO), pH/selective ion, and 

specific conductivity meters and probes for routine physical and chemical measurements are needed. Unless a 

test is being conducted to specifically measure the effect 

of DO or conductivity, a portable field-grade instrument 

is acceptable. 

6.3.3.14 See Table 6.1 for a list of additional equipment 

and supplies. 

6.3.4 Water-delivery System 

6.3.4.1 The waterdelivery system used in water-renewal 

testing can be one of several designs. The system 

should be capable of delivering water to each replicate 

test chamber. Mount and Brungs (1967) diluters have 

20 

been successfully modified for sediment testing. Other 

diluter systems have also been useful (Ingersoll and 

Nelson, 1990; Maki, 1977; Benoit et al., 1993; Zumwalt 

et al., 1994). The water-delivery system should be calibrated before the test by determining the flow rate of the 

overlying water. The general operation of the system 

should be visually checked daily throughout the length 

of the test. If necessary, the water-deliver-y system should 

be adjusted during the test. At any particular time during 

the test, flow rates through any two test chambers 

should not differ by more than 10%. 

6.3.4.2 The overlying water can be replaced manually 

(e.g., siphoning); however, manual systems take more 

time to maintain during a test- In addition, automated 

systems generally result in less suspension of sediment 

compared to manual renewal. 

6.3.5 Test Chambers 

6.3.5.1 Test chambers may be constructed in several 

ways and of various materials, depending on the experimental design and the contaminants of interest. Clear 

silicone adhesives, suitable for aquaria, sorb some organic compounds that might be difficult to remove. 

Therefore, as little adhesive as possible should be in 

contact with the test material. Extra beads of adhesive 

should be on the outside of the test chambers rather 

than on the inside. To leach potentially toxic compounds 

from the adhesive, all new test chambers constructed 

using silicone adhesives should be held at least 48 h in 

overlying water before use in a test. 

6.3.5.2 Test chambers for specific tests are described in 

Sections 11, 12, and 13. 

6.3.6 Cleaning 

6.3.6.1 All non-disposable sample containers, test chambers, and other equipment that have come in contact 

with sediment should be washed after use in the manner 

described below to remove surface contaminants. 

1. Soak 15 min in tap water, and scrub with detergent, 

or clean in an automatic dishwasher. 

2. Rinse twice with tap water. 

3. Carefully rinse once with fresh, dilute (10%, V:V) 

hydrochloric or nitric acid to remove scale, metals, 

and bases. To prepare a 10% solution of acid, add 

10 mL of concentrated acid to 90 ml of deionized 

water. 

4. Rinse twice with deionized water. 

5. Rinse once with full-strength, pesticide-grade acetone to remove organic compounds (use a fume 

hood or canopy). 

6. Rinse three times with deionized water. 
Table 6.1 Equipment and Supplies for Culturing and Testing Specific Test Organisms 

A. Biological Supplies 

Brood stock 01 test orgamsms 

Actlve dry yeast (HA) 

Cerophym (dried cereal leaves; HA) 

Trout food pellets (HA) 

TetratinB goldfish food (CT) 

Trout starter (LV) 

Helisoma sp. snails (optional; LV) 

Algae (e.g _, Selenasfrum capricornulum. Chlorella; CT) 

Dratoms (e.g., Nawcula sp; HA) 

B. Glassware 

Culture chambers 

Test chambers (300-mL high-form ltpless beaker: HA and CT) 

Test chambers (15.8 x 29.3- x 11.7~cm, W x L x H; LV) 

Juvemle holding beakers (e.g.. 1 L; HA) 

Crystallizing oishes or beakers (200- to 300-mL; CT) 

Erlenmeyer flasks (250 and 500 mL; CT) 

Larval rearmg chambers (e.g., 19-L capacity: CT) 

l/4” glass tubing (for asprrating flask; CT) 

Glass bowls (20-cm diameter; LV) 

Glass vials (10 mL; LV) 

Wide-bore pipets (4 to 6 mm ID) 

Glass drsposable ptpets 

Burettes (for hardness and alkalinity determinations) 

Graduated cylrnders (assorted sizes, 10 mL to 2 L) 

C. Instruments and Equipment 

Dissecting mrcroscope 

Stainless-steel sieves (e.g., U.S. Standard No. 2530 

3540.50 mesh) 

Delivery system for overlying water (See Appendrx B for a listing 

of equipment needed for water delivery systems) 

Photoperiod tamers 

Light meter 

Temperature controllers 

Thermometer 

Contrnuous recording thermometers 

Dissolved oxygen meter 

pH meter 

Ion-specrfic meter 

Ammonia electrode (or ammonia test kit) 

Specrficconductance meter 

Dryrng oven 

Desiccator 

Balance (0.01 mg sensitrvrty) 

C. Instruments and Equipment 

Blender 

Refrigerator 

Freezer 

Light box 

Hemacytometer (HA) 

Paper shredder, cutter, or scrssors (CT, LV) 

Ttssue homogenizer (LV) 

Electric dnll with starnless steel auger (diameter 7.6 cm, 

overall length 38 cm, auger brt length 25.4 cm (Sectron 8.3) 

D. Miscellaneous 

Ventrlation system for test chambers 

Air supply and airstones (011 free and regulated) 

Cotton surgical gauze or cheese cloth (HA) 

Stainless-steel screen (no. 60 mesh, for test chambers) 

Glass hole-cutting bits 

Stlicon adhesive caulking 

Plastic mesh (110 pm mesh opening; NytexB 110; HA) 

Alummum-weighmg pans 

Fluorescent-lrght bulbs 

Nalgene bottles (500 mL and 1000 mL for food preparation and 

storage) 

Deionized water 

Airline tubing 

Whrte plastic dish pan 

“Coiled-web material” (3-M, St. Paul. MN; HA) 

White paper towelrng (for substrate; CT) 

Brown paper toweling (for substrate; LV) 

Screening material (e.g., Nitex@ (110 mesh), window screen, or 

panty hose; CT) 

Water squirt bottle 

Dissecting probes (LV) 

Dental picks (LV) 

Shallow pans (plastic (lightcolored). glass, stamless steel) 

E. Chemicals 

Detergent (non-phosphate) 

Acetone (reagent grade) 

Hexane (reagent grade) 

Hydrochloric acid (reagent grade) 

Chloroform (LV) 

Methanol (LV) 

Copper Sulfate 

Potassium Chloride 

Reagents for reconstituting water 

Formalin (or Notox@) 

Sucrose 

HA = Hyalella azteca 

CT = Chironomus tentans 

LV = Lumbriculus vafiegalus 

6.3.6.2 All test chambers and equipment should be solution can be used in place of both the organic solvent 

thoroughly rinsed with the dilution water immediately and the acid (see ASTM, 1988a), but the solution might 

before use in a test. attack silicone adhesive and leave chromium residues 

on glass. A alternative to use of dichromate-sulfuric acid 

6.3.6.3 Many organic solvents leave a film that is could be to heat glassware for 8 h at 450%. 

insoluble in water. A dichromate-sulfuric acid cleaning 

21 
Section 7 

Water, Formulated Sediment, Reagents, and Standards 

7.1 Water 

7.1.1 Requirements 

7.1.1.1 Water used to test and culture organisms should 

be uniform in quality. Acceptable water should allow 

satisfactory survival, growth, or reproduction of the test 

organisms. Test organisms should not show signs of 

disease or apparent stress (e.g., discoloration, unusual 

behavior). If problems are observed in the culturing or 

testing of organisms, it is desirable to evaluate the 

characteristics of the water. See USEPA (1993a) and 

ASTM (1994a) for a recommended list of chemical 

analyses of the water supply. 

7.1.2 Source 

7.1.2.1 A natural water is considered to be of uniform 

quality if monthly ranges of the hardness, alkalinity, and 

specific conductance are less than 10% of their respective averages and if the monthly range of pH is less than 

0.4. Natural waters should be obtained from an uncontaminated well or spring, if possible, or from a 

surface-water source. If surface water is used, the intake should be positioned to (1) minimize fluctuations in 

quality and contamination, (2) maximize the concentration of dissolved oxygen, and (3) ensure low concentrations of sulfide and iron. Municipal-water supplies may 

be variable and may contain unacceptably high concentrations of materials such as copper, lead, zinc, fluoride, 

chlorine, or chloramines. Chlorinated water should not 

be used for culturing or testing because residual chlorine and chlorine-produced oxidants are toxic to many 

aquatic organisms. Use of tap water is discouraged 

unless it is dechlorinated and passed through a deionizer 

and carbon filter (USEPA, 1993a). 

7.1.2.2 For site-specific investigations, it is desirable to 

have the waterquality characteristics of the overlying 

water as similar as possible to the site water. For certain 

applications the experimental design might require use 

of water from the site where sediment is collected. 

7.1.2.3 Water that might be contaminated with facultative pathogens may be passed through a properly maintained ultraviolet sterilizer equipped with an intensity 

meter and flow controls or passed through a filter with a 

pore size of 0.45 μm or less. 

7.1.2.4 Water might need aeration using air stones, 

surface aerators, or column aerators. Adequate aeration 

will stabilize pH, bring concentrations of dissolved oxygen and other gases into equilibrium with air, and minimize oxygen demand and concentrations of volatiles. 

Excessive aeration may reduce hardness and alkalinity 

of hard water (e.g., 280 mg/L hardness as CaCO3; E.L. 

Brunson, NBS, Columbia, MO, personal communication). The concentration of dissolved oxygen in source 

water should be between 90 to 100% saturation to help 

ensure that dissolved oxygen concentrations are acceptable in test chambers. 

7.1.3 Reconstituted Water 

7.1.3.1 Ideally, reconstituted water should be prepared 

by adding specified amounts of reagent-grade chemicals to high-purity distilled or deionized water (ASTM, 

1988a; USEPA, 1993a). In some applications, acceptable high-purity water can be prepared using deionization, distillation, or reverse-osmosis units (Section 6.3.3.2; 

USEPA, 1993a). In some applications, test water can be 

prepared by diluting natural water with deionized water 

(Kemble et al., 1993). 

7.1.3.2 Deionized water should be obtained from a 

system capable of producing at least 1 mega-ohm water. If large quantities of high quality deionized water are 

needed, it may be advisable to supply the laboratory 

grade water deionizer with preconditioned water from a 

mixed-bed water treatment system. 

7.1.3.3 Conductivity, pH, hardness, dissolved oxygen, 

and alkalinity should be measured on each batch of 

reconstituted water. The reconstituted water should be 

aerated before use to adjust pH and dissolved oxygen to 

the acceptable ranges (e.g., Section 7.1.3.4.1). USEPA 

(1993a) recommends using a batch of reconstituted 

water for two weeks. 

7.1.3.4 Reconstituted Fresh Water 

7.1.3.4.1 To prepare 100 L of reconstituted fresh water, 

use the reagent grade chemicals as follows: 

1. Place about 75 L of deionized water in a properly 

cleaned container. 

22 
2. Add 5 g of CaSO4 and 5 g of CaCI2 to a 2-L aliquot of 

deionized water and mix (e.g., on a stir plate) for 30 

min or until the salts dissolve. 

3. Add 3 g of MgSO4, 9.6 g NaHCO3, and 0.4 g KCI to 

a second 2-L aliquot of deionized water and mix on 

a stir plate for 30 min. 

4. Pour the two 2-L aliquots containing the dissolved 

salts into the 75 L of deionized water and fill the 

carboy to 100 L with deionized water. 

5. Aerate the mixture for at least 24 h before use. 

6. The water quality of the reconstituted water should 

be approximately the following: hardness, 90 to 100 

mg/L as CaCO3, alkalinity 50 to 70 mg/L as CaCO3, 

conductivity 330 to 360 μS/cm, and pH 7.8 to 8.2. 

7.1.3.4.2 This reconstituted fresh water was developed 

by USEPA EMSL-Cincinnati (J.M. Lazorchak, USEPA, 

Cincinnati, OH, personal communication) and has been 

used successfully in round-robin testing with H. azteca, 

C. tentans, and C. riparius (Section 15). This reconstituted water has a higher proportion of chloride to sulfate 

compared to the reconstituted waters described in ASTM 

(1988a) and USEPA (1993a). Variable success has 

been reported using USEPA or ASTM reconstituted 

waters (USEPA, 1993a) with H. azteca. Research is 

ongoing to develop additional types of reconstituted 

waters suitable for these test organisms. 

7.1.3.5 Synthetic Seawater 

7.1.3.5.1 Reconstituted salt water can be prepared by 

adding commercial sea salts, such as FORTY FATHOMS®, HW MARINEMIX®, INSTANT OCEAN@, or 

equivalent to deionized water. 

7.1.3.5.2 A synthetic seawater formulation called GP2 

is prepared with reagent grade chemicals that can be 

diluted with deionized water to the desired salinity 

(USEPA, 1994c). 

7.1.3.5.3 Ingersoll et al. (1992) describe procedures for 

culturing H. azteca at salinities up to 15%. Reconstituted salt water was prepared by adding INSTANT 

OCEAN@ salts to a 25:75 (v/v) mixture of freshwater 

(hardness 283 mg/L as CaCO3) and deionized water 

that was held at least two weeks before use. Synthetic 

seawater was conditioned by adding 6.2 mL of Frit-zyme® 

#9 nitrifying bacteria (Nitromonas sp. and Nitrobacter 

sp.; Fritz Chemical Company, Dallas, TX) to each liter of 

water. The cultures were maintained by using static 

renewal procedures; 25% of the culture water was replaced weekly. Hyalella azteca have been used to evaluate the toxicity of estuarine sediments up to 15 % 

salinity (Nebeker and Miller, 1988; Roach et al., 1992; 

Winger et al., 1993). 

7.2 Formulated Sediment 

7.2.1 General Requirements 

7.2.1.1 Formulated sediments are mixtures of materials 

that mimic natural sediments. Formulated sediments 

have not been routinely applied to evaluate sediment 

contamination. A primary use of formulated sediment 

could be as a control sediment. Formulated sediments 

allow for standardization of sediment testing or as a 

basis for conducting sediment research. Formulated 

sediment provides a basis by which any testing program 

can assess the acceptability of their procedures and 

facilities. In addition, formulated sediment provides a 

consistent measure evaluating performance-based criteria necessary for test acceptability. The use of formulated sediment eliminates interferences caused by the 

presence of indigenous organisms. For toxicity tests 

with sediments spiked with specific chemicals, the use 

of a formulated sediment eliminates or controls the 

variation in sediment physico-chemical characteristics 

and provides a consistent method for evaluating the fate 

of chemicals in sediment. However, additional research 

is needed before formulated sediments are used routinely for sediment spiking procedures (e.g.. identifying 

standardized and representative sources of organic carbon). 

7.2.1.2 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) the materials used in formulation of the sediment should have 

consistent characteristics. Consistent material characteristics include (1) consistency of materials from batch 

to batch, (2) contaminant concentrations below concentrations of concern, and (3) availability to all individuals 

and facilities. 

7.2.1.3 Physico-chemical characteristics that might be 

considered when evaluating the appropriateness of a 

formulated sediment include percent sand, percent clay, 

percent silt, organic carbon content, cation exchange 

capacity (CEC), oxidation reduction potential (redox), 

pH, and carbon:nitrogen:phosphorus ratios. 

7.2.2 Sources of Materials 

7.2.2.1 A variety of methods describe procedures for 

making formulated sediments. These procedures often 

use similar constituents; however, they often include 

either a component or a formulation step that would 

result in variation from test facility to test facility. In 

addition, none of the procedures have been subjected to 

standardization and consensus approval or round-robin 

(ring) testing. 

7.2.2.2 Most formulated sediments include sand and 

clay/silt that meet certain specifications; however, some 

may be quite different. For example, three sources of 

clay and silt include Attagel® 50, ASP® 400, and ASP® 

23 
400P. Table 7.1 summarizes the characteristics of these 

materials. The percentage of clay ranges from 56.5 to 

88.5 and silt ranges from 11.5 to 43.5. These characteristics should be evaluated when considering the materials to use in a formulated sediment. 

Table 7.1 Characteristics of Three Sources of Clays and Sifts 

Used in Formulated Sediments 

Charactenstic Attage!@ 50 ASP@ 400 ASF+B 4OOP 

% Sand 

% Clay 

% Slk 

Soil class 

0.0 

88.50 

11.50 

Clay 

0.01 

68.49 

31.50 

Clay 

0.0 

56.50 

43.50 

Silty clay 

Note: Table 7.3 IS list of suppliers. 

7.2.2.3 A critical component of formulated sediment is 

the source of organic carbon. Many procedures have 

used peat as the source of organic carbon. Other sources 

of organic carbon listed in Table 7.2 have been evaluated including humus, potting soil, maple leaves, 

composted cow manure, rabbit chow, cereal leaves, 

chlorella, trout chow, Tetramin@ and Tetrafin@? Only 

peat, humus, potting soil, and composted cow manure 

have been used successfully without fouling the overlying water. The other sources of organic carbon (Table 

7.2) cause dissolved oxygen concentrations to fall to 

unacceptable levels (F.J. Dwyer, NBS, Columbia, MO, 

personal communication). If appropriate conditioning pr@ 

cedures can be determined these other sources of 

organic carbon may be acceptable. An important consideration in the selection of an organic carbon source is the 

ratio of carbon:nitrogen:phosphorus. As demonstrated in 

Table 7.2, percentage carbon ranged from 30 to 47, 

nitrogen ranged from 3 to 45 mg/g, and phosphorus 

ranged from below detection to 11 Mg for several 

different carbon sources. These characteristics should 

be evaluated when considering the materials to use in a 

formulated sediment. 

Table 7.2. Carbon, Nitrogen, Phosphorus Levels for Various 

Sources of Organic Carbon’ 

Organic carbon Carbon Nitrogen Phosphorus 

source WI OwQ) Wg) 

Peat 

Maple leaves 1 

Maple leaves 2 

Cow manure 

Rabbit chow 

Humic acid 

Cereal leaves 

Chlorella 

Trout chow 

Tetramin@ 

47 4 0.4 

42 6 1.3 

47 3 1.7 

30 11 8.2 

40 18 0.2 

40 3 - 

47 4 0.4 

40 41 5.7 

43 36 11 

37 45 9.6 

36 29 8.6 

7.2.3 Procedure 

7.2.3.1 A summary of procedures that have been used 

to formulate sediment are listed below. Suppliers of 

various components are listed in Table 7.3. 

1. 

2. 

3. 

4. 

Walsh et al. (1981): (1) Wash sand (Mystic White No 

85, 45, and 18-New England Silica Inc) and sieve 

into three grain sizes: coarse (500 to 1500 pm); 

medium (250 to 499 pm); and fine (63 to 249 pm). 

(2) Clay and silt were obtained from Engelhard 

Corp.; (3) Peat moss is milled and sieved through an 

840 pm screen. (4) Constituents are mixed dry in 

the following quantities: coarse sand (0.6%); medium sand (8.7%); fine sand (69.2%); silt (10.2%): 

clay (6.4%); and organic matter (4.9%). 

Clements, W.H. (Colorado State University, Ft. 

Collins, CO, personal communication): (1) Rinse 

peat moss then soak for 5 d in deionized water 

renewing water daily. (2) After acclimation for 5 d 

remove all water and spread out to dry. (3) Grind 

moss and sieve using the following sieve sizes: 1 .18 

mm (discard these particles); 1.00 mm {average 

size 1.09 mm); 0.85 mm (average size 0.925); 0.60 

(average size 0.725); 0.425 mm (average size 0.5125 

mm); retainer (average size 0.2125 mm). (4) Use a 

mixture of sizes that provides an average particle 

size of 840 m. (5) Wash sand (Mystic white #45) 

and dry. (6) Clay and silt are obtained using ASP 

400 (Englehard Corp). (7) ConstituerBs are mixed 

dry in the following quantities: sand (1242 g); silt and 

clay (219 g); dolomite (7.5 g); peat moss (31.5 g); 

and humic acid (0.15 g). (8) Sediment is mixed for 

an hour on a rolling mill and stored dry until ready for 

use. 

Hanes et al. (1991): (1) Sieve sand and retain two 

particle sizes (90 to 180 urn and 180 to 250 urn) 

which are mixed in a ratio of 2:l. (2) Potting soil is 

dried for 24 h at room temperature and sieved 

through a 1 -mm screen. Clay is commercially available sculptors clay. (3) Determine percent moisture 

of clay and soil after drying for 24 h at 60 to 100°C. 

(correct for percent moisture when mixing materials). (4) Constituents are mixed by weight in the 

following ratios: sand mixture (42%); clay (42%); 

and soil (16%). (5) After mixing, autoclave in a foilcovered container for 20 min. (6) Mixture can be 

stored indefinitely if kept covered after autoclaving. 

Naylor (1993): (1) Sand is acid-washed and sieved 

to obtain a 40 to 100 mm size. (2) Clay is kaolin light. 

(3) Peat moss is ground and sieved using a 2-mm 

screen (peat moss which is allowed to dry out will 

not rehydrate and will float on the water surface). (4) 

Adjust for the use of moist peat moss by determining 

moisture content (dry 5 samples of peat at 60°C until 

constant weight is achieved). (5) Constituents are 

mixed by weight in the following percentages: sand 

(69%); kaolin (20%); peat (10% [adjust for moisture 

1 F.J. Dwyer, NBS. Columbia, MO, personal communication 

24 
Table 7.3 Sources of Components Used in Formulated Sediments 

Component Sources 

Sand 

Kaolln!te 

Montmorillonite 

Clay 

Humus 

Peat 

Potttng so11 

Humic acid 

Cow manure 

Dolomite 

. Mystic White #18, #45. #65. #X+-New England Silica. Inc.. South Windsor, CT 

* Product No. 33094, BDH Chemical, Ltd.. Poole, England 

* ASP 400, ASP 4OOP, ASP 600, ASP 900-Englehard Corporation, Edison, NJ 

. Product No. 33059, BDH Chemical, Ltd., Poole, England 

. W.D. Johns, Source Clays, University of Missouri. Columbia. MO 

* Lewiscratt Sculptor’s Clay, available In hobby and artist supply stores 

. Sims Bark Co., Inc., Tuscumbia, AL 

. D.L. Brownmg Co., Mather, WI 

* Joseph Bentley. Ltd., Barrow-on-Humber, South Humberside. England 

* Melllnger’s, North Lima, OH 

. Zehr’s No Name Potting SolI, Mlssssauga, Ontario 

. Aldrich Chemical Co, Milwaukee, WI 

. A.H. Hoffman, Inc., LandiswIle, PA 

- Ward’s Natural Science Establishment, Inc., Rochester, NY 

- Southern Agri-mrnerals Corp., Hartford, AL 

5. 

content]); and CaCO, (1%). (6) Mix for 2 h in a soil 

shaker and store in sealed containers. 

Suede1 and Rodgers (1994): (1) Sand (Mystic White 

#18 and 90) is sieved to provide three different size 

fractions: coarse (2.0 to 0.5 mm), medium (0.5 to 

0.25 mm) and fine (0.25 to 0.05 mm). (2) Silt (ASP 

400), clay (ASP 600 and 900), montmorillonite clay, 

and dolomite are ashed at 550°C. for 1 h to remove 

organic matter. (3) Humus is dried (70°C) and milled 

to 2.0 mm. (4) Dolomite is added as 1% of the silt 

requirement. (5) Materials are aged for 7 d in flowing 

water before mixing. (6) Constituents are mixed to 

mimic the desired characteristics of the sediment of 

concern. 

7.2.3.2 The procedure for formulating a sediment is a 

combination of methods outlined in Section 7.2.3.1. The 

characteristics of this formulation would be sand 77%, 

silt/clay 17%. The organic matter would depend on the 

source of organic carbon. This approach could be modified to mimic specific characteristics of a sediment. If a 

formulated sediment is to be used as a control sediment, 

the physico-chemical characteristics of the formulated 

sediment should be within the tolerance limits of the test 

organism. 

1. Wash sand, sieve, and retain the following two size 

groups: medium (0.5 to 0.25 mm) and fine (0.25 to 

0.05 mm). Sand should be mixed at a ratio of 2:1, 

fine:medium. 

2. Clay and silt fractions are obtained using ASP@ 

400. Other clays or silts (e.g., Attagel@ 50, ASP@ 

4OOP, ASP@ 600, ASP@ 900, montmorillonite) might 

be used if specific characteristics are required. 

3. Organic matter (peat, humus, cow manure) should 

be dried, milled, and passed through a 0.64 mm 

sieve. 

4. Either CaMg(COJ, or CaCO, should be added to 

buffer the sediments. 

5. Att constituents are mixed on a percent dry weight 

basis. Mix in the following ratios: sand (77%); silt/ 

clay (17%); organic matter (5%); buffer (1%). 

7.3 Reagents 

7.3.1 Data sheets should be followed for reagents and 

other chemicals purchased from supply houses. The 

test material(s) should be at least reagent grade, unless 

a test on formulation commercial product, technical-grade, 

or use-grade material is specifically needed. Reagent 

containers should be dated when received from the 

supplier, and the shelf life of the reagent should not be 

exceeded. Working solutions should be dated when 

prepared and the recommended shelf life should not be 

exceeded. 

7.4 Standards 

7.4.1 Appropriate standard methods for chemical and 

physical analyses should be used when possible. For 

those measurements for which standards do not exist or 

are not sensitive enough, methods should be obtained 

from other reliable sources. 

25 
Section 8 

Sample Collection, Storage, Manipulation, and Characterization 

8.1 Collection 

8.1.1 Before the preparation or collection of sediment, a 

procedure should be established for the handling of 

sediments that might contain unknown quantities of 

toxic contaminants (Section 5). 

8.1.2 Sediments are spatially and temporally variable 

(Stemmer et al., 1990a). Replicate samples should be 

collected to determine variance in sediment characteristics. Sediment should be collected with as little disruption as possible; however, subsampling, cornpositing, or 

homogenization of sediment samples may be necessary 

for some experimental designs. Sampling may cause 

toss of sediment integrity, change in chemical speciation, or disruption of chemical equilibrium (ASTM, 1994b). 

A benthic grab or core should be used rather than a 

dredge to minimize disruption of the sediment sample. 

Sediment should be collected from a depth that will 

represent expected exposure. For example, oligochaetes may burrow 4 to 15 cm into sediment. Samples 

collected for evaluations of dredged material should 

include all sediment to project depth. Surveys of the 

toxicity of surficial sediment are often based on cores of 

the upper 2 cm sediment depth. 

8.1.3 Exposure to direct sunlight during collection should 

be minimized, especially if the sediment contains photolytic compounds. Sediment samples should be cooled 

to 4°C in the field before shipment (ASTM, 1994a). Dry 

ice can be used to cool samples in the field; however, 

sediments should never be frozen. Monitors can be 

used to measure temperature during shipping (e.g., 

TempTale Temperature Monitoring and Recording System, Sensitech, Inc., Beverly, MA). 

8.1.4 For additional information on sediment collection 

and shipment see ASTM (1994b). 

8.2 Storage 

8.2.1 Manipulation or storage can alter bioavailability of 

contaminants in sediment (Burton and Ingersoll, 1994); 

however, the alterations that occur may not substantially 

affect toxicity. Storage of sediment samples for several 

months at 4°C did not result in significant changes in 

chemistry or toxicity (T. Dillon and H. Tatem, USCOE, 

Vicksburg, MS, personal communication; G.T. Ankley 

and D. DeFoe, USEPA, Duluth, MN, unpublished data); 

however, others have demonstrated changes in spiked 

sediment within days to weeks (e.g., Burton, 1991; 

Stemmer et al., 1990a). Sediments primarily contaminated with nonionic, nonvolatile organics will probably 

change little during storage because of their relative 

resistance to biodegradation and sorption to solids. However, metals and metalloids may be affected by changing redox, oxidation, or microbial metabolism (such as 

with arsenic, selenium, mercury, lead, and tin; all of 

which are methylated by a number of bacteria and 

fungi). Metal-contaminated sediments may need to be 

tested relatively soon after collection with as little manipulation as possible (Burton and Ingersoll, 1994). 

8.2.2 Given that the contaminants of concern and the 

influencing sediment characteristics are not always known 

a priori, it is desirable to hold sediments in the dark at 

4°C and start tests soon after collection from the field. 

Recommended sediment holding time ranges from less 

than two (ASTM, 1994a) to less than eight weeks 

(USEPA-USCOE, 1994). If whole-sediment tests are 

started after two weeks of collection, it may be desirable 

to conduct additional characterizations of sediment to 

evaluate possible effects of storage on sediment. For 

example, concentrations of contaminants of concern 

could be measured in pore water within two weeks from 

sediment collection and at the start of the sediment test 

(Kemble et al., 1993). Ingersoll et al. (1993) recommend 

conducting a toxicity test with pore water within two 

weeks from sediment collection and at the start of the 

sediment test. Freezing and longer storage might further 

change sediment properties such as grain size or contaminant partitioning and should be avoided (ASTM, 

1994b; Schuytema et al., 1989; K.E. Day Environment 

Canada, Burlington, Ontario, personal communication). 

Sediment should be stored with no air over the sealed 

samples (no head space) at 4°C before the start of a test 

(Shuba et al., 1978; ASTM, 1994b). Sediment may be 

stored in containers constructed of suitable materials as 

outlined in Section 6. It is desirable to avoid contact with 

metals, including stainless steel and brass sieving 

screens, and some plastics. 

26 
8.3 Manipulation 

8.3.1 Homogenization 

8.3.1.1 Samples tend to settle during shipment. As a 

result, water above the sediment should not be discarded but should be mixed back into the sediment 

during homogenization. Sediment samples should not 

be sieved to remove indigenous organisms unless there 

is a good reason to believe indigenous organisms may 

influence the response of the test organism. However, 

large indigenous organisms and large debris can be 

removed using forceps. Reynoldson et al. (1994) observed reduced growth of amphipods, midges, and mayflies in sediments with elevated numbers of oligochaetes 

and recommended sieving sediments suspected to have 

high numbers of indigenous oligochaetes. If sediments 

must be sieved, it may be desirable to analyze samples 

before and after sieving (e.g., pore-water metals, DOC, 

AVS, TOC) to document the influence of sieving on 

sediment chemistry. 

8.3.1.2 If sediment is collected from multiple field 

samples, the sediment can be pooled and mixed using 

stirring or a rolling mill, feed mixer, or other suitable 

apparatus (see ASTM, 1994b). Homogenization of sediment can be accomplished using a modified 30-cm 

bench-top drill press (Dayton Model 32993) or a 

variable-speed hand-held drill outfitted with a stainlesssteel auger (diameter 7.6 cm, overall length 38 cm, 

auger bit length 25.4 cm; Part No. 800707, Augers 

Unlimited, Extort, PA; Kemble et al., 1994). These procedures could also be used to mix test sediment with a 

control sediment in dilution experiments. 

8.3.2 Sediment Spiking 

8.3.2.1 Test sediment can be prepared by manipulating 

the properties of a control sediment. Additional research 

is needed before formulated sediments are used routinely for sediment spiking procedures (e.g., identifying 

standardized and representative sources of organic carbon). Mixing time (Stemmer et al., 1990a) and aging 

(Word et al., 1987; Landrum, 1989; Landrum and Faust, 

1992) of spiked sediment can affect responses. Many 

studies with spiked sediment are often started only a few 

days after the chemical has been added to the sediment. This short time period may not be long enough for 

sediments to equilibrate with the spiked chemicals. Consistent spiking procedures should be followed in order to 

make interlaboratory comparisons. It is recommended 

that spiked sediment be aged at least one month before 

starting a test: however equilibration for some chemicals 

may not be achieved for long periods of time. 

8.3.2.1.1 The cause of sediment toxicity and the magnitude of interactive effects of contaminants can be estimated by spiking a sediment with chemicals or complex 

waste mixtures (Lamberson and Swartz, 1992). Sediments spiked with a range of concentrations can be 

used to generate either point estimates (e.g., LC50) or a 

minimum concentration at which effects are observed 

(lowest-observable-effect concentration; LOEC). Results 

of tests may be reported in terms of a BSAF (Ankley et 

al., 1992b). The influence of sediment physico-chemical 

characteristics on chemical toxicity can also be determined with sediment-spiking studies (Adams et al., 1985). 

8.3.2.2 The test material(s) should be at least reagent 

grade, unless a test on formulation commercial product, 

technical-grade, or use-grade material is specifically 

needed. Before a test is started, the following should be 

known about the test material: (1) the identity and concentration of major ingredients and impurities, (2) water 

solubility in test water, (3) estimated toxicity to the test 

organism and to humans, (4) if the test concentration(s) 

are to be measured, the precision and bias of the 

analytical method at the planned concentration(s) of the 

test material, and (5) recommended handling and disposal procedures. 

8.3.2.2.1 Organic compounds have been added in the 

dry form or coated on the inside walls of the container 

(Ditsworth et al., 1990). Metals are generally added in 

an aqueous solution (ASTM, 1994b; Carlson et al., 

1991; Di Toro et al., 1990). If an organic solvent is used, 

the solvent in the sediment should be at a concentration 

that does not affect the test organism. Concentrations of 

the chemical in the pore water and in the whole sediment should be monitored at the beginning and the end 

of a test. 

8.3.2.3 Use of a solvent other than water should be 

avoided if possible. Addition of organic solvents may 

dramatically influence the concentration of dissolved 

organic carbon in pore water (G.T. Ankley, USEPA, 

Duluth, MN, personal communication). If an organic 

solvent must be used, both a solvent-control and a 

negative-control sediment must be included in a test. 

The solvent control must contain the highest concentration of solvent present and must be from the same batch 

used to make the stock solution (see ASTM, 1988a). 

The same concentration of solvent should be used in all 

treatments. If an organic solvent is used as a carrier, it 

may be possible to perform successive washes of sediment to remove most of the solvent while leaving the 

compound of study (Harkey et al., 1994). 

8.3.2.4 If the concentration of solvent is not the same in 

all test solutions that contain test material, a solvent test 

should be conducted to determine whether survival, 

growth, or reproduction of the test organisms is related 

to the concentration of the solvent. 

8.3.2.4.1 If the test contains both a negative control and 

a solvent control, the survival, growth, or reproduction of 

the organisms tested should be compared. If a statistically significant difference is detected between the two 

controls, only the solvent control may be used for meeting the acceptability of the test and as the basis for 

calculating results. The negative control might provide 

additional information on the general health of the organisms tested. If no statistically significant difference is 

detected, the data from both controls should be used for 

27 
meeting the acceptability of the test and as the basis for 

calculating the results (ASTM, 1992). 

8.3.2.5 Test Concentration(s) for Laboratory 

Spiked Sediments 

8.3.2.5.1 If a test is intended to generate an LC50, the 

selected test concentrations should bracket the predicted LC50. The prediction might be based on the 

results of a test on the same or a similar test material 

with the same or a similar test organism. The LC50 of a 

particular compound may vary depending on physical 

and chemical sediment characteristics. If a useful prediction is not available, it is desirable to conduct a 

range-finding test in which the organisms are exposed 

to a control and three or more concentrations of the test 

material that differ by a factor of ten. Results from 

water-only tests could be used to establish concentrations to be tested in a whole-sediment test based on 

predicted pore-water concentrations (Di Toro et al., 1991). 

8.3.2.5.2 Bulk-sediment chemical concentrations might 

be normalized to factors other than dry weight. For 

example, concentrations of nonpolar organic compounds 

might be normalized to sediment organic-carbon content, and simultaneously extracted metals might be normalized to acid volatile sulfides (Di Toro et at, 1990; Di 

Toro et al., 199 1). 

8.3.2.5.3 In some situations it might be necessary to 

only determine whether a specific concentration of test 

material is toxic to the test organism, or whether adverse 

effects occur above or below a specific concentration. 

When there is interest in a particular concentration, it 

might only be necessary to test that concentration and 

not to determine an LC50. 

8.3.2.6 Addition of test material(s) to sediment may be 

accomplished using various methods, such as a (1) 

rolling mill, (2) feed mixer, or (3) hand mixing (ASTM, 

1994b). Modifications of the mixing techniques might be 

necessary to allow time for a test material to equilibrate 

with the sediment. Mixing time of spiked sediment should 

be limited from minutes to a few hours, and temperature 

should be kept low to minimize potential changes in the 

physico-chemical and microbial characteristics of the 

sediment (ASTM, 1994b). Duration of contact between 

the chemical and sediment can affect partitioning and 

bioavailability (Word et al., 1987). Care should be taken 

to ensure that the chemical is thoroughly and evenly 

distributed in the sediment. Analyses of sediment subsamples are advisable to determine the degree of mixing homogeneity (Ditsworth et al., 1990). Moreover, 

results from sediment-spiking studies should be compared to the response of test organisms to chemical 

concentrations in natural sediments (Lamberson and 

Swartz, 1992). 

8.4 Characterization 

8.4.1 All sediments should be characterized and at least 

the following determined: pH and ammonia of the pore 

water, organic carbon content (total organic carbon, 

TOC), particle size distribution (percent sand, silt, clay), 

and percent water content (ASTM, 1994a; Plumb, 1981). 

8.4.2 Other analyses on sediments might include biological oxygen demand, chemical oxygen demand, cation exchange capacity, Eh, total inorganic carbon, total 

volatile solids, acid volatile sulfides, metals, synthetic 

organic compounds, oil and grease, petroleum hydrocarbons, as well as interstitial water analyses for various 

physico-chemical parameters. 

8.4.3 Macrobenthos may be evaluated by subsampling 

the field-collected sediment. If direct comparisons are to 

be made, subsamples for toxicity testing should be 

collected from the same sample for analysis of sediment 

physical and chemical characterizations. Qualitative descriptions of the sediment may include color, texture, 

presence of macrophytes or animals. Monitoring the 

odor of sediment samples should be avoided because of 

potential hazardous volatile contaminants. 

8.4.4 Analytical Methodology 

8.4.4.1 Chemical and physical data should be obtained 

using appropriate standard methods whenever possible. 

For those measurements for which standard methods 

do not exist or are not sensitive enough, methods should 

be obtained from other reliable sources. 

8.4.4.2 The precision, accuracy, and bias of each 

analytical method used should be determined in the 

appropriate matrix: that is, sediment, water, tissue. Reagent blanks and analytical standards should be analyzed, and recoveries should be calculated. 

8.4.4.3 Concentration of spiked test material(s) in sediment, interstitial water, and overlying water should be 

measured as often as practical during a test. If possible, 

the concentration of the test material in overlying water, 

interstitial water and sediments should be measured at 

the start and end of a test. Measurement of test 

material(s) degradation products might also be desirable. 

8.4.4.4 Separate chambers should be set up at the start 

of a test and destructively sampled during and at the end 

of the test to monitor sediment chemistry. Test organisms might be added to these extra chambers depending on the objective of the study. 

8.4.4.5 Measurement of test material(s) concentration 

in water can be accomplished by pipeting water samples 

from about 1 to 2 cm above the sediment surface in the 

test chamber. Overlying water samples should not con28 
tain any surface debris, any material from the sides of 8.4.4.7 A variety of procedures have been used to 

the test chamber, or any sediment. isolate interstitial water including centrifugation, filtration, pressure, or by using an interstitial water sampler; 

8.4.4.6 Measurement of test material(s) concentration however, centrifugation without filtration is the recomin sediment at the end of a test can be taken by mended procedure (Ankley and Schubauer-Berigan, 

siphoning most of the overlying water without disturbing 1994). filtration may reduce concentrations of materials 

the surface of the sediment, then removing appropriate in interstitial water (Schults et al., 1992). Care should be 

aliquots of the sediment for chemical analysis. taken to ensure that contaminants do not transform, 

degrade, or volatilize during isolation or storage of the 

interstitial water sample. - 

29 
Section 9 

Quality Assurance and Quality Control 

9.1 Introduction 

9.1.1 Developing and maintaining a laboratory quality 

assurance (QA) program requires an ongoing commitment by laboratory management and also includes the 

following: (1) appointment of a laboratory 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 laboratory Standard Operating Procedures (SOPS) for test 

organism culturing, testing, 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. Additional guidance for QA can be obtained in 

USEPA (1989d). 

9.1.2 QA practices within a testing laboratory should 

address all activities that affect the quality of the final 

data, such as (1) sediment sampling and handling, (2) 

the source and condition of the test organisms, (3) 

condition and operation of equipment, (4) test conditions, (5) instrument calibration, (6) replication, (7) use 

of reference toxicants, (8) record keeping, and (9) data 

evaluation. 

9.1.3 Quality control (QC) practices, on the other hand, 

consist of the more focused, routine, day-to-day activities carried out within the scope of the overall QA 

program. For more detailed discussion of quality assurance, and general guidance on good laboratory practices related to testing see FDA (1978), USEPA (1979a), 

USEPA (1980a), USEPA (1980b), USEPA (1993a), 

USEPA (1994b), USEPA (1994c), DeWoskin (1984), 

and Taylor (1987). 

9.2 Performance-based Criteria 

9.2.1 USEPA Environmental Monitoring Management 

Council (EMMC) recommended the use of 

performance-based methods in developing standards 

for chemical analytical methods (Williams, 1993). 

Performance-based methods were defined by EMMC 

as a monitoring approach that permits the use of appropriate methods that meet pre-established demonstrated 

performance standards. Minimum required elements of 

performance, such as precision, reproducibility, bias, 

sensitivity, and detection limits should be specified, and 

the method should be demonstrated to meet the performance standards. 

9.2.2 Participants at a September 1992 USEPA sediment toxicity workshop arrived at a consensus on several culturing and testing methods for freshwater organisms (Appendix A, Section S.4). In developing guidance 

for culturing test organisms to be included in this manual 

for sediment tests, it was generally agreed that no single 

method must be used to culture organisms. Success of 

a test relies on the health of the culture from which 

organisms are taken for testing. Having healthy organisms of known quality and age for testing is the key 

consideration relative to culture methods. Therefore, a 

performance-based criteria approach is the preferred 

method through which individual laboratories should 

evaluate culture health rather than using control-based 

criteria. Performance-based criteria were chosen to allow each laboratory to optimize culture methods while 

providing organisms that produce reliable and comparable test results. See Tables 11.3, 12.3, and 13.4 for a 

listing of performance criteria for culturing and testing. 

9.3 Facilities, Equipment, and Test 

Chambers 

9.3.1 Separate areas for test organism culturing and 

testing must be provided to avoid loss of cultures due to 

cross-contamination. Ventilation systems should be designed and operated to prevent recirculation or leakage 

of air from chemical analysis laboratories or sample 

storage and preparation areas into test organism culturing or sediment testing areas, and from sediment testing 

laboratories and sample preparation areas into culture 

rooms. 

9.3.2 Equipment for temperature control should be 

adequate to maintain recommended test-water temperatures. Recommended materials should be used in 

the fabricating of the test equipment that comes in 

contact with the sediment or overlying water. 

9.3.3 Before a sediment test is conducted in a new 

facility, a “non-contaminant” test should be conducted in 

which all test chambers contain a control sediment and 

30 
overlying water. This information is used to demonstrate 

that the facility, control sediment, water, and handling 

procedures provide acceptable responses of test organisms (Section 9.14). 

9.4 Test Organisms 

9.4.1 The organisms should appear healthy, behave 

normally, feed well, and have low mortality in cultures, 

during holding (e.g., <20% for 48 h before the start of a 

test), and in test controls. The species of test organisms 

should be positively identified to species. 

9.5 Water 

9.5.1 The quality of water used for organism culturing 

and testing is extremely important. Overlying water used 

in testing and water used in culturing organisms should 

be uniform in quality. Acceptable water should allow 

satisfactory survival, growth, or reproduction of the test 

organisms. Test organisms should not show signs of 

disease or apparent stress (e.g., discoloration, unusual 

behavior). See Section 7 for additional details. 

9.6 Sample Collection and Storage 

9.6.1 Sample holding times and temperatures should 

conform to conditions described in Section 8. 

9.7 Test Conditions 

9.7.1 It is desirable to measure temperature continuously in at least one chamber during each test. Temperatures should be maintained within the limits specified for each test. Dissolved oxygen, alkalinity, water 

hardness, conductivity, ammonia, and pH should be 

checked as prescribed in Sections 11.3, 12.3, and 13.3. 

9.8 Quality of Test Organisms 

9.8.1 Monthly reference-toxicity tests should be conducted on all test organisms using procedures outlined 

in Section 9.16. If reference-toxicity tests are not conducted monthly, the lot of organisms used to start a 

sediment test must be evaluated using a reference 

toxicant. Physiological measurements such as lipid content might also provide useful information regarding the 

health of the cultures. 

9.8.2 The quality of test organisms obtained from an 

outside source must be verified by conducting a 

reference-toxicity test concurrently with the sediment 

test. The supplier should provide data with the shipment 

describing the history of the sensitivity of organisms 

from the same source culture. If the supplier has not 

conducted five reference toxicity tests with the test 

organism, it is the responsibility of the testing laboratory 

to conduct these five reference toxicity tests before 

starting a sediment test (Section 9.14.1). 

9.8.3 The supplier should also certify the species identification of the test organisms and provide the taxonomic references or name(s) of the taxonomic expert(s) 

consulted. 

9.9 Quality of Food 

9.9.1 Problems with the nutritional suitability of the food 

will be reflected in the survival, growth, or reproduction 

of the test organisms in cultures or in sediment tests. 

9.9.2 Food used to culture organisms used in bioaccumulation tests must be analyzed for compounds to be 

measured in the bioaccumulation tests. 

9.10 Test Acceptability 

9.10.1 For the test results to be acceptable, survival at 

10 d must equal or exceed 80% for H. azteca and 70% 

for C. tentans in the control sediment. Numbers of L. 

variegatus should not be reduced in test sediments 

relative to the control sediment and organisms should 

burrow into the test sediment. Avoidance of test sediment by L. variegatus will decrease bioaccumulation. 

See Table 11.3, 12.3, and 13.4 for additional requirements for acceptability of the tests. 

9.10.2 An individual test may be conditionally acceptable if temperature, dissolved oxygen, and other specified conditions fall outside specifications, depending on 

the degree of the departure and the objectives of the 

tests (see test condition summaries). The acceptability 

of a test will depend on the experience and professional 

judgment of the laboratory analyst and the reviewing 

staff of the regulatory authority. Any deviation from test 

specifications should be noted when reporting data from 

a test. 

9.11 Analytical Methods 

9.11.1 All routine chemical and physical analyses for 

culture and testing water, food, and sediment should 

include established quality assurance practices outlined 

in USEPA methods manuals (USEPA, 1979a; USEPA, 

1979b; USEPA, 1993a; USEPA, l994b). 

9.11.2 Reagent containers should be dated when received from the supplier, and the shelf life of the reagent 

should not be exceeded. Working solutions should be 

dated when prepared and the recommended shelf life 

should not be exceeded. 

9.12 Calibration and Standardization 

9.12.1 Instruments used for routine measurements of 

chemical and physical characteristics such as pH, dissolved oxygen, temperature, and conductivity should be 

calibrated before use each day according to the instrument manufacturer’s procedures as indicated in the 

general section on quality assurance (see USEPA Methods 150.1, 360.1, 170.1, and 120.1; USEPA, 1979b). 

Calibration data should be recorded in a permanent log. 

31 
9.12.2 A known-quality water should be included in the 

analyses of each batch of water samples (e.g., water 

hardness, alkalinity, conductivity). 

9.13 Replication and Test Sensitivity 

9.13.1 The sensitivity of sediment tests will depend in 

part on the number of replicates/treatment, the significance level selected, and the type of statistical analysis. 

If the variability remains constant, the sensitivity of a test 

will increase as the number of replicates is increased. 

The minimum recommended number of replicates varies with the objectives of the test and the statistical 

method used for analysis of the data (Section 14). 

9.14 Demonstrating Acceptable 

Performance 

9.14.1 It is the responsibility of a laboratory to demonstrate its ability to obtain consistent, precise results with 

reference toxicants before it performs sediment tests 

(see Section 9.16). lntralaboratory precision, expressed 

as a coefficient of variation (CV) of the range in response for each type of test to be used in a laboratory, 

should be determined by performing five or more tests 

with different batches of test organisms using the same 

reference toxicant at the same concentrations with the 

same test conditions (e.g., the same test duration, type 

of water, age of test organisms, feeding) and the same 

data analysis methods. This should be done to gain 

experience for the toxicity tests and as a point of reference for future testing. A reference toxicant concentration series (OS or higher) should be selected that will 

consistently provide partial mortalities at two or more 

concentrations of the test chemical (Section 15). 

9.14.2 Before conducting tests with contaminated sediment, the laboratory should demonstrate its ability to 

conduct tests by conducting five exposures in control 

sediment as outlined in Table 11 .I, 12.1, or 13.1. It is 

recommended that these five exposures with control 

sediment be conducted concurrently with the five reference toxicity tests descrit ed in Section 9.14.1. 

9.14.3 Laboratories should demonstrate that their personnel are able to recover an average of at least 90% of 

the organisms from whole sediment. For example, test 

organisms could be added to control sediment or test 

sediments and recovery could be determined after 1 h 

(Tomasovic et al., 1994). 

9.15 Documenting Ongoing Laboratory 

Performance 

9.15.1 Satisfactory laboratory performance on a continuing basis is demonstrated by conducting monthly 

water-only 96-h reference-toxicity tests with each test 

organism. For a given test organism, successive tests 

should be performed with the same reference toxicant at 

the same concentrations in the same type of water using 

the same data analysis method (Section 15). 

9.152 Outliers, which are data falling outside the control limits and trends of increasing or decreasing sensitivity are readily identified. If the reference toxicity datum 

from a given test falls outside the “expected” range (e.g., 

22 SD), the sensitivity of the organisms and the credibility of the test results are suspect. In this case, the test 

procedure should be examined for defects and should 

be repeated with a different batch of test organisms. 

9.153 A sediment test may be acceptable if specified 

conditions of a reference toxicity test fall outside the 

expected ranges (Section 9.10.2). Specifically, a sediment test should not automatically be judged unacceptable if the LC50 for a given reference toxicity test falls 

outside the expected range or if mortality in the control 

of the reference toxicity test exceeds 10%. All the performance criteria outlined in Tables 11.3, 12.3, and 13.4 

must be considered when determining the acceptability 

of a sediment test. The acceptability of the sediment test 

would depend on the experience and judgment of the 

investigator and the regulatory authority. 

9.15.4 Performance should improve with experience, 

and the control limits should gradually narrow, as the 

statistics stabilize. However, control limits of &2 SD, by 

definition, will be exceeded 5% of the time, regardless of 

how well a laboratory performs. For this reason, good 

laboratories that develop very narrow control limits may 

be penalized if a test result that falls just outside the 

control limits is rejected de facto. The width of the 

control limits should be considered in decisions regarding rejection of data (Section 15). 

9.16 Reference Toxicants 

9.16.1 Ideally, reference-toxicity tests should be conducted in conjunction with sediment tests to determine 

possible changes in condition of a test organism (Lee, 

1980). Water-only reference-toxicity tests should be conducted monthly. Deviations outside an established normal range may indicate a change in the condition of the 

test organism population. Results of reference-toxicity 

tests also enable interlaboratory comparisons of test 

organism sensitivity. 

9.16.2 Reference toxicants such as sodium chloride 

(NaCI), potassium chloride (KCI), cadmium chloride 

(CdCI ), and copper sulfate (CuSO,) are suitable for 

use. I4 o one reference toxicant can be used to measure 

the condition of test organisms with respect to another 

toxicant with a different mode of action (Lee, 1980). 

However, it may be unrealistic to test more than one or 

two reference toxicants routinely. KCI has been used 

successfully in round-robin water-only exposures with 

H. azteca and C. tentans (Section 15). 

9.16.3 Test conditions for conducting reference-toxicity 

tests with H. azteca, C. tentans, and L. variegates are 

outlined in Tables 9.1 and 9.2. Reference-toxicity tests 

can be conducted using one organism/chamber or muitiple organisms in each chamber. Some laboratories 
have observed low control survival when more than one 

midge/chamber is tested in water-only exposures. 

9.17 Record Keeping 

9.17.1 Proper record keeping is important. A complete 

file should be maintained for each individual sediment 

test or group of tests on closely related samples. This 

file should contain a record of the sample 

chain-of-custody; a copy of the sample log sheet; the 

original bench sheets for the test organism responses 

during the sediment test(s); chemical analysis data on 

the sample(s); control data sheets for reference toxicants; detailed records of the test organisms used in the 

test(s), such as species, source, age, date of receipt, 

and other pertinent information relating to their history 

and health; information on the calibration of equipment 

and instruments; test conditions used; and results of 

reference toxicant tests. Laboratory data should be recorded immediately to prevent the loss of information or 

inadvertent introduction of errors into the record. Original data sheets should be signed and dated by the 

laboratory personnel performing the tests. For additional 

detail see Section 14. 

Table 9.1 Recommended Test Conditions for Conducting Reference-Toxicity Tests with One Organism/Chamber 

Parameter Conditions 

1. Test type: 

2. Dilution series: 

3. Toxicant: 

4. Temperature: 

5. Light quality: 

6. Illuminance: 

7. Photoperiod: 

8. Renewal of water: 

9. Age of organisms: 

10. Test chamber: 

11. Volume of water: 

12. Number of organtsms/chamber: 

13. Number of replicate chambers/treatment: 

14. Feeding: 

15. Substrate: 

16. 

17. 

Aeration: 

Dilutron water: 

18. Test chamber cleaning: 

19. Water quality: 

20. Test duration: 

21. Endpoint: 

22. Test acceptabrlity: 

Water-only test 

Control and at least 5 test concentrations (0.5 dilutron factor) 

NaCI, KCI. Cd, or Cu 

23 f 1°C 

WIdespectrum fluorescent lights 

About 500 to 1000 Iux 

16L:BD 

H. azka: 7- to 14-d old 

C. tentans: third instar larvae’ 

L. vanegatus: adults 

30-mL plastic cups (covered with glass or plastrc) 

20 mL 

1 

10 minimum 

H. azfeca: 0.1 mL YCT (1800 mg/L stock) on Day 0 and 2 

C. tenfans: 0.25 mL Tetrafin@ (4 g/L stock) on Day 0 and 2 

L. vafiegatusr not fed 

H. azteca: NIted screen (110 mesh) 

C. fen&ins: sand (monolayer) 

L. variegafus: no substrate 

None 

Culture water, well water, surface water, site water, or 

reconstituted water 

None 

Hardness, alkalinity, conductivity, dissolved oxygen, and pH at 

the beginning and end of a test. Temperature daily 

96 h 

Survival (LC50) 

90% control survtval 

1 Age requirement: All anrmals must be third mstar or younger with at least 50% of the organisms at third instar. 

33 
Table 9.2 Recommended Test CondMions for Conducting Reference-Toxicity Tests with More Than One Organism/Chamber 

Parameter Conditions 

1. Test type: 

2. Drlution series: 

3. Towrcant: 

4. Temperature: 

5. Light qualrty: 

6. Illumrnance: 

7. Photopenod: 

8. Renewal of water: 

9. Age of organrsms: 

10. Test chamber: 

11. Volume of water: 

12. Number of organisms/chamber: 

13. Number of replrcate chambers/treatment: 

14. Feeding: 

15. Substrate: 

16. 

17. 

Aerahon: 

Drlutron water: 

18. 

19. 

Test chamber cleaning: 

Water quality: 

20. Test durahon: 

21. Endpomt: 

22. Test acceptability: 

Water-only test 

Control and at least 5 test concentrations (0.5 drlution factor) 

NaCI. KCI, Cd, or Cu 

23 1 1°C 

Wide-spectrum fluorescent lights 

About 500 to 1000 Iux 

16L:8D 

None 

ii. azteca: 7- to 14-d old 

C. tentans: third instar 

L. variegatus: adults 

250-mL glass beaker (covered with glass or plastrc) 

100 mL (mimmum) 

10 mmimum 

3 minimum 

H. azfeca: 0.5 mL YCT (1800 mqlL stock) on Day 0 and 2 

C. tentans: 1.25 mL Tetrafir% (4 g/L stock) on Day 0 and 2 

1. vanegatus: not fed 

H. azteca: Nrtex@ screen (110 mesh) 

C. tenfans: sand (monolayer) 

L. variegatusr no substrate 

None 

Culture water, well water, surface water, site water or reconstituted water 

None 

Hardness, alkalmrty, conductivrty, dissolved oxygen, and pH at 

the begmnmg and end of a test. Temperature daily 

96 h 

Survival (LC50) 

90% control survival 

1 Age requrrement: All animals must be third instar or younger with at least 50% of the organisms at third instar. 

34 
Section 10 

Collection, Culturing, and Maintaining Test Organisms 

10.1 Life Histories 

10.1.1 Hyalella azteca 

10.1.1.1 Hyalella azteca inhabit permanent lakes, ponds, 

and streams throughout North and South America (de 

March, 1981; Pennak, 1989). Occurrence of H. azteca is 

most common in warm (20 to 30°C for much of the 

summer) mesotrophic or eutrophic lakes that support 

aquatic plants. These amphipods are also found in 

ponds, sloughs, marshes, rivers, ditches, streams, and 

springs, but in lower numbers. Hyalella azteca have 

achieved densities of >1O,OOO m2 in preferred habitats 

(de March, 1981). 

10.1.1.2 Hyalella azteca are epibenthic detritivores that 

burrow into the sediment surface. Hargrave (1970) reported that H. azteca selectively ingest bacteria and 

algae. The behavior and feeding habits of H. azteca 

make them excellent test organisms for sediment assessments. 

10.1.1.3 Reproduction by H. azteca is sexual. The adult 

males are larger than females and have larger second 

gnathopods (de March, 1981). Males pair with females 

by grasping the females (amplexus) with their gnathopods 

while on the backs of the females. After feeding together 

for 1 to 7 d the female is ready to molt and the two 

organisms separate for a short time while the female 

sheds her old exoskeleton. Once the exoskeleton is 

shed, the two organisms reunite and copulation occurs. 

The male places sperm near the marsupium of the 

female and her pleopods sweep the sperm into the 

marsupium. The organisms again separate and the 

female releases eggs from her oviducts into the marsupium where they are fertilized. Hyalella azteca average 

about 18 eggs/brood (Pennak, 1989) with larger organisms having more eggs (Cooper, 1965). 

10.1.1.4 The developing embryos and newly hatched 

young are kept in the marsupium until the next molt. At 

24 to 28° hatching ranges from 5 to 10 d after fertilization (Embody, 1911; Bovee, 1950; Cooper, 1965). The 

time between molts for females is 7 to 8 d at 26 to 28°C 

(Bovee, 1950). Therefore, about the time embryos hatch, 

the female molts and releases the young. Hyalella azteca 

average 15 broods in 152 d (Pennak, 1989). Pairing of 

the sexes is simultaneous with embryo incubation of the 

previous brood in the marsupium. Hyalella azteca have 

a minimum of nine instars (Geisler, 1944). There are 5 to 

8 pre-reproductive instars (Cooper, 1965) and an indefinite number of post-reproductive instars. The first five 

instars form the juvenile stage of development, instar 

stages 6 and 7 form the adolescent stage when sexes 

can be differentiated, instar stage 8 is the nuptial stage, 

and all later instars are the adult stages of development 

(Pennak, 1989). 

10.1.1.5 Hyalella azteca have been successfully cultured at illuminance of about 500 to 1000 Iux (Ingersoll 

and Nelson, 1990; Ankley et al., 1991a; Ankley et al., 

1991 b). Hyalella azteca avoid bright light, preferring to 

hide under litter and feed during the day. 

10.1.1.6 Temperatures tolerated by H. azteca range 

from 0 to 33°C (Embody, 1911; Bovee, 1949; Sprague, 

1963). At temperatures less than 10°C the organisms 

rest and are immobile (de March, 1977; de March, 

1978). At temperatures of 10 to 18°C reproduction can 

occur. Juveniles grow more slowly at colder temperatures and become larger adults. Smaller adults with 

higher reproduction are typical when organisms are 

grown at 18 to 28°C. The highest rates of reproduction 

occur at 26 to 28°C (de March, 1978) while lethality 

occurs at 33 to 37% (Bovee, 1949; Sprague, 1963). 

10.1.1.7 Hyalella azteca are found in waters of widely 

varying types. Hyalella azteca can inhabit saline waters 

up to 29%; however, their distribution in these saline 

waters has been correlated to water hardness (Ingersoll 

et al., 1992). Hyalella azteca inhabit water with high Mg 

concentrations at conductivities up to 22,000 μS/cm, but 

only up to 12,000 μS/cm in Na-dominated waters 

(Ingersoll et al., 1992). De March (1981) reported H. 

azteca were not collected from locations where calcium 

was less than 7 mg/L. Hyalella azteca have been cultured in water with a salinity up to 15% in reconstituted 

salt water (Ingersoll et al., 1992; Winger and Lasier, 

1993). In laboratory studies, Sprague (1963) reported a 

24-h LC50 for dissolved oxygen at 20°C of 0.7 mg/L. 

Pennak and Rosine (1976) reported similar findings. 

Nebeker et al. (1992) reported 48-h and 30-d LC5Os for 

H. azteca of less than 0.3 mg/L dissolved oxygen. 

Weight and reproduction of H. azteca were reduced 

after 30-d exposure to 1.2 mg/L dissolved oxygen. 

35 
10.1.1.8 Hyalella azteca tolerate a wide range of substrates. Ingersoll and Nelson (1990) and Ingersoll et al. 

(1993) reported that H. azteca tolerated sediments ranging from more than 90% silt- and clay-sized particles to 

100% sand-sized particles without detrimental effects 

on either survival or growth. Hyalella azteca tolerated a 

wide range in grain size and organic matter in 10-d tests 

with formulated sediment (Suedel and Rodgers, 1994). 

Ankley et al. (1994a) evaluated the effects of natural 

sediment physico-chemical characteristics on the results of 10-d laboratory toxicity tests with H. azteca. C. 

tentans, and L. variegatus. Tests were conducted with 

and without the addition of exogenous food. Survival of 

organisms was decreased in tests without added food. 

Physico-chemical sediment characteristics including grain 

size and TOC were not significantly correlated to the 

response of H. azteca in either fed or unfed tests. 

10.1.2 Chironomus tentans 

10.1.2.1 Chironomus tentans have a holarctic distribution (Townsend et al., 1981) and are commonly found in 

eutrophic ponds and lakes (Flannagan, 1971; Driver, 

1977). Midge larvae are important in the diet of fish and 

waterfowl (Sadler, 1935; Siegfried, 1973; Driver et al., 

1974; McLarney et al., 1974). Larvae of C. tentans 

usually penetrate a few cm into sediment. In both lotic 

and lentic habitats with soft bottoms, about 95% of the 

chironomid larvae occur in the upper 10 cm of substrates, 

and very few larvae are found below 40 cm (Townsend 

et al., 1981). Larvae were found under the following 

conditions in British Columbia lakes by Topping (1971): 

particle size <0.15 mm to 2.0 mm, temperature 0 to 

23.3°C, dissolved oxygen 0.22 to 8.23 mg/L, pH 8.0 to 

9.2, conductivity 481 to 4,136 μmhos/cm, and sediment 

organic carbon 1.9 to 15.5%. Larvae were absent from 

lakes if hydrogen sulfide concentration in overlying water exceeded 0.3 mg/L. Abundance of larvae was positively correlated with conductivity, pH, amount of food, 

percentages of particles in the 0.59 to 1.98 mm size 

range, and concentrations of Na. K, Mg, Cl. SO, and 

dissolved oxygen. Others (e.g., Curry, 1962; Oliver, 

1971) have reported a temperature range of 0 to 35°C 

and a pH range of 7 to 10. 

10.1.2.2 Chironomus tentans are aquatic during the 

larval and pupal stages. The life-cycle of C. tentans can 

be divided into four distinct stages: (1) an egg stage, (2) 

a larval stage, consisting of four instars, (3) a pupal 

stage, and (4) an adult stage. Mating behavior has been 

described by Sadler (1935) and others (ASTM, 1994a). 

Males are easily distinguished from females because 

males have large, plumose antennae and a much thinner abdomen with visible genitalia. The male has paired 

genital claspers on the posterior tip of the abdomen 

(Townsend et al., 1981). The adult female weighs about 

twice as much as the male, with about 30% of the 

female weight contributed by the eggs. After mating, 

adult females oviposit a single transparent, gelatinous 

egg mass directly into the water. At ERL-D, the females 

oviposit eggs within 24 h after emergence. An egg mass 

contains about 2,300 eggs (Sadler, 1935) and will hatch 

in 2 to 4 d at 23°C. Under optimal conditions larvae will 

pupate and emerge as adults after about 21 d at 23°C. 

Larvae begin to construct tubes (or cases) on the second or third day after hatching. The cases lengthen and 

enlarge as the larvae grow with the addition of small 

particles bound together with threads from the mouths of 

larvae (Sadler, 1935). The larvae draw food particles 

inside the tubes and also feed in the immediate vicinity 

of either end of the open-ended tubes with their caudal 

extremities anchored within the tube. The four larval 

stages are followed by a black-colored pupal stage 

(lasting about 3 d) and emergence to a terrestrial adult 

(imago) stage. The adult stage lasts for 3 to 5 d, during 

which time the adults mate during flight and the females 

oviposit their egg masses (2 to 3 d post-emergence: 

Sadler, 1935). 

10.1.2.3 Chironomus tentans tolerate a wide range of 

substrates. Survival or growth of C. tentans was not 

reduced over a wide range in sediment grain sized in 

10-d tests with formulated sediment; however, survival 

was reduced in artificial sediments below 0.91% organic 

matter when organisms were not fed (Suedel and 

Rodgers, 1994). Ankley et al. (1994a) evaluated the 

effects of natural sediment physico-chemical characteristics on the results of 10-d laboratory toxicity tests with 

H. azteca, C. tentans, and L. variegatus. Tests were 

conducted with and without the addition of exogenous 

food. Survival and growth of organisms was decreased 

in tests without added food. Physico-chemical sediment 

characteristics including grain size and TOC were not 

significantly correlated to survival of C. tentans in tests 

in which organisms were fed. However, linear modeling 

indicated growth of C. tentans was influenced by grain 

size distribution of the test sediments (growth slightly 

increased in coarser sediment). 

10.1.3 Lumbriculus variegatus 

10.1.3.1 Lumbriculus variegatus inhabit a variety of 

sediment types throughout the United States and Europe (Chekanovskaya, 1962; Cook, 1969; Spencer, 1980; 

Brinkhurst, 1986). Lumbriculus variegatus typically tunnel in the upper aerobic zone of sediments of reservoirs, 

rivers, lakes, ponds, and marshes. When not tunneling, 

they bury their anterior portion in sediment and undulate 

their posterior portion in overlying water for respiratory 

exchange. 

10.1.3.2 Adults of L. variegatus can reach a length of 40 

to 90 mm, diameter of 1.0 to 1.5 mm, and wet weight of 

5 to 12 mg (Call et al., 1991; Phipps et al., 1993). Lipid 

content is about 1.0% (wet weight, Ankley et al., 1992b). 

Lumbriculus variegatus most commonly reproduce 

asexually, although sexual reproduction has been reported (Chekanovskaya, 1962). Newly hatched worms 

have not been observed in cultures (Call et al., 1991; 

Phipps et al., 1993. Cultures consist of adults of various 

sizes. Populations of laboratory cultures double (number of organisms) every 10 to 14 d at 20°C (Phipps et al., 

1993). 

36 
10.1.3.3 Lumbriculus variegatus tolerate a wide range 

of substrates. Ankley et al. (1994a) evaluated the effects 

of natural sediment physico-chemical characteristics on 

the results of 1 O-d laboratory toxicity tests with H. azteca, 

C. tentans, and L. variegatus. Tests were conducted 

with and without the addition of exogenous food. Survival and reproduction of organisms was decreased in 

tests without added food. Physico-chemical sediment 

characteristics including grain size and TOC were not 

significantly correlated to reproduction or growth of L. 

variegatus in either fed or unfed tests. 

10.2 General Culturing Procedures 

10.2.1 Acceptability of a culturing procedure is based in 

part on performance of organisms in culture and in the 

sediment test (Section 1.4 and 9.2). No single technique 

for culturing test organisms is required. What may work 

well for one laboratory may not work as well for another 

laboratory. While a variety of culturing procedures are 

outlined in Section 10.3 for H. azteca, in Section 10.4 for 

C. tentans, and in Section 10.5 for L. variegatus, organisms must meet the test acceptability requirements listed 

in Tables 11.3, 12.3, or 13.4. 

10.2.2 All organisms in a test must be from the same 

source. Organisms may be obtained from laboratory 

cultures, from commercial, or government sources (Table 

10.1). The test organism used should be identified using 

TaMe 10.1 Sources of Test Organisms 

Source 

U.S. Envtronmental Protection Agency 

Species 

Envlronmental Research Laboratory-Duluth 

6201 Congdon Boulevard 

Duluth. MN 55804 

Teresa Norberg-King (218/720-5500) 

H. azteca 

C. tentans 

1. var/egatus 

U.S. EnvIronmental Protection Agency 

Enwronmental Monitoring System Laboratory 

3411 Church Street 

Cincinnati, OH 45244 

Jim Lazorchak (513/569-7076) 

H. azteca 

C. fentans 

MIdwest Science Center 

Nattonal Biological Survey 

4200 New Haven Road 

Columbia, MO 65201 

Eugene Greer (3141875-5399) 

H. azteca 

c. renrans 

1. variegatus 

Great Lakes Environmental Research 1. vanegafus 

Laboratory, NOAA 

2205 Commonwealth Boulevard 

Ann Arbor, MI 48105-1593 

Peter Landrum (3 13/74 l-2276) 

Wrrght State Umversity 

Department of BiologIcal Sciences 

Dayton, OH 45435 

Allen Burton (513,‘873-2201) 

H. azteca 

C. tenfans 

L. vafiegatus 

Mlchlgan State University 

Department of Fisheries and Wildltife 

No. 13 Natural Resources Building 

East Lansing, MI 48824-l 222 

John Giesy (517/‘353-2000) 

H. azteca 

C. tentans 

L. varregatus 

an appropriate taxonomic key, and verification should 

be documented. Obtaining organisms from wild populations should be avoided unless organisms are cultured 

through several generations in the laboratory. In addition, the ability of the wild population of sexually reproducing organisms to cross-breed with the existing laboratory population must be determined. Sensitivity of the 

wild population to select contaminants (e.g., Table 1.4) 

should also be documented. 

10.2.3 Test organisms obtained from commercial sources 

should be shipped in well-oxygenated water in insulated 

containers to maintain temperature during shipment. 

Temperature and dissolved oxygen of the water in the 

shipping containers should be measured on arrival to 

determine if the organisms might have been subjected 

to low dissolved oxygen or temperature fluctuations. 

The temperature of the shipped water should be gradually adjusted to the desired culture temperature at a rate 

not exceeding 2°C per 24 h. Additional reference-toxicity 

testing is required if organisms are not cultured at the 

testing laboratory (Section 9.16). 

10.2.4 A group of organisms should not be used for a 

test if they appear to be unhealthy, discolored, or otherwise stressed (e.g., >20% mortality for 48 h before the 

start of a test). if the organisms fail to meet these 

criteria, the entire batch should be discarded and a new 

batch should be obtained. All organisms should be as 

uniform as possible in age and life stage. Test organisms should be handled as little as possible. When 

handling is necessary, it should be done as gently. 

carefully, and as quickly as possible. 

10.2.5 H. azteca, C. tentans, and L. variegatus can be 

cultured in a variety of waters. Water of a quality sufficient to culture fathead minnows (Pimephales promelas) 

or cladocerans will generally be adequate. 

10.251 Variable success has been reported using 

reconstituted waters described in ASTM (1988a) or 

USEPA (1993a) to culture or test H. azteca (USEPA. 

1992). However, the reconstituted water described in 

Section 7.1.3.4 has been used to successfully culture t-i. 

azteca (J.M. Lazorchak, USEPA, Cincinnati, OH, personal communication). The reconstituted water described 

in Section 7.1.3.4 has a higher proportion of chloride to 

sulfate compared to the reconstituted waters described 

in ASTM (1988a) and USEPA (1993a). H. artecacan be 

cultured and tested at salinities up to 15 ?60 (Ingersoll et 

al., 1992; Winger et al., 1993). 

10.252 Organisms can be cultured using either static 

or renewal procedures. Renewal of water is recommended to limit loss of the culture organisms from a 

drop in dissolved oxygen or a buildup of waste products. 

In renewal systems, there should be at least one volume 

addition/d of culture water to each chamber. In static 

systems, the overlying water volume should be changed 

at least weekly by siphoning down to a level just above 

the substrate and slowly adding fresh water. Extra care 

should be taken to ensure that proper water quality IS 

37 
maintained in static systems. For example, aeration is 

needed in static systems to maintain dissolved oxygen 

at >40% of saturation. 

be more uniform in size than organisms within a range of 

7-d old. 

10.253 A recirculating system using an under-gravel 

filter has been used to culture amphipods and midges 

(P.V. Wlnger. NBS, Athens, GA, personal communication). The approach for using a recirculating system to 

culture organisms has been described by New et al. 

(1974). Crandall et al. (1981), and Rottmann and 

Campton (1989). Under-gravel filters can be purchased 

from aquarium suppliers and consist of an elevated 

plate with holes that fit on the bottom of an aquarium. 

The plate has a standpipe to which a pump can be 

attached. Gravel or an artificial substrate (e.g., plastic 

balls or multi-plate substrates) are placed on the plate. 

The substrates provide surface area for microorganisms 

that use nitrogenous compounds. A simple example of a 

recirculating system is two aquaria positioned one above 

the other with a total volume of 120 L. The bottom 

aquarium contains the under-gravel filter system, gravel, 

or artificial substrate, and a submersible pump. The top 

aquarium is used for culture of animals and has a hole in 

the bottom with a standpipe for returning overflow water 

to the bottom aquarium. Water lost to evaporation is 

replaced weekly, and water is replaced at one- to twomonth intervals. Cultures fed foods such as Tetramin@ 

should include limestone gravel to help avoid depression in pH. Recirculating systems require less maintenance than static systems. 

10.3.2 The following procedure described by Call et al. 

(1994) and USEPA (1993) can be used to obtain 

known-age amphipods to start a test. Mature amphipods 

(50 organisms.+O-d old at 23°C) are held in 2-L glass 

beakers contalnlng 1 L of aerated culture water and 

cotton gauze as a substrate. Cotton gauze should be 

soaked in water for 24 h before use and should be 

renewed on a weekly basis. Amphipods are fed 10 mL of 

a the yeast-Cerophyl@trout chow (YCT) mixture (Appendix C), 10 mL of the green algae Se/en&rum 

capricornutum (about 35 x lo6 cells/ml), and 10 mL of 

the diatom, Navicuia spp. (1 .O x 1 Og cells/ml) on Monday. Five mL of each food is added to cultures on 

Wednesdays and Fridays. 

10.2.6 Cultures should be maintained at 23°C with a 

16L:8D photoperiod at a illuminance of about 500 to 

1000 Iux (ASTM, (1994a) and Appendix A). Cultures 

should be observed daily. Water temperature should be 

measured daily or continuously, and dissolved oxygen 

should be measured weekly. Reference-toxicity tests 

should be conducted at least monthly. If referencetoxicity 

tests are not conducted monthly, the lot of organisms 

used to start a sediment test must be evaluated using a 

reference toxicant. Culture water hardness, alkalinity, 

ammonia, and pH should be measured at least quarterly 

and the day before the start of a sediment test. If 

reconstituted water is used to culture organisms, water 

quality should be measured on each batch of reconstituted water. Culture procedures should be evaluated 

and adjusted as appropriate to restore or maintain the 

health of the culture. 

38 

10.3.2.1 Water in the culture chambers is changed 

weekly. Survival of adults and juveniles and production 

of young amphipods should be measured at this time. 

The contents of the culture chambers are poured into a 

translucent white plastic or white enamel pan. After the 

adults are removed, the remaining amphipods will range 

in age from cl- to 7-d old. Young amphipods are transferred with a pipet into a 1-L beaker containing culture 

water and are held for one week before starting a toxicity 

test. Presoaked cotton gauze is placed in the beakers, 

and organisms are fed 10 mL of YCT and 10 mL of 

green algae, and 10 mL of diatoms with renewal of 

water, and 5 mL of each food on Wednesdays and 

Fridays (Appendix C). Survival of young amphipods 

should be 90% during this one week holding period. 

Records should be kept on the number of surviving 

adults, number of breeding pairs, and young production 

and survival. This information can be used to develop 

control charts which are useful in determining if cultures 

are maintaining a vigorous reproductive rate indicative 

of culture health. Some of the adult amphipods can be 

expected to die in the culture chambers, but mortality 

greater than about 50% should be cause for concern. 

Reproductive rates in culture chambers containing 60 

adults can be as high as 500 young per week. A 

decrease in reproductive rate may be caused by a 

change in water quality, temperature, food quality, or 

brood stock health. Adult females will continue to reproduce for several months; however, young production 

gradually decreases after about three months. 

10.3 Culturing Procedures for Hyalella 

azteca 

10.3.1 The cultunng procedures described below are 

based on methods described in USEPA (1993), Ankley 

et al. (1994a), Call et al. (1994), Tomasovic et al. (1994), 

Greer ( 1993), Ingersoll and Nelson (1990), ASTM (1993a) 

and Appendix A. The culturing procedure must produce 

7- to 14-d old amphipods to start a sediment test (Table 

11.3). A narrower age range of organisms used to star-l a 

test may be desirable when growth is measured as an 

endpoint. Amphipods within a range of l- to 2-d old will 

10.3.3 A second procedure for obtaining known-age 

amphipods is described by Borgmann et al. (1989). 

Known-age amphipods are cultured in 2.5-L chambers 

containing about 1 L of culture water and between 5 and 

25 adult H. azteca. Each chamber contains pieces of 

cotton gauze presoaked in culture water. Once a week 

the test organisms are isolated from the gauze and 

collected using a sieve. Amphipods are then rinsed into 

petri dishes where the young and adults are sorted. The 

adults are returned to the culture chambers containing 

fresh water and food. 
10.3.4 A third procedure for obtaining known-age amphipods is described by Greer (1993) and Tomasovic et 

al. (1994). Mass cultures of mixed-age amphipods are 

maintained in 80-L glass aquaria containing about 50 L 

of water (Ingersoll and Nelson, 1990). Tetramin@ is 

added to each culture chamber receiving daily water 

renewals to provide about 20 g dry solids/50 L of water 

twice weekly in an 80-L culture chamber. Additional 

Tetramin@ is added when most of the Tetramin@ has 

been consumed. Laboratories using static systems should 

develop lower feeding rales specific to their systems. 

Each culture chamber has a substrate of maple leaves 

and artificial substrates (six 20cm diameter sections per 

80-L aquaria of “coiled-web material”; 3-M, St. Paul, 

MN). Before use, leaves are soaked in water for about 

30 d. The leaves are then flushed with water to remove 

residuals of naturally occurring tannic acid before placement in the cultures. 

10.3.4.1 To obtain known-age amphipods, a U.S. Standard Sieve #25 (710 pm mesh) is placed underwater in a 

chamber containing mixed-age amphipods. A #25 sieve 

will retain mature amphipods, and immature amphipods 

will pass through the mesh. Two or three pieces of 

artificial substrate (3-M coiled-web material) or a mass 

of leaves with the associated mixed-age amphipods are 

quickly placed into the sieve. The sieve is brought to the 

top of the water in the culture chamber keeping all but 

about 1 cm of the sieve under water. The artificial 

substrates or leaves are then shaken under water several times to dislodge the attached amphipods. The 

artificial substrates or leaves are taken out of the sieve 

and placed back in the culture chamber. The sieve is 

agitated in the water to rinse the smaller amphipods 

back into the culture chamber. The larger amphipods 

remaining in the sieve are transferred with a pipet into a 

dish and then placed into a shallow glass pan (e.g., pie 

pan) where immature amphipods are removed. The 

remaining mature amphipods are transferred using a 

pipet into a second #25 sieve which is held in a glass 

pan containing culture water. 

10.3.4.2 The mature amphipods are feft in the sieve in 

the pan overnight to collect any newborn amphipods 

that are released. After 24 h, the sieve is moved up and 

down several times to rinse the newborn amphipods 

(Q4-h old) into the surrounding water in the pan. The 

sieve is removed from the pan, and the mature amphipods are placed back into their culture chamber or 

placed in a second pan containing culture water if additional organisms are needed for testing. The newborn 

amphipods are moved with a pipet and placed in a 

culture chamber with flowing water during a grow-out 

period. The newborn amphipods should be counted to 

determine if adequate numbers have been collected for 

the test. 

10.3.4.3 Isolation of about 1500 (750 pairs) adults in 

amplexus provided about 800 newborn amphipods in 24 

h and required about six man-hours of time. Isolation of 

about 4000 mixed-age adults (some in amplexus and 

others not in amplexus) provided about 800 newborn 

amphipods in 24 h and required less than one man-hour 

of time. The newborn amphipods should be held for 6 to 

13 d to provide 7- to 14-d old organisms to start a test. A 

few maple leaves and a small amount of Tetramin@ is 

placed into the grow-out culture chamber to provide 

food. 

10.3.5 Laboratories that use mixed-age amphipods for 

testing must demonstrate that the procedure used to 

isolate amphipods will produce test organisms that are 

7- to 14-d old. For example, amphipods passing through 

a U.S. Standard #35 sieve (500 pm), but stopped by a 

#45 sieve (355 pm) averaged 1.54 mm (SD 0.09) in 

length (P.V. Winger, NBS, Athens, Ga, unpublished 

data). The mean length of these sieved organisms corresponds to that of 6-d old amphipods (Figure 10.1). 

After holding for 3 d before testing to eliminate organisms injured during sieving, these amphipods were about 

9-d old (length 1.84 mm, SD 0.11) at the start of a 

toxicity test. 

10.3.5.1 Ingersoll and Nelson (1990) and ASTM (1994a) 

describe the following procedure for obtaining mixed-age 

amphipods of a similar size to start a test. Smaller 

amphipods are isolated from larger amphipods using a 

stack of U.S. Standard sieves: #30 (600 pm), #40 (425 

pm), and #60 (250 pm). Sieves should be held under 

water to isolate the amphipods. Amphipods may float on 

the surface of the water if they are exposed to air. 

Artificial substrate or leaves are placed in the #30 sieve. 

Culture water is rinsed through the sieves and small 

amphipods stopped by the #60 sieve are washed tnto a 

collecting pan. Larger amphipods in the #30 and #40 

sieves are returned to the culture chamber. The smaller 

amphipods are then placed in 1-L beakers containing 

culture water and food (about 200 amphipods per beaker) with gentle aeration. 

10.3.5.2 Amphipods should be held and fed at a rate 

similar to the mass cultures for least 2 d before the start 

of a test to eliminate animals injured during handling. 

10.3.6 See Section 10.2.6 for procedures used to 

evaluate the health of cultures. 

10.4 Culturing Procedures for 

Chironomus tentans 

10.4.1 The culturing methods described below are 

based on methods described in USEPA (1993), Ankley 

et al. (1994a), Call et al. (1994), Greer (1993), ASTM 

(1994a), and Appendix A. Sediment tests must be started 

with third instar larvae (at least 50% of the larvae must 

be third instar with the remaining larvae second instar; 

Table 12.3). At a temperature of 23”C, larvae should 

develop to the third instar by 9 to 11 d after hatching 

(about 11 to 13 d post-oviposition). The instar of midges 

at the start of a test must be determined using head 

capsule width (Table 10.2). It IS also desirable to monrtor 

the weight or length of midges at the start of a sediment 

test. 

39 
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 

Day 

Mean (+I- 2SD) 

Figure 10.1 Length and relative age of H@e//e azteca collected by sleving in comparison with length of known-age organfsms. 

P.V. Wlnger, NBS, Athens, GA, unpublished data. 

Table 10.2 Chironomus tentans lnstar and Head Capsule 

Widths’ 

lwtar Days after Mean (mm) flange (mm) 

hatching 

First 1 to 4.4 0.10 0.09to 0.13 

Second 4.4 to 8.5 0.20 0.18 to 0.23 

Third 8.5 to 12.5 0.38 0.33to 0.45 

Fourth 212.5 0.67 0.63to 0.71 

: T.J. Norberg-King, USEPA. Duluth, MN. unpublished data. 

10.4.2 Recent research has indicated that the third 

instar C. tenfans were frequently referred to as the 

second instar in previous literature (T.J. Norberg-King, 

USEPA, Duluth, MN, unpublished data). When C. tentans 

larvae were measured daily, the C. tentans raised at 22 

to 24°C were third instar, not second instar, by 9 to 11 d 

after hatching. 

10.4.3 Both silica sand and shredded paper toweling 

have been used as substrates to culture C. tentans. 

Either substrate may be used if a healthy culture can be 

maintained. Greer (1993) used sand or paper toweling 

to culture midges; however, sand was preferred due to 

the ease in removing larvae for testing. Sources of sand 

are listed in Section 7. 

10.4.3.1 Paper towels are prepared according to a 

procedure adapted from Batac-Catalan and White (1982). 

Plain white kitchen paper towels are cut into strips. Cut 

toweling is loosely packed into a 2-L beaker, submersed 

in acetone, covered and placed in a fume hood, and 

soaked overnight to solubilize organic contaminants. 

The acetone is drained completely, and deionized water 

is added, brought to a boil, and stirred to drive off any 

remaining acetone vapors. This process is repeated two 

more times. Finally, the toweling is rinsed three times 

with cold deionized water. A mass of the toweling sufficient to fill a 150-mL beaker is placed into a blender 

containing 1 L of deionized water, and blended for 30 

set or until the strips are broken apart in the form of a 

pulp. The pulp is then sieved using a 710 pm sieve and 

rinsed well with deionized water to remove the shortest 

fibers. 

10.4.3.2 Dry shredded paper toweling loosely packed 

into a 2-L beaker will provide sufficient substrate for 

about ten 19-L chambers (USEPA, 1993). The shredded 

toweling placed in a 150-mL beaker produces enough 

substrate for one 19-L chamber. Additional substrate 

can be frozen in deionized water for later use. 

10.4.4 Five egg masses will provide a sufficient number 

of organisms to start a new culture chamber. Egg masses 

should be held at 23°C in a glass beaker or crystallizing 

40 
dish containing about 100 to 150 mL of culture water 

(temperature change should not exceed 2°C per d). 

Food is not added until the embryos start to hatch (in 

about 2 to 4 days at 23°C) to reduce the risk of oxygen 

depletion. A small amount of green algae (e.g., a thin 

green layer) is added to the water when embryos start to 

hatch. When most of the larvae have left the egg mass, 

150 to 200 larvae should be placed into a culture chamber. Crowding of larvae will reduce growth. Larvae that 

have formed cases can be transferred to aquaria or 

culture chambers using a gentle stream of water from a 

squeeze bottle. See Section 10.45 1 or 10.4.6.1 for a 

description of feeding rates. Larvae should reach the 

third instar by about 10 d after median hatch (about 12 to 

14 d after the time the eggs were laid). 

IO.45 Chironomus tentans are cultured in soft water at 

the USEPA laboratory in Duluth in glass aquaria (19.0-L 

capacity, 36- x 21- x 26-cm high). A water volume of 

about 6 to 8 L in these flow-through chambers can be 

maintained by drilling an overflow hole in one end 1 1 cm 

from the bottom. The top of the aquarium is covered with 

a mesh material to trap emergent adults. Pantyhose with 

the elasticized waist is positioned around the chamber 

top and the legs are cut off. Fiberglass-window screen 

glued to a glass-strip (about 2- to 3cm wide) rectangle 

placed on top of each aquarium has also been used by 

Call et al. (1994). About 200 to 300 mL of sand is placed 

in each chamber. 

10.451 Tetrafin@ food is added to each culture chamber to provide a final food concentration of about 0.04 

mg dry solids/ml of culture water. A stock suspension of 

the solids is prepared in culture water such that a total 

volume of 5.0 mL of food suspension is added daily to 

each culture chamber. For example, if a culture chamber volume is 8 L, 300 mg of food would be added daily 

by adding 5 mL of a 56-g/L stock suspension. The stock 

suspension should be well mixed immediately before 

removing an aliquot for feeding. Each batch of food 

should be refrigerated and can be used for up to two 

weeks (Appendix C). Laboratories using static systems 

should develop lower feeding rates specific to their 

systems. 

10.4.6 Chironomus tentans are cultured by Greer (1993) 

in Rubbermaid@ 5.7-L polyethylene cylindrical containers. The containers are modified by cutting a semicircle 

into the lid 17.75 cm across by 12.5 cm. Stainless-steel 

screen (20 mesh/O.4 cm) is cut to size and melted to the 

plastic lid. The screen provides air exchange, retains 

emerging adults, and is a convenient way to observe the 

culture. Two holes about 0.05 cm in diameter are drilled 

through the uncut portion of the lid to provide access for 

an air line and to introduce food. The food access hole is 

closed with a No. 00 stopper. Greer (1993) cultures 

midges under static conditions with moderate aeration, 

and about 90% of the water is replaced weekly. Each 

5.7-L culture chamber contains about 3 L of water and 

about 25 mL of fine sand. Eight to 10 chambers are used 

to maintain the culture. 

10.4.6.1 Midges In each chamber are fed 2 ml/d of a 

100 g/L Tetrafin@ suspension on Tuesday, Wednesday, 

Thursday, Friday, and Sunday. A 2-mL chlorella suspension (deactivated “Algae-Feast@ Chlorelld’, Earthrise Co., Callpatria, CA) is added to each chamber on 

Saturday and on Monday. The chlorella suspension is 

prepared by adding 5 g of dry chlorella powder/L of 

water. The mixture should be refrigerated and can be 

used for up to two weeks. 

10.4.6.2 The water should be replaced more often if 

animals appear stressed (e.g.. at surface or pale color at 

the second instar) or if the water is cloudy. Water is 

replaced by first removing emergent adults with an 

aspirator. Any growth on the sides of the chamber 

should be brushed off before water is removed. Care 

should be taken not to pour or siphon out the larvae 

when removing the water. Larvae will typically stay near 

the bottom; however, a small mesh sieve or nylon net 

can be used to catch any larvae that float out. After the 

chambers have been cleaned, temperature-adjusted 

culture water is poured back into each chamber. The 

water should be added quickly to stir up the larvae. 

Using this procedure, the approximate size, number, 

and the general health of the culture can be observed. 

10.4.7 Adult emergence will begin about three weeks 

after hatching at 23°C. Once adults begin to emerge, 

they can be gently siphoned into a dry aspirator flask on 

a daily basis. An aspirator can be made using a 250- or 

500-mL Erlenmeyer flask, a two-hole stopper, some 

short sections of 0.25 inch glass tubing, and Tygon@ 

tubing for collecting and providing suction (Figure 10.2). 

Adults should be aspirated with short inhalations to 

avoid injuring the organisms. The mouth piece on the 

aspirator should be replaced or disinfected between 

use. Sex ratio of the adults should be checked to ensure 

that a sufficient number of males are available for mating and fertilization. One male may fertilize more than 

one female. However, a ratio of one female to three 

males ensures good fertilization. 

10.4.7.1 A reproduction and oviposit chamber may be 

prepared in several different ways (Figure 10.2). Culture 

water (about 50 to 75 mL) can be added to the aspiration 

flask in which the adults were collected (Figure 10.2; 

Batac-Catalan and White, 1982). ERL-Duluth (USEPA, 

1993) uses a 500-mL collecting flask with a length of 

Nitex@ screen positioned vertically and extending into 

the cutture water (Figure 10.2). The Nitex@ screen is 

used by the females to position themselves just above 

the water during oviposition. The two-hole stopper and 

tubing of the aspirator should be replaced by screened 

material or a cotton plug for good arr exchange in the 

oviposition chamber. 

10.4.7.2 Greer (1993) used an oviposition box to hold 

emergent adults. The box is constructed of a 5.7-L 

chamber with a 20-cm tall cylindrical chamber on top. 

The top chamber is constructed of stainless steel screen 

(35 meshI2.54 cm) melted onto a plastic lid with a 17.75 

cm hole. A 5cm hole is cut into the side of the bottom 

41 
Tygon Tubing 

B 

m 2 Hole Stopper 

\ 

500 ml Erlenrneyer 

Mesh Cover 

Figure 10.2 Aspirator chamber (A) and reproduction and oviposit chamber (B) for adult midges. 

42 
chamber and a #l 1 stopper is used to close the hole. 

Egg masses are removed by first sliding a piece of 

plexiglass between the top and bottom chambers. Adult 

midges are then aspirated from the bottom chamber. 

The top chamber with plexiglass is removed from the 

bottom chamber and a forceps is used to remove the 

egg masses. The top chamber is put back on top of the 

bottom chamber, the plexiglass is removed, and the 

aspirated adults are released from the aspirator into the 

chamber through the 5-cm hole. 

10.4.8 About two to three weeks before the start of a 

test, at least 3 to 5 egg masses should be isolated for 

hatching using procedures outhned in Section 10.4.3. 

10.4.9 Records should be kept on the time to first 

emergence and the success of emergence for each 

culture chamber. It is also desirable to monitor growth 

and head capsule width periodically in the cultures. See 

Section 10.2.6 for additional detail on procedures for 

evaluating the health of the cultures. 

10.5 Culturing Procedures for 

Lumbricuius variegatus 

10.51 The culturing procedures described below are 

based on methods described in Phipps et al. (1993), 

USEPA (1993), Call et al. (1994), E.L. Brunson (NBS, 

Columbia, MO, unpublished data), and Appendix A. 

Bioaccumulation tests are started with adult organisms. 

1 OS.2 Lumbriculus variegatus are generally cultured 

with daily renewal of water (57- to 80-L aquaria containing 45 to 50 L of water). 

10.5.3 Paper towels can be used as a substrate for 

culturing L. variegates (Phipps et al., 1993). Substrate is 

prepared by cutting unbleached brown paper towels into 

strips either with a paper shredder or with a scissors. 

Cut toweling is loosely packed into a 2-L beaker, submersed in acetone, covered and placed in a fume hood, 

and soaked overnight to solubilire organic contaminants. The acetone is discarded, and the towels are 

rinsed several times with deionized water. Deionized 

water is added, brought to a boil, and stirred three times 

to drive off the acetone vapors. This is repeated two 

more times. The strips are conditioned for at least one 

week by placing 4 L of strips into an aquarium equipped 

with two water lines each having a flow capacity of about 

100 mUmin. One line is placed below and one line is 

placed above the towel mass. Glass weights (several 

2.5-cm x 25.4-cm glass strips standing on edge and 

glued on both ends to glass strips about 50 cm in length) 

can be placed on the strips to prevent floating. This 

approach creates a uniform water flow throughout the 

strips and minimizes fouling. Following conditioning, the 

strips are removed and evenly placed on the bottom of 

the culture chamber. Glass weights keep the strips in 

place. Conditioned strips can be frozen in deionized 

water for later use. The substrate is renewed with condrtioned towels when thin or bare areas appear. The 

substrate in the chamber will generally last for about two 

months. 

10.54 Oligochaetes probably obtain nourishment from 

ingesting the organic matter in the substrate (Pennak. 

1989). Lumbriculus variegafus in each of the culture 

chambers are fed a lo-mL suspension of 6 g of trout 

starter 3 times/week. The particles will temporarrly disperse on the surface film, break through the surface 

tension, and settle out over the substrate. Laboratories 

using static systems should develop lower feedlng rates 

specific to their systems. Food and substrate used to 

culture oligochaetes should be analyzed for compounds 

to be evaluated in bioaccumulation tests. If the concentration of the test compound is above the detectton level 

and the food is not measured, the test may be invalidated. Recent studies in other laboratories, for example. 

have indicated elevated concentrations of PCBs in substrate and/or food used for culturing the oligochaete (J. 

Amato, AScl Corporation, Duluth, MN, personal communication). 

10.5.5 Phipps et al. (1993) recommend starting a new 

culture with 500 to 1000 worms. Conditioned paper 

toweling should be added when the substrate In a 

culture chamber is thin. 

10.5.6 On the day before the start of a test, oligochaetes 

can be isolated by transferring substrate from the cultures into a beaker using a fine mesh net. Additional 

organisms can be removed using a glass pipet (20 cm 

long, 5 mm i.d.) (Phipps et al., 1993). Water can be 

slowly trickled into the beaker. The oligochaetes will 

form a mass and most of the remaining substrate will be 

flushed from the beaker. On the day the test is started, 

organisms can be placed in glass or stainless-steel 

pans. A gentle stream of water from the pipet can be 

used to spread out clusters of oligochaetes. The remaining substrate can be siphoned from the pan by allowing 

the worms to reform in a cluster on the bottom of the 

pan. For bioaccumulation tests, aliquots of worms to be 

added to each test chamber can be transferred using a 

blunt dissecting needle or dental pick. Excess water can 

be removed during transfer by touching the mass of 

oligochaetes to the edge of the pan. The mass of 

oligochaetes is then placed in a tared weigh boat. qurckly 

weighed, and immediately introduced into the appropriate test chamber. Organisms should not be blotted with 

a paper towel to remove excess water. 

10.5.7 The culture population generally doubles (number of organisms) in about 10 to 14 d. See Section 

10.2.6 for additional detail on procedures for evaluating 

the health of the cuttures. 

43 
Section 11 

Test Method 100.1 

Hyalella azteca 10-d Survival Test for Sediments 

11.1 Introduction 

11.1.1 Hyalella azteca (Saussure) have many desirable 

characteristics of an ideal sediment toxicity testing organism including relative sensitivity to contaminants associated with sediment, short generation time, contact 

with sediment, ease of culture in the laboratory, and 

tolerance to varying physico-chemical characteristics of 

sediment. Their response has been evaluated in interlaboratory studies and has been confirmed with natural 

benthos populations. Many investigators have successfully used H. azteca to evaluate the toxicity of freshwater 

sediments (e.g., Nebeker et al., 1984a; Borgmann and 

Munwar, 1989; Ingersoll and Nelson, 1990; Ankley et 

al., 1991a, Ankley et al., 1991b; Burton et al., 1989; 

Winger and Lasier, 1993; Kemble et al., 1994). Ii. 

azteca has been used for a variety of sediment assessments (Ankley et al., 1991; West et al., 1993; Hoke et 

al., 1994; West et at., 1994; and Hoke et al., 1994). 

Hyalella azteca can also be used to evaluate the toxicity 

of estuarine sediments (up to 15% salinity; Nebeker 

and Miller, 1988; Roach et al., 1992, Winger et al., 

1993). Endpoints typically monitored in sediment toxicity 

tests with H. azteca Include survival and growth. 

11.1.2 A specific test method for conducting a 10-d 

sediment toxicity test is described in Section 11.2 for H. 

azteca. Methods outlined in Appendix A and the literature cited in Table A.2 were used for developing test 

method 100.1. Results of tests using procedures different from the procedures described in Section 11.2 may 

not be comparable, and these different procedures may 

alter contaminant bioavailability. Comparison of results 

obtained using modified versions of these procedures 

might provide useful information concerning new concepts and procedures for conducting sediment tests with 

aquatic organisms. If tests are conducted with procedures different from the procedures described in this 

manual. additional tests are required to determine comparability of results (Section 1.3). 

11.2 Recommended Test Method for 

Conducting a 10-d Sediment 

Toxicity Test with Hyalella azteca 

11.2.1 Recommended conditions for conducting a 10-d 

sediment toxicity test with H. azteca are summarized in 

Table 11.1. A general activity schedule is outlined in 

Table 11.2. Decisions concerning the various aspects of 

experimental design, such as the number of treatments, 

number of test chambers/treatment, and water-quality 

characteristics should be based on the purpose of the 

test and the methods of data analysis (Section 14). The 

number of replicates and concentrations tested depends 

in part on the significance level selected and the type of 

statistical analysis. When variability remains constant, 

the sensitivity of a test increases as the number of 

replicates increase. 

11.2.2 The recommended 10-d sediment toxicity test 

with H. azteca must be conducted at 23°C with a 16L:8D 

photoperiod at an illuminance of about 500 to 1000 Iux 

(Table 11.1). Test chambers are 300-mL high-form lipless 

beakers containing 100 mL of sediment and 175 mL of 

overlying water. Ten 7- to 14-d old amphipods are used 

to start a test. The number of replicates/treatment depends on the objective of the test. Eight replicates are 

recommended for routine testing (Section 14). Amphipods in each test chamber are fed 1.5 mL of YCT food 

daily (Appendix C). Each chamber receives 2 volume 

additions/d of overlying water. Water renewals may be 

manual or automated, and Appendix B describes 

water-renewal systems that can be used to deliver overlying water. Overlying water can be culture water, well 

water, surface water, site water, or reconstituted water. 

For site-specific evaluations, the characteristics of the 

overlying water should be as similar as possible to the 

site where sediment is collected. Requirements for test 

acceptability are summarized in Table 11.3. 

11.3 General Procedures 

11.3.1 Sediment into Test Chambers: The day before 

the sediment test is started (Day -1) each sediment 

should be thoroughly mixed and added to the test chambers (Section 8.3.1). Sediment should be visually inspected to judge the degree of homogeneity. Excess 

water on the surface of the sediment can indicate separation of solid and liquid components. If a quantitative 

measure of homogeneity is required, replicate subsamples should be taken from the sediment batch and 

analyzed for TOC, chemical concentrations, and particle 

size. 

11.3.1.1 Each test chamber should contain the same 

amount of sediment, determined either by volume or by 

44 
Table 11.1 Test Conditions for Conducting a 10-d Sediment Toxicity Test with Hyalella azteca 

Parameter Conditions 

1. Test type: 

2. Temperature: 

3. Light quality: 

4. Illuminance: 

5. Photoperiod: 

6. Test chamber: 

7. Sediment volume: 

8. Overlying water volume: 

9. Renewal of overlying water: 

10. 

11. 

12. 

Age of organisms: 

Number of organisms/chamber: 

Number of replicate 

chambers/treatment: 

13. Feeding: 

14. Aeration: 

15. Overlying water: 

16. lest chamber cleaning: 

17. Overlying water quality: 

18. Test duration: 

19. Endpoints: 

20. Test acceptability: 

Whole-sediment toxicity test with renewal of overlying water 

23±1°C 

Wide-spectrum fluorescent lights 

About 500 to 1000 lux 

16L:8D 

300-mL high-form lipless beaker 

100 ml 

175 mL 

2 volume additions/d; continuous or intermittent (e.g., one volume 

addition every 12 h) 

7- to 14-d old at the start of the test 

10 

Depends on the objective 

of the test. Eight replicates are recommended for routine testing (see 

Section 14) 

YCT food, fed 1.5 mL daily to each test chamber 

None, unless dissolved oxygen in overlying water drops below 40% of 

saturation 

Culture water, well water, surface water, site water, or reconstituted 

water 

If screens become clogged during a test, gently brush the outside of 

the screen (Appendix B) 

Hardness, alkalinity, conductivity, pH. and ammonia at the beginning 

and end of a test. Temperature and dissolved oxygen daily 

10d 

Survival (growth optional) 

Minimum mean control survival of 80% and performance-based criteria 

specifications outlined in Table 11.3 

Table 11.2 General Activity Schedule for Conducting a Sediment Toxicity Test with HyaIe//a azteca 1 

Day Activity 

-7 Separate known-age amphipods from the cultures and place in holding chambers, Begin preparing food for the test. 

-6 to -2 

-1 

0 

Feed and observe isolated amphipods, monitor water quality (e.g., temperature and dissolved oxygen). 

Feed and observe isolated amphipods, monitor water quality. Add sediment into each test chamber. place chambers into 

exposure system, and start renewing overlying water. 

Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductivity, ammonia). Transfer 10 7- to 

14-day old amphipods into each test chamber. Release organisms under the surface of the water. Add 1.5 mL of YCT into 

each test chamber. Archive 20 test organisms for weight or length determination. Observe behavior of test organisms. 

1 to 8 Add 1.5 mL of YCT food to each test chamber. Measure temperature and dissolved oxygen. Observe behavior of test 

organisms. 

9 Same as Day 1. Measure total water quality. 

10 Measure temperature and dissolved oxygen. End the test by collecting the amphipods with a sieve. Count survivors and set 

aside organisms for weight or length measurements. 

Modified from Call et al., 1994 

45 
Table 11.3 Test Acceptability Requirements for a l&d Sediment Toxicity Test with Hyalella azteca 

A. It IS recommended for conducting a 1 O-d test wrth Hyalelfa azteca that the following performance cntena be met: 

1 Age ot H. azteca at the start of the test must be between 7- to 14-d old. 

2. Average survrval of H. azteca In the control sediment must be greater than or equal to 80% at the end of the test. 

3 Haraness, alkalrmty. pti. and ammonia In the overlying water withn a treatment should not vary by more than 50% during the test. 

8. Performance-based crrterra for culturmg H. azfeca Include 

1. Laboratones should perform monthly 96-h water-only reference-toxrcity tests to assess the sensrirvrty of culture organisms. If reference-toxicity tests are not conducted monthly, the lot of organisms used to start a sediment test must be evaluated using a reference 

toxlcant (Section 9.16). 

2. Laboratones should track parentat survrval In the cultures and record ths nformatron usng control charts 11 known-age cultures are 

marrtained. Records shodId also be kept on the frequency of restartrng cultures and the age of brood organisms. 

3. Laboratones shculd record the followmg water-quality charactershcs of the cultures at least quarterly and the day before the start of a 

sedrment test: pf-l. hardness. alkalin,ty, and ammonra. Dssolved oxygen should be measured weekly. Temperature should be 

recorded daily. 

4. Laboratories should charactenze and monitor background contamination and nutrient quality of food if problems are observed in 

cultunng or testing organisms. 

5. Physologrcal measurements such as lipid content might provrde useful Information regarding the health of the cultures. 

C. AdrWonal reourrements: 

1. All orgamsms In a test must be from the same source. 

2. It IS desirable to start tests soon after collectron of sedrment from the field (see Section 8.2 for additlonal detarl). 

3. All test charrbers (and compartments) should be rdentlcal and should contam the same amount of sediment and overlymg water. 

4. Negatrve-control sediment and appropriate solvent controls must be included III a test. The concentratron of solvent used must not 

aoversely affect test organisms. 

5 Test orgamsms must be cultured and tested at 23°C. 

6. The dally mean !est temperature must be within -1°C of the desired temperature. The Instantaneous temperature must always be 

wrthrn -3’C of the desred temperature. 

7. Natural physco-chemical characteristrcs of test sediment collected from the field should be withm the tolerance llmrts of the test 

organisms. 

.- - -- - 

weight. Overlying water is added to the chambers in a 

manner that minimrzes suspension of sediment. This 

can be accomplished by gently pouring water along the 

sides of the chambers or by pouring water onto a baffle 

(e.g., a circular piece of Teffor@ with a handle attached) placed above the sediment to dissipate the 

torte of the water. Renewal of overlying water is started 

on Day - 1. A test begins when the organisms are added 

to the test chambers (Day 0). 

11.3.2 Renewal of overlying water: Renewal of overlying water IS required during a test. At any particular 

time during the test, flow rates through any two test 

chambers should not differ by more than 10%. Mount 

and Brungs (1967) diluters have been modified for 

sediment testing, and other automated water delivery 

systems have also been used (Maki, 1977; Ingersoll 

and Nelson. 1990; Benoit et al., 1993; Zumwalt et al., 

1994). The water-delivery system should be calibrated 

before a test is started to verify that the system is 

functioning properly. Renewal of overlying water is 

started on Day -1 before the addition of test organisms 

or food on Day 0. Appendix B describes water-renewal 

systems that can be used for conducting sediment 

tests. 

11.3.2.1 In water-renewal tests with one to four volume 

additions of overlying water/d, water-quality characteristics generally remain similar to the inflowing water 

(Ingersoll and Nelson, 1990; Ankley et al., 1993); however, in static tests, water quality may change profoundly during the exposure (Shuba et al., 1978). For 

example, in static whole-sediment tests, the alkalinity, 

hardness, and conductivity of overlying water more than 

doubled in several treatments during a four-week exposure (Ingersoll and Nelson, 1990). Additionally, concentrations of metabolic products (e.g., ammonia) may also 

increase during static exposures, and these compounds 

can either be directly toxic to the test organisms or may 

contribute to the toxicity of the contaminants in the 

sediment. Furthermore, changes in water-quality characteristics such as hardness may influence the toxicity 

of many inorganic (Gauss et al., 1985) and organic 

(Mayer and Ellersieck, 1986) contaminants. Although 

contaminant concentrations are reduced in the overlying 

water in water-renewal tests, organisms in direct contact 

with sediment generally receive a substantial proportion 

of a contaminant dose directly from either the whole 

sediment or from the interstitial water. 

46 
11.3.3 Acclimation: Test organisms must be cultured 

and tested at 23°C. Ideally, test organisms should be 

cultured in the same water that will be used in testing. 

However, acclimation of test organisms to the test water 

is not required. 

11.3.4 Placing Organisms in Test Chambers: Test 

organisms should be handled as little as possible. Amphipods should be introduced into the overlying water 

below the air-water interface. Test organisms can be 

pipetted directly into overlying water (Ankley et al., 1993). 

Alternatively, test organisms can be placed into 30-mL 

counting cups that are floated in the test chambers for 

15 min before organisms are introduced into the overlying water (Ingersoll and Nelson, 1990). Length or weight 

should be measured on a subset of at least 20 organisms used to start the test. 

11.3.5 Monitoring a Test: All chambers should be 

checked daily and observations made to assess test 

organism behavior such as sediment avoidance. However, monitoring effects on burrowing activity of test 

organisms may be difficult because the test organisms 

are often not visible during the exposure. The operation 

of the exposure system should be monitored daily. 

11.3.5.1 Measurement of Overlying Wafer-quality 

Characteristics: Conductivity, hardness, pH, alkalinity, 

and ammonia should be measured in all treatments at 

the beginning and end of a test. Overlying water should 

be sampled just before water renewal from about 1 to 2 

cm above the sediment surface using a pipet. It may be 

necessary to pool water samples from individual replicates. The pipet should be checked to make sure no 

organisms are removed during sampling of overlying 

water. Hardness, alkalinity, pH, conductivity, and ammonia in the overlying water with a treatment should not 

vary by more than 50% during a test. 

11.3.5.1 .l Dissolved oxygen should be measured daily 

and should be between 40 and 100% saturation (ASTM, 

1988a). If a probe is used to measure dissolved oxygen 

in overlying water, it should be thoroughly inspected 

between samples to make sure that organisms are not 

attached and should be rinsed between samples to 

minimize cross contamination. Aeration can be used to 

maintain dissolved oxygen in the overlying water above 

40% saturation. Dissolved oxygen and pH can be measure directly in the overlying water with a probe. 

11.3.5.1.2 Temperature should be measured at least 

daily in at least one test chamber from each treatment. 

The temperature of the water bath or the exposure 

chamber should be continuously monitored. The dally 

mean test temperature must be within +l”C of the 

desired temperature. The instantaneous temperature 

must always be within +3*C of the desired temperature. 

11.3.6 Feeding: Without addition of food, the test 

organisms may starve during exposures. However, the 

addition of the food may alter the availability of the 

contaminants in the sediment (Wiederholm et al.. 1987; 

Harkey et al., 1994). Furthermore, if too much food is 

added to the test chamber or if the mortality of test 

organisms is high, fungal or bacterial growth may develop on the sediment surface. Therefore, the amount of 

food added to the test chambers is kept to a minimum. 

11.3.6.1 Suspensions of food should be thoroughly 

mixed before aliquots are taken. If excess food collects 

on the sediment, a fungal or bacterial growth may develop on the sediment surface, in which case feeding 

should be suspended for one or more days. A drop in 

dissotved oxygen betow 40% of saturation during a test 

may indicate that the food added is not being consumed. 

Feeding should be suspended for the amount of time 

necessary to increase the dissolved oxygen concentration (ASTM, 1994a). If feeding is suspended In one 

treatment, it should be suspended in all treatments. 

Detailed records of feeding rates and the appearance of 

the sediment surface should be made daily. 

11.3.7 Ending a Test: Any of the surviving amphipods 

in the water column or on the surface of the sediment 

can be pipetted from the beaker before sieving the 

sediment. Immobile organisms isolated from the sediment surface or from sieved material should be considered dead. Ankley et al. (1994a) recommend using a 

#25 sieve (710 pm mesh) to remove amphipods from 

sediment. Alternatively, Kemble et al. (1994) recommend sieving sediment using the following procedure: 

(1) pour about half of the overlying water through a tt50 

(300 pm) U.S. Standard mesh sieve, (2) swirl the remaining water to suspend the upper 1 cm of sediment. 

(3) pour this slurry through the #50 mesh sieve and 

wash the contents of the sieve into an examination pan, 

(4) rinse the coarser sediment remaining in the test 

chamber through a #40 (425 pm) mesh sieve and wash 

the contents of this second sieve into a second examination pan. Surviving test organisms can be removed 

from the two pans and preserved in 8% sugar tormalin 

solution for growth measurements (Ingersoll and Nelson, 

1990). NoTox@ (Earth Safe Industries, Belle Mead. NJ) 

can be used as a substitute for formalin (Unger et al.. 

1993). 

11.3.7.1 A consistent amount of time should be taken to 

examine sieved material for recovery of test organisms 

(e.g., 10 minkeplicate). Laboratories should demonstrate that their personnel are able to recover an average of at least 90% of the organisms from whole sediment. For example, test organisms could be added to 

control or test sediments, and recovery could be determined after 1 h (Tomasovic et al., 1994). 

11.3.8 Test Data: Survival is the prrmary endpoint 

recorded at the end of the 10-d sediment toxicity test 

with H. azleca. Measuring growth is optional; however. 

growth of amphipods may be a more sensitive toxicity 

endpoint compared to survival (Burton and Ingersoll. 

1994; Kemble et al., 1994). The duration of the 1 O-d test 

started with 7- to 14-d old amphipods is not long enough 

to determine sexual maturation or reproductive effects. 

47 
11.3.8.1 Amphipod body length (kO.1 mm) can be 

measured from the base of the first antenna to the tip of 

the third uropod along the curve of the dorsal surface. 

Ingersoll and Nelson (1990) describe the use of a digitizing system and microscope to measure lengths of H. 

azteca. Antenna1 segment number can also be used to 

estimate length or weight of amphipods (EL Brunson, 

NBS, Columbia, MO, personal communication). Wet or 

dry weight measurements have also been used to estimate growth of H. azteca (ASTM, 1994a). Dry weight of 

amphipods should be determined by pooling all living 

organisms from a replicate and drying Jhe sample at 

about 60 lo 90% to a constant weight. The sample is 

brought to room temperature in a desiccator and weighed 

to the nearest 0.01 mg to obtain mean weight per 

surviving organism per replicate. 

11.4 Interpretation of Results 

11.4.1 Section 14 describes general information for 

interpretation of test results. The following sections describe species-specific information that is useful in helpIng to interpret the results of sediment toxicity tests with 

H. azteca. 

11.4.2 Age Sensitivity: The sensitivity of H. azteca 

appears to be relatively similar up to at least 24-to 26-d 

oid organisms (Collyard et al., 1994). For example, the 

toxicity of diazinon, Cu, Cd, and Zn was similar in 96-h 

water-only exposures starting with 0- to 2-d old organisms through 24- to 26-d old organisms (Figure 11.1). 

The toxicity of alkylphenol ethoxylate (a surfactant) tended 

to increase with age. In general, this suggests that tests 

started with 7- to 14-d old amphipods would be representative of the sensitivity of H. azteca up to at least the 

adult life stage. 

11.4.3 Grain Size: Hyalella azteca are tolerant of a 

wide range of substrates. Physicochemical characteristics (e.g., grain size or TOC) of sediment were not 

significantly correlated to the response of H. azteca in 

toxicity tests in which organisms were fed (Section 

10.1 .1.8; Ankley et al., 1994a). 

11.4.4 isolating Organisms at the End of a Test: 

Quantitative recovery of young amphipods (e.g., 0- to 

7-d old) is difficult given their small size (Figure 11.2, 

Tomasovic et al., 1994). Recovery of older and larger 

amphipods (e.g., 21d old) is much easier. This was a 

primary reason for deciding to start 10-d tests with 7- to 

14-d old amphipods (organisms are 17- to 24-d old at 

the end of the 1Od test). 

11.4.5 Influence of Indigenous Organisms: Survival 

of H. azleca in 28-d tests was not reduced in the 

presence of oligochaetes in sediment samples 

(Reynoldson et al., 1994). However, growth of amphipods was reduced when high numbers of oligochaetes 

were placed in a sample. Therefore, it is important to 

determine the number and biomass of indigenous organisms in field-collected sediment in order to better 

interpret growth data (Reynoldson et al., 1994). Furthermore, presence 01 predators may also influence the 

response of test organisms in sediment (Ingersoll and 

Nelson, 1990). 

48 
8 

'I 

O-2 2-4 6-8 8-10 12-14 1818 20-22 24-26 O-2 2-4 4-6 6-8 lC!-12 16-18 20-22 24-26 

AgeClass AgeClass 

6Q2 

i f 

f 

OJ, r 

o-2 2-4 4-6 6-8 lo-12 18-20 20-22 o-2 2-4 4-6 10-12 16-18 24-26 

AgeClass AgeClass (d) 

, 

=j 400- 

5 

2 

Q 

> 

2 .- 

N 

300- 11 

200 

1 

o-2 2-4 4-6 8-10 12-14 24-26 

AgeClass (d) 

Figure 11.1 Lifestage sensitivity of Hyalella azfeca in 96-h water-only exposures. 

49 
80 

60 

z 

e 

5 

$ 40 

8 CT 

20 --------------------------------- 

- - 

0 

1 7 14 21 

Age (days) 

Figure 11.2 Average recovery of different age Hyafe//a azteca from sediment by 7 individuals. 

50 
Section 12 

Test Method 100.2 

Chironomus tentans 10-d Survival and Growth Test for Sediments 

12.1 Introduction 

12.1.1 Chironomus tentans (Fabricius) have many desirable characteristics of an ideal sediment toxicity testing organism including relative sensitivity to contaminants associated with sediment, contact with sediment, 

ease of culture in the laboratory, tolerance to varying 

physico-chemical characteristics of sediment, and short 

generation time. Their response has been evaluated in 

interlaboratory studies and has been confirmed with 

natural benthos populations. Many investigators have 

successfully used C. tentans to evaluate the toxicity of 

freshwater sediments (e.g., Wentsel et al., 1977; Nebeker 

et al., 1984a; Nebeker et al., 1988; Adams et al., 1985; 

Giesy et al., 1988; Hoke et al., 1990; West et al., 1993; 

Ankley et al., 1993; Ankley et al., 1994a; Ankley et 

al.,1994b). C. tentans has been used for a variety of 

sediment assessments (West et al., 1993; Hoke et al., 

1994; West et al., 1994; and Ankley et al., 1994c). 

Endpoints typically monitored in sediment toxicity tests 

with C. tentans include survival and growth (ASTM, 

1994a). 

12.1.2 A specific test method for conducting a 10-d 

sediment toxicity test is described in Section 12.2 for C. 

tentans. Methods outlined in Appendix A and the literature cited in Table A.3 were used for developing test 

method 100.2. Results of tests using procedures different from the procedures described in Section 12.2 may 

not be comparable and these different procedures may 

alter contaminant bioavailability. Comparison of results 

obtained using modified versions of these procedures 

might provide useful information concerning new concepts and procedures for conducting sediment tests with 

aquatic organisms. If tests are conducted with procedures different from the procedures described in this 

manual, additional tests are required to determine comparability of results (Section 1.3). 

12.2 Recommended Test Method for 

Conducting a 10-d Sediment 

Toxicity Test with Chironomus 

tentans 

12.2.1 Recommended conditions for conducting a 10-d 

sediment toxicity test with C. tentans are summarized in 

Table 12.1. A general activity schedule is outlined in 

Table 12.2. Decisions concerning the various aspects of 

experimental design, such as the number of treatments, 

number of test chambers/treatment, and waterquality 

characteristics should be based on the purpose of the 

test and the methods of data analysis (Section 14). The 

number of replicates and concentrations tested depends 

in part on the significance level selected and the type of 

statistical analysis. When variability remains constant, 

the sensitivity of a test increases as the number of 

replicates increases. 

12.2.2 The recommended 10-d sediment toxicity test 

with C. tentans must be conducted at 23°C with a 

16L:8D photoperiod at an illuminance of about 500 to 

1000 lux (Table 12.1). Test chambers are 300-mL 

high-form lipless beakers containing 100 mL of sediment and 175 mL of overlying water. Ten third-instar 

midges are used to start a test. All organisms must be 

third instar or younger with at least 50% of the organisms at third instar. The number of replicates/treatment 

depends on the objective of the test. Eight replicates are 

recommended for routine testing (see Section 14). Midges 

in each test chamber are fed 1.5 mL of a 4-g/L Tetrafin® 

suspension daily. Each chamber receives 2 volume 

additions/d of overlying water. Water renewals may be 

manual or automated, and Appendix B describes 

water-renewal systems that can be used to deliver overlying water. Overlying water can be culture water, well 

water, surface water, site water, or reconstituted water. 

For site-specific evaluations, the characteristics of the 

overlying water should be as similar as possible to the 

site where sediment is collected. Requirements for test 

acceptability are summarized in Table 12.3. 

12.3 General Procedures 

12.3.1 Sediment into Test Chambers: The day before 

the sediment test is started (Day -1) each sediment 

should be thoroughly mixed and added to the test chambers (Section 8.3.1). Sediment should be visually inspected to judge the extent of homogeneity. Excess 

water on the surface of the sediment can indicate separation of solid and liquid components. If a quantitative 

measure of homogeneity is required, replicate subsamples should be taken from the sediment batch and 

analyzed for TOC, chemical concentrations, and particle 

size. 

51 
Table 12.1 Recommended Test Conditions for Conducting a 10-d Sediment Toxicity Test with Chironomus tentans 

Parameter Conditions 

1. Test type: 

2. Temperature: 

3. Light quality: 

4. Illuminance: 

5. Photoperiod: 

6. Test chamber: 

7. Sediment volume: 

8. Overlying water volume: 

9. Renewal of overlying water: 

10. Age of organisms: 

11. Number of organisms/ 

chamber: 

12. Number of replicate 

chambers/treatment: 

13. Feeding: 

14. Aeration: 

15. Overlying water: 

16. Test chamber cleaning: 

17. Overlying water quality: 

18. Test duration: 

19. Endpoints: 

20. Test acceptability: 

Whole-sediment toxicity test with renewal of overlying water 

23 ± 1°C 

Widespectrum fluorescent lights 

About 500 to 1000 Iux 

16L:8D 

high-form lipless beaker 

100 mL 

175 mL 

2 volume additions/d;continuous or intermittent (e.g., one volume addition 

every 12 h) 

Third instar larvae (All organisms must be third instar or younger with at 

least 50% of the organisms at third instar) 

10 

Depends on the objective of 

the test. Eight replicates are recommended for routine testing (see 

Section 14) 

Tetrafin® goldfish food, fed 1.5 mL daily to each test chamber (1.5 mL 

contains 4.0 mg of dry solids) 

None, unless dissolved oxygen in overlying water drops below 40% of 

saturation 

Culture water, well water, surface water, site water, or reconstituted water 

If screens become clogged during a test; gently brush the outside of the 

screen (Appendix 6) 

Hardness, alkalinity, conductivity, pH, and ammonia at the beginning and 

end of a test. Temperature and dissolved oxygen daily 

10d 

Survival and growth (dry weight) 

Minimum mean control survival of 70% and mean weight per surviving 

control organism of 0.6 mg. Performance-based criteria specifications 

outlined in Table 12.3 

12.3.1.1 Each test chamber should contain the same 

amount of sediment, determined either by volume or by 

weight. Overlying water is added to the chambers in a 

manner that minimizes suspension of sediment. This 

can be accomplished by gently pouring water along the 

sides of the chambers or by pouring water onto a baffle 

(e.g., a circular piece of Teflon with a handle attached) 

placed above the sediment to dissipate the force of the 

water. Renewal of overlying water is started on Day -1. 

A test begins when the organisms are added to the test 

chambers (Day 0). 

12.3.2 Renewal of overlying water: Renewal of overlying water is required during a test. At any particular 

time during the test, flow rates through any two test 

chambers should not differ by more than 10%. Mount 

and Brungs (1967) diluters have been modified for sediment testing, and other automated water delivery systems have also been used (Maki, 1977; Ingersoll and 

Nelson, 1990; Benoit et al., 1993; Zumwalt et al., 1994). 

Each water-delivery system should be calibrated before 

a test is started to verify that the system is functioning 

properly. Renewal of overlying water is started on Day 

-1 before the addition of test organisms or food on Day 

0. Appendix B describes water-renewal systems that 

can be used for conducting sediment tests. 

12.3.2.1 In water-renewal tests with one to four volume 

additions of overlying water/d, water-quality characteristics generally remain similar to the inflowing water 

(Ingersoll and Nelson, 1990; Ankley et al., 1993); however, in static tests, water quality may change profoundly during the exposure (Shuba et al., 1978). For 

example, in static whole-sediment tests, the alkalinity, 

hardness, and conductivity of overlying water more than 

doubled in several treatments during a four-week exposure (Ingersoll and Nelson, 1990). Additionally, concentrations of metabolic products (e.g., ammonia) may also 

increase during static exposures, and these compounds 

can either be directly toxic to the test organisms or may 

52 
Table 12.2 General Activity Schedule for Conducting a Sediment Toxicity Test with Chifonomus tenfans 1 

Day Activity 

-14 Isolate adults for production of egg masses. 

-13 Place newly deposited egg masses Into hatchmg dishes. 

-12 A larval reanng chamber IS prepared with new substrate. 

-11 Examtne egg masses for hatchrng success. If egg masses have hatched, transfer first instar larvae and any renaming unhatched 

embryos from the crystallizing dishes Into the larval rearrng chamber. Feed orgamsms. 

-10 SameasDay-11. 

-9 to -2 Feea and observe midges. Measure water quality (e.g., temperature and dissolved oxygen). 

-1 Add Iood to each larval rearing chamber and measure temperature and dissolved oxygen. Add sediment Into each test chamber, 

place chamber into exposure system, and start renewing overlying water. 

0 Measure total water quality (temperature, pH. hardness, alkalinity, dissolved oxygen, conductMy, ammonia). Remove third-mstar 

larvae from the culture chamber substrate. Add 1.5 mL of Tetrafin@ (4.0 g/L) into each test chamber. Transfer 10 larvae into each 

test chamber. Release organisms under the surface of the water. Archive 20 test organisms for instar determination and weight or 

length determmation. Observe behavior of test orgamsms. 

1 to 8 Add 1.5 mL of food to each test chamber. Measure temperature and dissolved oxygen. Observe behavior of test organisms. 

9 Same as Day 1. Measure total water quallty. 

10 Measure temperature and dissolved oxygen. End the test by collecting the rnldges with a sieve. Measure weight or length of surviving 

larvae. 

’ Modtfred from Call et al.. 1994 

contribute to the toxicity of the contaminants in the 

sediment. Furthermore, changes in water-quality characteristics such as hardness may influence the toxicity 

of many inorganic (Gauss et al., 1985) and organic 

(Mayer and Ellersieck, 1986) contaminants. Although 

contaminant concentrations are reduced in the overlying 

water in water-renewal tests, organisms in direct contact 

with sediment generally receive a substantial proportion 

of a contaminant dose directly from either the whole 

sediment or from the interstitial water. 

12.3.3 Acclimation: Test organisms must be cultured 

and tested at 23°C. Ideally, test organisms should be 

cultured in the same water that will be used in testing. 

However, acclimation of test organisms to the test water 

is not required. 

12.3.4 Piacing Organisms in Test Chambers: Test 

organisms should be handled as little as possible. Midges 

should be introduced into the overlying water below the 

air-water interface. Test organisms can be pipetted directly into overlying water (Ankley et al., 1993). Alternatively, test organisms can be placed into 30-mL counting 

cups that are floated in the test chambers for 15 min 

before organisms are introduced into the overlying water 

(Ingersoll and Nelson, 1990). Length or weight should 

be measured on a subset of at least 20 organisms used 

to start the test. Head capsule width of midges must be 

measured on this subset of test organisms to determine 

the instar used to start the test (Table 10.2). 

12.3.5 Monitoring a Test: All chambers should be 

checked daily and observations made to assess test 

organism behavior such as sediment avoidance. However, monitoring effects on burrowing activity of test 

organisms may be difficult because the test organisms 

are often not visible during the exposure. The operation 

of the exposure system should be monitored daily. 

12.3.5.1 Measurement of Overlying Water-quality Characteristics: Conductivity, hardness, pH, alkalinity, and 

ammonia should be measured in all treatments at the 

beginning and end of a test. Overlying water should be 

sampled just before water renewal from about 1 to 2 cm 

above the sediment surface using a pipet. It may be 

necessary to pool water samples from individual replicates. The pipet should be checked to make sure no 

organisms are removed during sampling of overlying 

water. Hardness, alkalinity, pH, conductivity, and ammonia in the overlying water within a treatment should not 

vary by more than 50% during a test. 

12.3.5.1 .l Dissolved oxygen should be measured daily 

and should be between 40 and 100% saturation (ASTM, 

1988a). If a probe is used to measure dissolved oxygen 

in overlying water, it should be thoroughly inspected 

between samples to make sure that organisms are not 

attached and should be rinsed between samples to 

minimize cross contamination. Aeration can be used to 

maintain dissolved oxygen in the overlying water above 

40% saturation. Dissolved oxygen and pH can be measured directly in the overlying water with a probe. 

12.3.5.1.2 Temperature should be measured at least 

daily in at least one test chamber from each treatment. 

The temperature of the water bath or the exposure 

53 
Table 12.3 Test Acceptability Requirements for a 10-d Sediment Toxicity Test with Chironomus tentans 

A. It IS recommended for conductmg a 1 O-d test with C. fentans that the following performance cntena be met: 

1. Tests must be started with third-mstar and younger larvae. At least 50% of the larvae must be in the third mstar at the start of the test 

2. Average survival of C. tentans in the control sediment must be greater than or equal to 70% at the end of the test. 

3. Average size of C. fentans In the control sediment must be at least 0.6 mg at the end of the test. 

4. Hardness, alkallnlty, pH, and ammonia In the overlying water wlthln a treatment should nol vary by more than 50% during the test. 

9. Performance-based criteria for culturmg C. tentans Include 

1. Laboratones should perform monthly 96-h water-only reference-toxlclty tests to assess the sensitivity of culture organisms. If reference-toxicity tests are not conducted monthly, the lot of organisms used to start a sediment test must be evaluated using a reterence 

toxlcant (Section 9.16). 

2. Laboratones should keep a record of t:me to first emergence for each culture and record this InformatIon using control charts. Records 

should also be kept on the frequency of restarting cultures. 

3. Laboratories should record the following waterquallty charactensbcs of the cultures at least quarterly and the day before the start of a 

sediment test: pH, hardness, alkahmty, and ammonia. Dissolved oxygen should be measured weekly. Temperature should be 

recorded dally. 

4. Laboratones should charactenze and monitor background contam~natlon and nutrient quality of food if problems are observed in 

culturing or testing organisms. 

5. Physlologlcal measurements such as lipid content might provide useful information regarding the health of the cultures. 

C. Additionat requirements: 

1. All organisms in a test must be from the same source. 

2. It is desirable to start tests soon after collection of sediment from the field (see Sectlon 8.2 for additional detail) 

3. All test chambers (and compartments) should be identical and should contain the same amount of sediment and overlylng water, 

4. Negative-control sediment and appropriate solvent controls must be included in a test. The concentration 01 solvent used must not 

adversely affect test orgamsms. 

5. Test orgamsms must be cultured and tested at 23°C 

6. The daily mean test temperature must be within tl”C of the desired temperature. The instantaneous temperature must always be 

within +3”C of the desired temperature. 

7. Natural physico-chemical charactenstlcs of test sediment collected from the field should be wlthin the tolerance Ilmlts of the test 

organisms. 

chamber should be continuously monitored. The daily 

mean test temperature must be within +l% of the 

desired temperature. The instantaneous temperature 

must always be within 53°C of the desired temperature. 

12.3.6 feeding: Without addition of food, the test 

organisms may starve during exposures. However, the 

addition of the food may alter the availability of the 

contaminants in the sediment (Wiederholm et al., 1987; 

Harkey et al., 1994). Furthermore, if too much food is 

added to the test chamber or if the mortality of test 

organisms is high, fungal or bacterial growth may develop on the sediment surface. Therefore, the amount of 

food added to the test chambers is kept to a minimum. 

12.3.6.1 Suspensions of food should be thoroughly 

mixed before aliquots are taken. If excess food collects 

on the sediment, a fungal or bacterial growth may develop on the sediment surface, in which case feeding 

should be suspended for one or more days. A drop in 

dissolved oxygen below 40% of saturation during a test 

may indicate that the food added is not being consumed. 

Feeding should be suspended for the amount of time 

necessary to increase the dissolved oxygen concentration (ASTM, 1994a). If feeding is suspended in one 

treatment, it should be suspended in all treatments. 

Detailed records of feeding rates and the appearance of 

the sediment surface should be made daily. 

12.3.7 Ending a Test: Immobile organisms isolated 

from the sediment surface or from sieved material should 

be considered dead. Ankley et al. (1994a) recommend 

using a #25 sieve (710 m mesh) to remove midges 

from sediment. Alternatively, Kemble et al. (1994) recommend sieving sediment using the following procedure: (1) pour about half of the overlying water through a 

#50 (300 pm) U.S. Standard mesh sieve, (2) pour about 

half of the sediment through the #50 mesh sieve and 

wash the contents of the sieve into an examination pan, 

(3) rinse the coarser sediment remaining in the test 

54 
chamber through a #40 (425 pm) mesh sieve and wash 

the contents of this second sieve into a second examination pan. Surviving midges can then be isolated from 

these pans. See Section 12.3.8.1 and 12.3.8.2 for the 

procedures for measuring weight or length of midges. 

12.3.7.1 A consistent amount of time should be taken to 

examine sieved material for recovery of test organisms 

(e.g., 10 minlreplicate). Laboratories should demonstrate that their personnel are able to recover an average of at least 90% of the organisms from whole sediment. For example, test organisms could be added to 

control sediment and recovery could be determined 

after 1 h (Tomasovic et al., 1994). 

12.3.8 Test Data: Dry weight and survival are the 

endpoints measured at the end of the 10-d sediment 

toxicity test with C. lentans. The duration of the 10-d test 

starting with third instar larvae is not long enough to 

determine emergence of adults. Average size of C. 

tentans in the control sediment must be at least 0.6 mg 

at the end of the test (Ankley et al., 1993; ASTM, 1994b; 

Section 15). 

12.3.8.1 Head capsule width can be measured on 

surviving midges at the end of the test before dry weight 

is determined. Dry weight of midges should be determined by pooling all living larvae from a replicate and 

drying the sample at about 60 to 90°C to a constant 

weight. The sample is brought to room temperature in a 

desiccator and weighed to the nearest 0.01 mg to obtain 

mean weights per surviving organism per replicate. Pupae or adult organisms must not be included in the 

sample to estimate dry weight. 

12.3.8.2 Measurement of length is optional. Separate 

replicate beakers should be set up to sample lengths of 

midges at the end of an exposure. An 8% sugar formalin 

solution can be used to preserve samples for length 

measurements (Ingersoll and Nelson, 1990). NoTox@ 

(Earth Safe Industries, Belle Mead, NJ) can be used as 

a substitute for formalin (Unger et al., 1993). Midge body 

length (kO.1 mm) can be measured from the anterior of 

the labrum to the posterior of the last abdominal segment (Smock, 1980). Kemble et al. (1994) photographed 

midges at magnification of 3.5x and measured the images using a computer-interfaced digitizing tablet. A 

digitizing system and microscope can also be used to 

measure length (Ingersoll and Nelson, 1990). 

12.4 Interpretation of Results 

12.4.1 Section 14 describes general information for 

interpretation of test results. The following sections describe species-specific information that is useful in helping to interpret the results of sediment toxicity tests with 

C. tentans. 

12.4.2 Age Sensitivity: Midges are perceived to be 

relatively insensitive organisms in toxicity assessments 

(Ingersoll, 1994). This conclusion is based on the practice of conducting short-term tests with 4th instar larvae 

in water-only exposures, a procedure that may underestimate the sensitivity of midges to toxicants. The first and 

second instars of chironomids are more sensitive to 

contaminants than the third or fourth instars. For example, first instar C. lentans larvae were 6 to 27 times 

more sensitive than 4th instar larvae to acute copper 

exposure (Nebeker et al., 198413; Gauss et al., 1985; 

Figure 12.1) and first instar C. rioarius larvae were 127 

times more sensitive than second instar larvae to acute 

cadmium exposure (Williams et al., 1986b; Figure 12.1). 

In chronic tests with first instar larvae, midges were 

often as sensitive as daphnids to inorganic and organic 

compounds (Ingersoll et al., 1990). Sediment tests should 

be started with uniform age and size midges because of 

the dramatic differences in sensitivity of midges by age. 

While third instar midges are not as sensitive as younger 

organisms, the larger larvae are easier to handle and 

isolate from sediment at the end of a test. 

12.4.3 Grain Size: Chironomus fentans are tolerant of a 

wide range of substrates. Physicochemical characteristics (e.g., grain size or TOC) of sediment were not 

significantly correlated to the survival of C. tentans in 

toxicity tests in which organisms were fed. However, 

linear modeling indicated that growth of C. tentans may 

have been slightly influenced by grain size distribution of 

the test sediments (Section 10.1.2.3; Ankley et al., 

1994a). Survival of C. tenfans was reduced below 0.91% 

organic matter in 10-d tests with formulated sediment 

(Suede1 and Rodgers, 1994); however these organisms 

did not receive a supplemental source of nutrition. 

12.4.4 isolating Organisms at the End of a Test: 

Quantitative recovery of larvae at the end of a 10-d 

sediment test should not be a problem. The larvae are 

red and typically greater than 5-mm long. 

12.4.5 influence of indigenous Organisms: The influence of indigenous organisms on the response of C. 

tentans in sediment tests has not been reported. Survival of a closely related species, C. riparius was not 

reduced in the presence of oligochaetes in sediment 

samples (Reynoldson et al., 1994). However, growth of 

C. riparius was reduced when high numbers of oligochaetes were placed in a sample. Therefore, it is important 

to determine the number and biomass of indigenous 

organisms in field-collected sediment in order to better 

interpret growth data (Reynoldson et al., 1994). Furthermore, presence of predators may also influence the 

response of test organisms in sediment (Ingersoll and 

Nelson, 1990). 

55 
A. Chironomus riparius: Cadium 

2 

0.5 

2nd 3rd 4th 

INSTAK 

Williams et al. (1966) 

8. Cbironomus tentans: Copper 

4th 

INSTAR 

Nebeker et al. (1964) 

Figure 12.1 Lifestage sensitivity of chironomids. 

56 
Section 13 

Test Method 100.3 

Lumbriculus variegatus Bioaccumulation Test for Sediments 

13.1 Introduction 

13.1.1 Lumbriculus variegatus (Oligochaeta) have many 

desirable characteristics of an ideal sediment bioaccumulation testing organism including contact with sediment, ease of culture in the laboratory, and tolerance to 

varying physico-chemical characteristics of sediment. 

The response of L. variegatus in laboratory exposures 

has been confirmed with natural benthos populations. 

Many investigators have successfully used L. variegatus 

in toxicity or bioaccumulation tests. Toxicity studies have 

been conducted in water-only tests (Bailey and Liu, 

1980; Hornig, 1980; Ewell et al., 1986; Nebeker et al., 

1989; Ankley et al., 1991a, Ankley et al., 1991b), in 

effluent tests (Hornig, 1980), and in whole-sediment 

tests (Nebeker et al., 1989; Ankley et al., 1991a, Ankley 

et al., 1991b; Ankley et al., 1992a; Call et al., 1991; 

Carlson et al., 1991; Phipps et al., 1993; West et al., 

1993). Several studies have reported the use of L. 

variegatus to examine bioaccumulation of chemicals 

from sediment (Schuytema et al., 1988; Nebeker et al., 

1989; Ankley et al., 1991b; Call et al., 1991; Carlson et 

al., 1991; Ankley et al., 1993; Kukkonen and Landrum, 

1994; and E.L. Brunson, NBS, Columbia, MO, unpublished data). However, interlaboratory studies have not 

yet been conducted with L. variegatus. 

13.1.2 Additional research is needed on the standardization of bioaccumulation procedures with sediment. 

Therefore, Section 13.2 describes general guidance for 

conducting a 28-d sediment bioaccumulation test with L. 

variegatus. Methods outlined in Appendix A and the 

literature cited in Table A.4 were used for developing 

this general guidance. Results of tests using procedures 

different from the procedures described in Section 13.2 

may not be comparable, and these different procedures 

may alter bioavailability. Comparison of results obtained 

using modified versions of these procedures might provide useful information concerning new concepts and 

procedures for conducting sediment tests with aquatic 

organisms. If tests are conducted with procedures different from the procedures described in this manual, additional tests are required to determine comparability of 

results (Section 1.3). 

13.2 Procedure for Conducting 

Sediment Bioaccumulation Tests 

with Lumbriculus variegatus 

13.2.1 Recommended test conditions for conducting a 

28-d sediment bioaccumulation test with L. variegatus 

are summarized in Table 13.1. Table 13.2 outlines procedures for conducting sediment toxicity tests with L. 

variegatus. A general activity schedule is outlined in 

Table 13.3. Decisions concerning the various aspects of 

experimental design, such as the number of treatments, 

number of test chambers/treatment, and water-quality 

characteristics should be based on the purpose of the 

test and the methods of data analysis (Section 14). The 

number of replicates and concentrations tested depends 

in par-t on the significance level selected and the type of 

statistical analysis. When variability remains constant, 

the sensitivity of a test increases as the number of 

replicates increases. 

13.2.2 The recommended 28-d sediment bioaccumulation test with L. variegatus can be conducted with adult 

oligochaetes at 23°C with a 16L:8D photoperiod at a 

illuminance of about 500 to 1000 Iux (Table 13.1). Test 

chambers can be 4 to 6 L that contain 1 to 2 L of 

sediment and 1 to 4 L of overlying water. The number of 

replicates/treatment depends on the objective of the 

test. Five replicates are recommended for routine testing (Section 14). To minimize depletion of sediment 

contaminants, the ratio of total organic carbon in sediment to dry weight of organisms should be about 50:1. A 

minimum of 1 g/replicate with up to 5 g/replicate should 

be tested. Oligochaetes are not fed during the test. Each 

chamber receives 2 volume additions/d of overlying 

water. Appendix B describes water-renewal systems 

that with minor modifications can be used to deliver 

overlying water. Overlying water can be culture water, 

well water, surface water, site water, or reconstituted 

water. For site-specific evaluations, the characteristics 

of the overlying water should be as similar as possible to 

the site where sediment is collected. Requirements for 

test acceptability are outlined in Table 13.4. 

13.2.2.1 Before starting a 28-d sediment bioaccumulation test with L. variegatus, a toxicity screening test 

should be conducted for at least 4 d using procedures 

outlined in Table 13.2 (E.L. Brunson, NBS, Columbia, 

57 
Table 13.1 Recommended Test Conditions for Conducting a 28-d Sediment Bioaccumulation Test with Lumbriculus variegatus 

Parameter Conditions 

1. Test type: Whole-sediment bioaccumulation test with renewal of overlying water 

2. Temperature: 23 ± 1°C 

3. Light quality: Wide-spectrum fluorescent lights 

4. Illuminance: About 500 to 1000 lux 

5. Photoperiod: 16L:8D 

6. Test chamber: 4- to 6-L aquaria with stainless steel screens or glass standpipes 

7. Sediment volume: 1 L or more depending on TOC 

8. Overlying water volume: 1 L or more depending on TOC 

9. Renewal of overlying water: 2 volume additions/d; continuous or intermittent (e.g., one volume addition every 12 h) 

10. Cl. Age of test organisms: Adults 

11. Loading of organisms Ratio of total organic carbon in sediin chamber: ment to organism dry weight should be no less than 50:1. Minimum of 1 g/replicate. Preferably 5 g/replicate 

12. Number of replicate Depends on the objective of the 

chambers/treatment: test. Five replicates are recommended for routine testing (see Section 14) 

13. Feeding: None 

14. Aeration: None, unless dissolved oxygen in overlying water drops below 40% of saturation 

15. Overlying water: Culture water, well water, surface water, site water, or reconstituted water 

16. Test chamber cleaning: It screens become clogged during the test, gently brush the outside of the screen (Appendix B) 

17. Overtying water quality: Hardness, alkalinity, conductivity, pH. and ammonia at the beginning and end of a test. Temperature and 

dissolved oxygen daily 

18. Test duration: 28d 

19. Endpoint: Bioaccumulation 

20. Test acceptability: Performance-based criteria specifications outlined in Table 13.4. 

MO, unpublished data). The preliminary toxicity screening test is conducted at 23°C with a 16L:8D photoperiod 

at a illuminance of about 500 to 1000 lux. Test chambers 

are 3OO-mL high-form lipless beakers containing 100 mL 

of sediment and 175 mL of overlying water. Ten adult 

oligochaetes/replicate are used to start a test. Four 

replicates are recommended for routine testing. Oligochaetes are not fed during the test. Each chamber 

receives 2 volume additions/d of overlying water. Ap 

pendix B describes water-renewal systems that can be 

used to deliver overlying water. Overlying water should 

be similar to the water to be used in the bioaccumulation 

test. Endpoints monitored at the end of a toxicity test are 

number of organisms and behavior. Numbers of L. 

variegatus in the toxicity screening test should not be 

significantly reduced in the test sediment relative to the 

control sediment. Test organisms should burrow into 

test sediment. Avoidance of test sediment by L variegatus 

may decrease bioaccumulation. 

13.3 General Procedures 

13.3.1 Sediment into Test Chambers: The day before 

the sediment test is started (Day -1) each sediment 

should be thoroughly mixed and added to the test chambers (Section 8.3.1). Sediment should be visually inspected to judge the extent of homogeneity. Excess 

water on the surface of the sediment can indicate separation of solid and liquid components. If a quantitative 

measure of homogeneity is required, replicate subsamples should be taken from the sediment batch and 

analyzed for TOG chemical concentrations, and particle 

size. 

13.3.1.1 Each test chamber should contain the same 

amount of sediment, determined either by volume or by 

weight. Overlying water is added to the chambers in a 

manner that minimizes suspension of sediment. This 

can be accomplished by gently pouring water along the 

sides of the chambers or by pouring water onto a baffle 

(e.g., a circular piece of Teflon@ with a handle attached) 

placed above the sediment to dissipate the force of the 

water. Renewal of overlying water is started on Day -1. 

A test begins when the organisms are added to the test 

chambers (Day 0). 

13.3.2 Renewal of Overlying Water: Renewal of overlying water is recommended during a test. At any particular time during the test, flow rates through any two 

test chambers should not differ by more than 10%. 

Mount and Brungs (1967) diluters have been modified 

for sediment testing, and other automated water delivery 

systems have also been used (Maki, 1977; Ingersoll and 

Nelson, 1990; Benoit et al.. 1993; Zumwalt et al., 1994). 

Each water-delivery system should be calibrated before 

a test is started to verify that the system is functioning 

properly. Renewal of overlying water is started on Day 
Table 13.2 Recotnnrended Test Conditions for Conducting a Preliminary 4-d Sedhnent Toxicity Screenhg Test with Lumbnkulus 

variegahrs 

Parameter Conditions 

1. Test type: 4-d Whole-sediment toxicity test with renewal of overlying water 

2. Temperature: 23 f “C 

3. Light quality: Wide-spectrum fluorescent lights 

4. Illuminance: about 500 to loo0 lux 

5. Photopenod: 16L:BD 

6. Test chamber: 300-mL high-form lipless beaker 

7. Sediment volume: 1OOmL 

8. Overlying water volume: 175 mL 

9. Renewal of overlyrng water: 2 volume additions/d; continuous or intermittent (e.g., one volume addition every 12 h) 

10. Age of test organisms: Adults 

11. Number of 10 

organisms/chamber: 

12. Number of replicate 

chambers/treatment: 

t3. Feeding: 

14. Aeration: 

15. OverlyIng water: 

16. Test chamber cleaning: 

17. Overlymg water quality: 

16. Test duratron: 

19. Endpoints: 

20. Test acceptability: 

4 minlmum 

None 

None, unless dissolved oxygen in overlymg water drops below 40% of saturation 

Culture water, well water. surfa water, site water, or reconstituted water 

If screens become clogged during the test. gently brush the outside of the screen 

Hardness, alkalinity, conductivity. pH, and ammonia at the beginning and end of a test. Temperature and 

dissolved oxygen daily 

4 d (minimum; up to 10 d) 

Number of organisms and behavior. There should be no significant reduction in number of orgamsms in a test 

sediment relative to the control 

Performance-based criteria spcifcations outlined in Table 13.4 

-1 before the addition of test organisms or food on Day 0 

(Appendix B). 

13.3.2.1 In water-renewal tests with one to four volume 

additions of overlying water/d, waterquality characteristics generally remain similar to the inflowing water 

(Ingersoll and Nelson, 1990; Ankley et al., 1993); however, in static tests, water quality may change pr* 

foundly during the exposure (Shuba et al., 1978). For 

example, in static whole-sediment tests, the alkalinity, 

hardness, and conductivity of overlying water more than 

doubled in several treatments during a four-week exp@ 

sure (Ingersoll and Nelson, 1990). Additionally, concentrations of metabolic products (e.g., ammonia) may also 

increase during static exposures, and these compounds 

can either be directly toxic to the test organisms or may 

contribute to the toxicity of the contaminants in the 

sediment. Furthermore, changes in waterquality characteristics such as hardness may influence the toxicity 

of many inorganic (Gauss et al., 1985) and organic 

(Mayer and Ellersieck, 1986) contaminants. Although 

contaminant concentrations are reduced in the overlying 

water in water-renewal tests, organisms in direct contact 

with sediment generally receive a substantial proportion 

of a contaminant dose directly from either the whole 

sediment or from the interstitial water. 

13.3.3 Acclimafion: Test organisms must be cultured 

and tested at 23°C. Ideally, test organisms should be 

cultured in the same water that will be used in testing. 

However, acclimation of test organisms to the test water 

is not required. 

13.3.4 Placing Org8nisms in Test Cnambersr Isolate 

oligochaetes for starting a test as described in Section 

10.5.6.. A subset of L. variegatus at the start the test 

should be sampled to determine starting concentrations 

of contaminants of concern. Mean group weights should 

be measured on a subset of at least 100 organisms 

used to start the test. The ratio of total organic carbon in 

sediment to dry weight of organisms at the start of the 

test should be no less than 50: 1. 

13.3.4.1 Oligochaetes added to each replicate should 

not be blotted to remove excess water (Section 10.56). 

Oligochaetes can be added to each replicate at about 

1.33x of the target stocking weight (E-L. Brunson, NBS, 

Columbia, MO, unpublished data). This additional 33% 

should account for the excess weight from water in the 

sample of nonblotted oligochaetes at the start of the 

test. 

59 
Table 13.3 General Activity Schedule for Conducting a 28-d Sediment Bioaccumulation Test with Lumbricutus varkgatus 

A. Conductmg a 4-d Toxrc~ty Scrcemng Test (conducted before the 28-d broaccumulatron test) 

W Activity 

-1 Isolate worms for conducting toxrcity screenrng test. Add sediment Into each test chamber, place chambers Into exposure system, 

and star1 renewrng overlying water. 

0 Measure total water quality (pH, temperature, drssolved onygen, hardness, alkakmty, conductrvrty, ammonia). Transter 10 worms 

into each test chamber. Measure weight of a subset of 20 organisms used to start the test. Observe behavtor of test orgamsms. 

l-2 Measure temperature and drssolved oxygen. Observe behavror of test organisms. 

3 Same as Day 1. Measure total water quality. 

4 Measure temperature and dissolved oxygen. End the test by collectmg the oligochaetes with a sieve and determme weight of 

survrvors. Broaccumulatron tests should not be conducted with L. variegafus 11 a test sediment slgnrficantly reduces number of 

olrgochaetes relatrve to the control sediment or if oligochaetes avoid the sediment. 

6. Conductmg a 28-d Broaccumulatron Test 

Day Activrty 

-1 Isolate worms for conductrng bioaccumulation test. Add sediment into each test chamber, place chambers into exposure system, 

and start renewing overlying water. 

0 Measure total water quality (pH, temperature, dissolved oxygen, hardness, alkalinity, conductrvrty, ammonra). Transfer appropriate 

amount of worms (based on weight) into each test chamber. Sample a subset of worms used to start the test for residue analyses. 

Observe behavror of test organisms. 

l-6 Measure temperature and dissolved oxygen. Observe behavior of test organisms. 

7 Same as Day 1. Measure total water quality. 

8-13 Same as Day 1 

14 Same as Day 7 

15-20 Same as Day 1 

21 Same as Day 7 

22-26 Same as Day 1 

27 Same as Day 1. Measure total water quality. 

28 Measure temperature and drssolvedoxygen. End uptake by collectingthe worms with a sieve. Separate any mdrgenous organisms 

from L. variegarus. Determine weight of survivors. Eliminate gut contents of survivtng worms in water for 24 h. 

29 Sample surviving worms after 24 h ot elimination for chemical analysis. 

13.35 Monitoring a Test: All chambers should be 

checked daily and observations made to assess test 

organism behavior such as sediment avoidance. However, monitoring effects on burrowing activity of test 

organisms may be difficult because the test organisms 

are often not visible during the exposure. The operation 

of the exposure system should be monitored daily. 

13.351 Measurement of Overlying Water-quality 

Characteristics: Conductivity, hardness, pH, alkalinity, 

and ammonia should be measured in all treatments at 

the beginning and end of a test. Overlying water should 

be sampled just before water renewal from about 1 to 2 

cm above the sediment surface using a pipet. It may be 

necessary to pcol water samples from individual replicates. The pipet should be checked to make sure no 

organisms are removed during sampling of overlying 

water. Hardness, alkalinity, pH, conductivity, and ammonia in the overlying water within a treatment should not 

vary by more than 500/o during a test. 

13.3.5.1 .l Dissolved oxygen should be measured 

daily and should be between 40 and 100% saturation 

(ASTM, 1988a). If a probe is used to measure dissolved oxygen in overlying water, it should be thoroughly inspected between samples to make sure that 

organisms are not attached and should be rinsed between samples to minimize cross contamination. Aeration can be used to maintain dissolved oxygen in the 

overlying water above 40% saturation. Dissolved oxygen and pH can be measured directly in the overlying 

water with a probe. 

13.3.5.1.2 Temperature should be measured at least 

daily in at least one test chamber from each treatment. 

The temperature of the water bath or the exposure 

chamber should be continuously monitored. The daily 

mean test temperature must be within ?l”C of the 

desired temperature. The instantaneous temperature 

must always be within +3”C of the desired temperature. 

60 
Table 13.4 Test Acceptability Requirements for a 28-d Sediment Bioaccumulation lest with lumbriculus vtwfepfus 

A. It is recommended for conducting a 28-d test with L. variegatus that the following performance criteria be met: 

1. Numbers of L. warregalus in a 4-d toxicity screening test should not be significantly reduced in the test sediment relative to the control 

sediment. 

2. Test orgamsms should burrow Into test sediment. Avoidance of test sedrment by L. wariegafus may decrease bioaccumulatlon. 

3. Hardness, alkalmity. pH, and ammonia in the overlymg water within a treatment should not vary by more than 50% dunng the test. 

0. Performance-based criteria for culturing L. variegatus include: 

1. Laboratones should perform monthly 96-h water-only reference-toxicity tests to assess the sensittvity of culture organisms. If 

reference-toxicity tests are not conducted monthly, the lot of organisms used to start a sediment test must be evaluated using a 

reference toxicant (Section 9.16). 

2. Laboratones should monitor the frequency with which the population is doubling in the culture (number of organisms) and record thts 

informatlon using control charts (doubling rate would need to be estimated on a subset of ammals from a mass culture). Records 

should also be kept on the frequency of restarting cultures. 

3. Food used to culture organisms should be analyzed before the start of a test for compounds to be evaluated in the bloaccumulation 

test. 

4. laboratones should record the following water-quality characteristics of the cultures at least quarterly and the day before the start of 

a sediment test: pl-l, hardness, alkalinity, and ammonia. Dissolved oxygen should be measured weekly. Temperature should be 

recorded daily. 

5. Laboratones should characterize and monitor background contamination and nutrient quality of food if problems are observed in 

culturing or testing organisms. 

6. Physiological measurements such as lipid content might provide useful information regarding the health of the cultures. 

C. AdditIonal requirements: 

1. All organisms in a test must be from the same source. 

2. It is desirable to start tests soon after collection of sediment from the field (see Section 8.2 for additional detail). 

3. All test chambers (and compartments) should be identical and should contain the same amount of sediment and overlying water. 

4. Negative-control sediment and appropriate solvent controls must be included in a test. The concentration of solvent used must not 

adversely affect test organisms. 

5. Test organisms must be cultured and tested at 23°C. 

6. The daily mean test temperature must be within +l”C of the desired temperature. The instantaneous temperature must always be 

within +3”C of the desired temperature. 

7. Natural physico-chemical characteristics of test sediment collected from the field should be within the tolerance limits of the test 

organisms. 

13.3.6 Feeding: Lumbriculus variegatus should not be 

fed during a bioaccumulation test. 

13.3.7 En&g B Tesl Sediment at the end of the test 

can be sieved through a fine-meshed screen sufficiently 

small to retain the oligochaetes (e.g., U.S. Standard 

Sieve 35 (500 pm mesh) or 40 (425 pm mesh)). The 

sieved material should be quickly transferred to a shallow pan to keep oligochaetes from moving through the 

screen. Immobile organisms should be considered dead. 

Live oligochaetes are transferred to a 1-L beaker containing overlying water without sediment for 24 h to 

eliminate gut contents. Oligochaetes should not be placed 

in clean sediment to eliminate gut contents. Clean sediment can add 15 to 20% to the dry weight of the 

oligochaetes which would result in a dilution of contaminant concentrations on a dry weight basis (Peter 

Landrum, NOAA, Ann Arbor, MI, personal communication). The elimination beakers may need to be aerated to 

maintain dissolved oxygen above 40% of saturation. 

Worms will clear more than 90% of the gut contents in 

24 h (Call et al., 1991). Following the 24-h elimination 

period, oligochaetes should be collected, placed in a 

tared weigh boat, blotted to remove excess water, and 

weighed to determine wet weight. Each sample should 

then be split into appropriate aliquots (e.g., metals, 

organics), placed in clean containers, and frozen for 

later analysis. Containers should be placed inside secondary freezer containers to minimize “freezer burn” or 

dehydration during storage. 

13.3.7.1 Fieldcollected sediments may include indigenous oligochaetes. The behavior and appearance of 

indigenous oligochaetes are usually different from L. 

variegatus. It may be desirable to test extra chambers 

without the addition of L. variegatus to check for the 

presence of indigenous oligochaetes in field-collected 

sediment (Phipps et al., 1993). Bioaccumulation of con61 
Table C3.5 Grams of Lumbrkutus variegatvs Tissue (Wet 

Weight) Required for Various Analytes at Selected 

Lower Llmlts oi Detection 

Analyte 

Grams of Tissue 

1.0 2.0 5.0 

Lower Ltm~t of Detection (pg/g) 

PC& 

PCB (totai’) 

PCB (congene+) 

Level of chlorination 

mono-tnchloro 

tetra-hexachloro 

hepta-octachloro 

nona-decachloro 

Organochlorine pest&es 1 

P.P’ DDE 

p.p’ - ODD 

p,p’ - DDT 

o,p’ - DDE 

0,p’ ODD 

o,p’ DOT 

Alpha-chlordane 

Gamma-chfordane 

Dieldrin 

Endrin 

Heptachlorepoxlde 

Oxychlordane 

Mirex 

Trans - nonachlor 

Toxaphene 

PAM 3 

PAHs 

Diox,ns ’ 

TCDO 

Inorganic 4 

Cadmium 

Copper 

Lead 

Ztnc 

0.600 0.300 0.120 

0.025 0.0125 0.005 

0.050 0.025 0.010 

0.075 0.0375 0.015 

0.125 0.0625 0.025 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.05a 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.050 0.025 0.010 

0.600 0.300 0.120 

0.012 0.006 0.002 

0.020 

Wg) 

0.004 

(Wg) 

0.005 0.0025 0.001 

0.005 0.0025 0.001 

0.005 0.0025 0.001 

0.005 0.0025 0.00 1 

’ Schmitt et al., 1990 

2 USEPA. 19% 

3 Vassllaros et al., 1982 

’ USEPA. 199Cd 

5 Schmitt and Finger, 1967 

Table 13.6 DetecSOn Limlts (ng) of Individual PAHs by HPLG 

FD’ 

Analyle 

Benzo(a)pyrene 

Pyrene 

Benzo(k)fluoranthene 

Dbenz(a,h)anthracene 

Anthracene 

Benz(a)anthracene 

Benzo(e)pyrene 

Benzo(b)fluoranthene 

Benzo(g,h.l)perylene 

3-Methyleholanthrene 

’ Obana et al., 1961 

Detectron L,mlt (ng) 

0.01 

0.03 

0.03 

0.03 

0.10 

o.to 

0.10 

0.10 

0.10 

0.10 

laminants by indigenous oligochaetes exposed in the 

same chamber with introduced L. variegates in a 28-d 

test has been evaluated (E.L. Bruson, NBS, Columbia, 

MO, personal communication). Peak concentrations of 

select PAHs and DOT were similar in the indigenous 

oligochaetes and L. variegalus exposed in the same 

chamber for 28 d. 

13.3.7.2 Care should be taken to isolate at least the 

minimum amount of tissue mass from each replicate 

chamber needed for analytical chemistry. 

13.3.8 Test Data: Tissue masses required for various 

analyses at selected lower limits of detection are listed 

in Table 13.5. Detection limits for individual PAHs in 

tissue are listed in Table 13.6. A minimum of 1 g/ 

replicate and preferably 5 g/replicate should be tested. 

13.3.8.1 If an estimate of dry weight is needed, a 

subsample should be dried to a constant weight at about 

60 to 90°C. The sample is brought to room temperature 

in a desiccator and weighed to the nearest 0.01 mg. 

Lumbriculus vafiegatus typically contain about 1% lipid 

(dry weight). 

13.3.8.2 Depending on specific study objectives, total 

lipids can be measured on a subsample of the total 

tissue mass of each thawed replicate sample. Gardner 

et al. (1985) describe procedures for measuring lipids in 

1 mg of tissue. Different methods of lipid analysis can 

yield different results (Randall et al., 1991). The analytical method used for lipid analysis should be calibrated 

against the chloroform-methanol extraction method described by Folch et al. (1957) and Bligh and Dyer (1959). 

13.3.8.2.1 A number of studies have demonstrated that 

tipids are the major storage site for organic contaminants in a variety of organisms (Robens et al., 1977; 

62 
Oliver and Niimi, 1983; de Boer, 1988). Because of the 

importance of lipids, it may be desirable to normalize 

bioaccumulated concentrations of nonpolar organics to 

the tissue lipid concentration. Lipid concentration is one 

of the factors required in deriving the BSAF (Section 14). 

However, the difficulty with using this approach is that 

each lipid method generates different lipid concentrations (see Kates (1986) for discussion of lipid methodology). The differences in lipid concentrations directly 

translate to a similar variation in the lipid-normalized 

contaminant concentrations or BSAF. 

13.3.8.2.2 For comparison of lipid-normalized tissue 

residues or BASFs, it is necessary to either promulgate 

a standard lipid technique or to intercalibrate the various 

techniques. Standardization of a single method is difficult because the lipid methodology is often intimately 

tied in with the extraction procedure for contaminant 

analysis. As an interim solution, the Bligh-Dyer lipid 

method (Bligh and Dyer, 1959) is recommended as a 

temporary “intercalibration standard” (Lee et al., 1994). 

13.3.8.2.3 The potential advantages of Bligh-Dyer include its ability to extract neutral lipids not extracted by 

many other solvent systems and the wide use of this 

method (or the same solvent system) in biological and 

toxicological studies (e.g., Roberts et al., 1977; Oliver 

and Niimi, 1983; de Boer, 1988; Landrum, 1989). Because the technique is independent of any particular 

analytical extraction procedure, it will not change when 

the extraction technique is changed. Additionally, the 

method can be modified for small tissue sample sizes as 

long as the solvent ratios are maintained (Herbes and 

Allen, 1983; Gardner et al., 1985). 

13.3.8.2.4 If the Bligh-Dyer method is not the primary 

lipid method used, the chosen lipid analysis method 

should be compared with Bligh-Dyer for each tissue 

type. The chosen lipid method can then be converted to 

“Bligh-Dyer” equivalents and the lipid-normalized tissue 

residues reported in “Bligh-Dyer equivalents.” In the 

interim, it is suggested that extra tissue of each species 

be frozen for future lipid analysis in the event that a 

different technique proves more advantageous (Lee et 

al., 1994). 

13.4 Interpretation of Results 

13.4.1 Section 14 describes general information for 

interpretation of test results. The following sections describe species-specific information that is useful in helping to interpret the results of sediment bioaccumulation 

tests with L. wariegatus. 

13.4.2 Durafion of Exposure: Because data from 

bioaccumulation tests often will be used in ecological or 

human health risk assessments, the procedures are 

designed to generate quantitative estimates of 

steady-state tissue residues. Eighty percent of 

steady-state is used as the general criterion (Lee et al., 

1994). Because results from a single or few species 

often will be extrapolated to other species, the procedures are designed to maximize exposure to 

sediment-associated contaminants so as not to systematically underestimate residues in untested species. 

13.4.2.1 A kinetic study can be conducted to estimate 

steady-state concentrations instead of conducting a 28-d 

bioaccumulation test (e.g., sample on Day 1, 3, 7, 14, 

28; E.L. Brunson, NBS, Columbia, MO, unpublished 

data; USEPA-USCOE, 1991). A kinetic test conducted 

under the same test conditions outlined above, can be 

used when 80% of steady-state will not be obtained 

within 28 d or when more precise estimates of 

steady-state tissue residues are required. Exposures 

shorter than 28 d may be used to determine whether 

compounds are bioavailable (i.e., bioaccumulation potential). 

13.4.2.2 DDT reportedly reached 90% of steady state 

by Day 14 of a 56 d exposure with L. variegafus. 

However, low molecular weight PAHs (e.g., 

acenaphthylene, fluorene, phenanthrene) generally 

peaked at Day 3 and tended to decline to Day 56 (E.L. 

Bruson, NBS, Columbia, MO, unpublished data). In 

general, concentrations of high molecular weight PAHs 

(e.g., benzo(b)fluoranthene, benzo(e)pyrene, indeno 

(1,2,3c,d)pyrene) either peaked at Day 28 or continued 

to increase during the 56 d exposure. 

13.4.3 influence of Indigenous Organisms: 

Field-collected sediments may include indigenous oligochaetes. Phipps et al. (1993) recommend testing 

extra chambers without the addition of L. variegates to 

check for the presence of indigenous oligochaetes in 

fieldcollected sediment. 

13.4.4 Sediment Toxicity in Bt’oaccumulation Tests: 

Toxicity or altered behavior of organisms in a sample 

may not preclude use of bioaccumulation data; however, information on adverse effects of a sample should 

be included in the report. 

13.4.4.1 Grain Size: Lumbriculus wariegatus are tolerant of a wide range of substrates. Physicochemical 

characteristics (e.g., grain size) of sediment were not 

significantly correlated to the growth or reproduction of 

L. variegatus in 10-d toxicity tests (see Section 10.1.3.3; 

Ankley et al., 1994a). 

13.4.4.2 Sediment Organic Carbon: Reduced growth 

of L. variegatus may result from exposure to sediments 

with low organic carbon concentrations (G.T. Ankley, 

USEPA, Duluth, MN, personal communication). Therefore, number of organisms and behavior in the 4-d 

toxicity screening test should be the criteria used to 

judge the acceptability of a bioaccumulation test. 

63 
Section 14 

Data Recording, Data Analysis and Calculations, and Reporting 

14.1 Data Recording 

14.1.1 Quality assurance project plans with data quality 

objectives and standard operating procedures should be 

developed before starting a test. Procedures should be 

developed by each laboratory to verify and archive data. 

14.1.2 A file should be maintained for each sediment 

test or group of tests on closely related samples (Section 9). This file should contain a record of the sample 

chain-of-custody; a copy of the sample log sheet; the 

original bench sheets for the test organism responses 

during the sediment test(s); chemical analysis data on 

the sample(s); control data sheets for reference toxicants; detailed records of the test organisms used in the 

test(s), such as species, source, age, date of receipt, 

and other pertinent information relating to their history 

and health; information on the calibration of equipment 

and instruments; test conditions used; and results of 

reference toxicant tests. Original data sheets should be 

signed and dated by the laboratory personnel performing the tests. 

14.1.3 Example data sheets are included in Appendix 

D. 

14.2 Data Analysis 

14.2.1 Statistical methods are used to make inferences 

about populations, based on samples from those populations. In most sediment toxicity and bioaccumulation 

tests, test organisms are exposed to contaminated sediment to estimate the response of the population of 

laboratory organisms. The organism response to these 

contaminated sediments is usually compared with the 

response to a control or reference sediment, or in some 

analyses of bioaccumulation test data, with a fixed standard such as an Food and Drug Administration (FDA) 

action level. In any toxicity or bioaccumulation test, 

summary statistics such as means and standard errors 

for response variables (e.g., survival, contaminant levels 

in tissue) should be provided for each treatment (e.g., 

pore-water concentration, sediment). 

14.2.1.1 Types of Data. Two types of data can be 

obtained from sediment toxicity or bioaccumulation tests. 

The most common endpoint in toxicity testing is mortality, which is a dichotomous or categorical type of data. 

Other endpoints commonly encountered in sublethal 

evaluations are growth (e.g. in sediment toxicity tests 

conducted with amphipods and midges) and tissue concentrations (e.g. in sediment bioaccumulation tests conducted with oligochaetes or polychaetes and mollusks; 

USEPA, 1994a). These types of endpoints are representative of continuous data. 

14.2.1.2 Sediment Testing Scenarios. Sediment tests 

are conducted to determine whether contaminants in 

sediment are harmful to or are bioaccumulated in benthic 

organisms. Sediment tests are commonly used in studies designed to (1) evaluate hazards of dredged material, (2) assess site contamination in the environment 

(e.g., to rank areas for cleanup), and (3) determine 

effects of specific contaminants, or combinations of 

contaminants, through the use of sediment spiking techniques. Each of these broad study designs has specific 

statistical design and analytical considerations, which 

are detailed below. 

14.2.1.2.1 Dredged Material Hazard Evaluation. In 

these studies, n sites are compared individually to a 

reference sediment. The statistical procedures appropriate for these studies are generally pairwise comparisons. Additional information on toxicity testing of dredged 

material and analysis of data from dredged material 

hazard evaluations is available in USEPA-USCOE 1994. 

14.2.1.2.2 Site Assessment of Field Contamination. 

Surveys of sediment toxicity or bioaccumulation often 

are included in more comprehensive analyses of biological, chemical, geological, and hydrographic data. Statistical correlation can be improved and costs may be 

reduced if subsamples are taken simultaneously for 

sediment toxicity or bioaccumulation tests, chemical 

analyses, and benthic community structure determinations. There are several statistical approaches to field 

assessments, each with a specific purpose. If the objective is to compare the response or residue level at all 

sites individually to a control sediment, then the pairwise 

comparison approach described below is appropriate. If 

the objective is to compare among all sites in the study 

area, then a multiple comparison procedure that employs an experiment-wise error rate is appropriate. If the 

objective is to compare among groups of sites, then 

orthogonal contrasts are a useful data analysis technique. 

64 
14.2.1.2.3 Sediment-Spiking Experiments. Sediments 

spiked with known concentrations of contaminants can 

be used to establish cause and effect relationships 

between chemicals and biological responses. Results of 

toxicity tests with test materials spiked into sediments at 

different concentrations may be reported in terms of an 

LC50, EC50, IC50, NOEC, or LOEC. Results of bioaccumulation tests with either field or spiked samples may 

be reported in terms of a BSAF (biota-sediment accumulation factor, Ankley et al. 1992b). The statistical 

approach outlined above for spiked sediment toxicity 

tests also applies to the analysis of data from sediment 

dilution experiments or water-only reference toxicant 

tests. 

14.2.2 The guidance outlined below on the analysis of 

sediment toxicity and bioaccumulation test data is 

adapted from a variety of sources including Lee et al. 

(1994), USEPA (1993a), USEPA (1993b), USEPA 

(1993c), and USEPA-USCOE (1994). The objectives of 

a sediment toxicity or bioaccumulation test are to quantify contaminant effects on or accumulation in test organisms exposed to natural or spiked sediments or 

dredged materials and to determine whether these effects are statistically different from those occurring in a 

control or reference sediment. Each experiment consists of at least two treatments: the control and one or 

more test treatment(s). The test treatment(s) consist(s) 

of the contaminated or potentially contaminated 

sediment(s). A control sediment is always required to 

ensure that no contamination is introduced during the 

experiment setup and that test organisms are healthy. A 

control sediment is used to judge the acceptability of the 

test. Some designs will also require a reference sediment that represents an environmental condition or potential treatment effect of interest. 

14.2.2.1 Experimental Unit. During toxicity testing, 

each test chamber to which a single application of 

treatment is applied is an experimental unit. During 

bioaccumulation testing, however, the test organism 

may be the experimental unit if individual members of 

the test species are evaluated and they are large enough 

to provide sufficient biomass for chemical analysis. The 

important concept is that the treatment (sediment) is 

applied to each experimental unit as a discrete unit. 

Experimental units should be independent and should 

not differ systematically. 

14.2.2.2 Replication. Replication is the assignment of 

a treatment to more than one experimental unit. The 

variation among replicates is a measure of the 

within-treatment variation and provides an estimate of 

within-treatment error for assessing the significance of 

observed differences between treatments. 

14.2.2.3 Minimum Detectable Difference (MDD). As 

the minimum difference between treatments which the 

test is required or designed to detect decreases, the 

number of replicates required to meet a given significance level and power increases. Because no consensus currently exists on what constitutes a biologically 

acceptable MDD, the appropriate statistical minimum 

significant difference should be a data quality objective 

(DQO) established by the individual user (e.g., program 

considerations) based on their data requirements, the 

logistics and economics of test design, and the ultimate 

use of the sediment toxicity or bioaccumulation test 

results. 

14.2.2.4 Minimum Number of Replicates. Four replicates per treatment or control are the absolute minimum 

number of replicates for a sediment toxicity test. However, USEPA recommends five replicates for marine 

testing (USEPA, 1994) or eight replicates for freshwater 

testing for each control or experimental treatment. It is 

always prudent to include as many replicates in the test 

design as are economically and logistically possible. A 

minimum of five replicates per treatment also is recommended for bioaccumulation testing. USEPA sediment 

toxicity testing methods recommend the use of 20 organisms per replicate for marine testing (USEPA, 1994a) 

or 10 organisms per replicate for freshwater testing. An 

increase in the number of organisms per replicate in all 

treatments, including the control, is allowable only if (1) 

test performance criteria for the recommended number 

of replicates are achieved and (2) it can be demonstrated that no change occurs in contaminant availability 

due to the increased organism loading. 

14.2.2.5 Randomization. Randomization is the unbiased assignment of treatments within a test system and 

to the exposure chambers ensuring that no treatment is 

favored and that observations are independent. It is also 

important to (1) randomly select the organisms (but not 

the number of organisms) for assignment to the control 

and test treatments (e.g., a bias in the results may occur 

if all of the largest animals are placed in the same 

treatment), (2) randomize the allocation of sediment 

(e.g., do not take all the sediment in the top of a jar for 

the control and the bottom for spiking), and (3) randomize the location of exposure units. 

14.2.2.6 Pseudoreplication. The appropriate assignment of treatments to the replicate exposure chambers 

is critical to the avoidance of a common error in design 

and analysis termed “pseudoreplication” (Hurlbert, 1984). 

Pseudoreplication occurs when inferential statistics are 

used to test for treatment effects even though the treatments are not replicated or the replicates are not statistically independent (Hurlbert, 1984). The simplest form 

of pseudoreplication is the treatment of subsamples of 

the experimental unit as true replicates. For example, 

two aquaria are prepared, one with control sediment, the 

other with test sediment, and 10 organisms are placed in 

each aquarium. Even if each organism is analyzed 

individually, the 10 organisms only replicate the biological response and do not replicate the treatment (i.e., 

sediment type). In this case, the experimental unit is the 

10 organisms and each organism is a subsample. A less 

obvious form of pseudoreplication is the potential systematic error due to the physical segregation of exposure chambers by treatment. For example, if all the 

control exposure chambers are placed in one area of a 

65 
room and all the test exposure chambers are in another, 

spatial effects (e.g., different lighting, temperature) could 

bias the results for one set of treatments. Random 

physical intermixing of the exposure chambers or randomization of treatment location may be necessary to 

avoid this type of pseudoreplication. Pseudoreplication 

can be avoided or reduced by properly identifying the 

experimental unit, providing replicate experimentat units 

for each treatment, and applying the treatments to each 

experimental unit in a manner that includes random 

physical intermixing (interspersion) and independence. 

However, avoiding pseudoreplication completely may 

be difficult or impossible given resource constraints. 

14.2.2.7 Composithg Samples. Compositing is used 

primarily in bioaccumufation experiments when the biomass of an individual organism is insufficient for chemicat analysis. Compositing consists of combining samples 

(e.g., organisms, sediment) and chemically analyzing 

the mixture rather than the individual samples. The 

chemical analysis of the mixture provides an estimate of 

the average concentration of the individual samples 

making up the composite. Compositing also may be 

used when the cost of analysis is high. Each organism 

or sediment sample added to the composite should be 

of equal size (i.e., wet weight) and the composite should 

be completely homogenized before taking a sample for 

chemical analysis. If compositing is performed in this 

manner, the value obtained from the analysis of the 

composite is the same as the average obtained from 

analyzing each individual sample (within any sampling 

and analytical errors). If true replicate composites (not 

subsample composites) are made, the variance of the 

replicates wilt be less than the variance of the individual 

samples, providing a more precise estimate of the mean 

value. This increases the power of a test between 

means of composites over a test between means of 

individuals or samples for a given number of samples 

analyzed. If cornpositing reduces the actual number of 

replicates, however, the power of the test will also be 

reduced. If composites are made of individuals or samples 

varying in size, the value of the composite and the mean 

of the individual organisms or sediment samples are no 

longer equivalent. The variance of the replicate composites wilt increase, decreasing the power of any test 

between means. In extreme cases, the variance of the 

composites can exceed the population variance (Tetra 

Tech, 1986). Therefore, it is important to keep the 

individuals or sediment samples comprising the composite equivalent in size. If sample sizes vary, consuIt 

the tables in Schaeffer and Janardan (1978) to determine if replicate composite variances will be higher than 

individual sample variances, which would make 

cornpositing inappropriate. 

14.2.3 The purpose of a toxicity or bioaccumulation test 

is to determine if the biological response to a treatment 

sample differs from the response to a control sample. 

Figure 14. t presents the possible outcomes and decisions that can be reached in a statistical test of such a 

hypothesis. The null hypothesis is that no difference 

exists among the mean control and treatment responses. 

Decision TR =Control TR > Control 

TR =Control Correct 

1 -(I 

Type 11 

Error 

P 

TR > Control Type 1 

Er*or 

U 

Correct 

1-P 

(Power) 

Treatment response (TR). Alpha (a) represents the probability of 

making a Type I statistical error (false poative); beta (p) represents 

the probabildy of making a Type II statistical error (false negative). 

Figure 14.1 Treatment response for a Type I and Type II error. 

The alternative hypothesis of greatest interest in sediment tests is that the treatments are toxic, or contain 

concentrations of bioaccumulatabte compounds, relative to the control or reference sediment. 

14.2.3. l Statistical tests of hypotheses can be designed 

to control for the chances of making incorrect decisions. 

In Figure 14.1, alpha (a) represents the probability of 

making a Type I statistical error. A Type I statistical error 

in this testing situation results from the false conclusion 

that the treated sample is toxic or contains chemicat 

residues not found in the control or reference sample. 

Beta (6) represents the probability of making a Type It 

statistical error, or the likelihood that one erroneously 

concludes there are no differences among the mean 

responses in the treatment, control or reference samples. 

Traditionally. acceptable values for a have ranged from 

0.1 to 0.01 with 0.05 or 5% used most commonly. This 

choice should depend upon the consequences of rnaking a Type I error. Historically, having chosen a, environmental researchers have ignored p and the associated 

power of the test (1-S). 

14.2.3.2 Fairweather (1991) presents a review of the 

need for, and the practical implications of, conducting 

power analysis in environmental monitoring studies. This 

review also includes a comprehensive bibliography of 

recent publications on the need for, and use of, power 

analyses in environmental study design and data analysis. The consequences of a Type It statistical error in 

environmental studies should never be ignored and may 

in fact be the most important criterion to consider in 

experimental designs and data analyses that include 

statistical hypothesis testing. To paraphrase Fairweather 

(1991). “The commitment of time, energy and people to 

a false positive (a Type I error) will only continue until the 

mistake is discovered. In contrast, the cost of a false 

negative (a Type If error) will have both short- and 

long-term costs (e.g., ensuing environmental degradation and the eventual cost of its rectification).“ 

14.2.3.3 The critical components of the experimental 

design associated with the test of hypothesis outlined 

above are (1) the required MDD between the treatment 

and control or reference responses, (2) the variance 

among treatment and control replicate experimental units, 

66 
(3) the number of replicate units for the treatment and 

control samples, (4) the number of animals exposed 

within a replicate exposure chamber, and (5) the selected probabilities of Type I (a) and Type II (@) errors. 

14.2.3.4 Sample size or number of replicates may be 

fixed due to cost or space considerations or may be 

varied to achieve a priori probabilities of a and 13. The 

MDD should be established ahead of time based upon 

biological and program considerations. The investigator 

has little control of the variance among replicate exposure chambers. However, this variance component can 

be minimized by selecting test organisms that are as 

biologically similar as possible and maintaining test conditions within prescribed quality control (QC) limits. 

14.2.3.5 The MDD is expressed as a percentage change 

from the mean control response. To test the equality of 

the control and treatment responses, a two-sample t-test 

1 

0.9 

0.8 

with its associated assumptions is the appropriate parametric analysis. If the desired MDD, the number of 

replicates per treatment, the number of organisms per 

replicate and an estimate of typical among replicate 

variability, such as the coefficient of variation (CV) from 

a control sample, are available, it is possible to use a 

graphical approach as in Figure 14.2 to determine how 

likely it is that a 20’70 reduction will be detected in the 

treatment response relative to the control response. The 

CV is defined as 1000/0 x (standard deviation divided by 

the mean). In a test design with 8 replicates per treatmen! and with an a level of 0.05, high power (i.e., XI.8) 

to detect a 20% reduction from the control mean occurs 

only if the CV is 15% or less (Figure 14.2). The choice of 

these variables also affects the power of the test. If 5 

replicates are used per treatment (Figure 14.3), the CV 

needs to be 10% or lower to detect a 20% reduction in 

response relative to the control mean with a power of 

90%. 

20 30 40 50 60 70 

% Reduction of Control Mean 

Figure 14.2 Power of the test vs. percent reduction in treatment response relative to the control mean at various CVs (8 replicates, 

alpha = 0.05 (one-tailed)). 

67 
0.8 

0.7 

0.6 

b 

3 0.5 

8 

0.4 

0.3 

0.2 

0.1 

0 

0 10 20 30 40 50 60 70 

% Reduction of Control Mean 

Figure 14.3 Power of the test vs. percent reduction in treatment response relative to the control mean at various CVs (5 replicates, alpha = 0.05 (one-tailed)). 

14.2.3.6 Relaxing the a level of a statistical test increases the power of the test. Figure 14.4 duplicates 

Figure 14.2 except that a is 0.10 instead of 0.05. Selection of the appropriate a level of a test is a function of the 

costs associated with making Type I and II statistical 

errors. Evaluation of Figure 14.2 illustrates that with a 

CV of 15% and an a level of 0.05, there is an 80% 

probability (power) of detecting a 20% reduction in the 

mean treatment response relative to the control mean. 

However, if a is set at 0.10 (Figure 14.4) and the CV 

remains at 15%, then there is a 90% probability (power) 

of detecting a 20% reduction relative to the control 

mean. The latter example would be preferable if an 

environmentally conservative analysis and interpretation of the data is desirable. 

14.2.3.7 lncreasrng the number of replicates per treatment will increase the power to detect a 20% reduction 

In treatment response relative to the control mean (Figure 14.5). Note, however, that for less than 8 replicates 

per treatment it is difficult to have high power (i.e., 

~0.80) unless the CV is less than 15%. If space or cost 

limit the number of replicates to fewer than 8 per treatment, then it may be necessary to find ways to reduce 

the among replicate variability and consequently the CV. 

Options that are available include selecting more uniform organisms to reduce biological variability or increasing the a level of the test. For CVs in the range of 

30% to 40%, even eight replicates per treatment is 

inadequate to detect small reductions (5 20%) in response relative to the control mean. 

14.2.3.8 The effect of the choice of a and p on number 

of replicates for various CVs is illustrated in Figure 14.6 

in which the combined total probability of Type I and 

Type II statistical errors is fixed and assumed to be 0.25. 

An a of 0.10 therefore establishes a b of 0.15. In Figure 

14.6, if a = p = 0.125, the number of replicates required 

to detect a difference of 20% relative to the control is at 

a minimum. As a or b decrease, the number of replicates required to detect the same 20% difference relative to the control increases. However, the curves are 

68 
0.8 

0.6 

O 10 20 30 40 50 60 70 

% Reduction of Control Mean 

Figure 14.4 Power of the test vs. percent reduction in treatment response relative to the control mean at various CVs (8 replicates, 

alpha = 0.10 (one-tailed)). 

relatively flat over the range of 0.05 to 0.20 and the least one of these texts and associated statistical tables 

curves are very dependent upon the choice of the on hand. A nonparametric statistics text such as Conover 

combined total of a + j3. Limiting the total of a + p to 0.10 (1980) may also be helpful. 

greatly increases the number of replicates necessary to 

detect a preselected percentage reduction in mean treat- 14.2.4.1 Mean. The sample mean (x) is the average 

ment response relative to the control mean. value, or Exi /r-r, where 

14.2.4 Figure 14.7 outlines a decision tree for analysis 

of survival and growth data subjected to hypothesis 

testing. In the tests described herein, samples or observations refer to replicates of treatments. Sample size n 

is the number of replicates (i.e., exposure chambers) in 

an individual treatment, not the number of organisms in 

an exposure chamber. Overall sample size N is the 

combined total number of replicates in all treatments. 

The statistical methods discussed in this section are 

described in general statistics texts such as Steel and 

Torrie (1980), Sokal and Rohlf (1981), Dixon and Massey 

(1983), Zar (1984) and Snedecor and Cochran (1989). 

It is recommended that users of this manual have at 

n = number of observations (replicates) 

x, = ith observation 

Ix, = every x summed = x, + xp + xJ + . . . + x, 

14.2.4.2 Standard Deviation. The sample standard 

deviation (s) is a measure of the variation of the data 

around the mean and is equivalent to ,z. The sample 

variance, s*, is given by the following “machine” or 

“calculation” formula: 

69 
0.8 - 

f 

2 4 6 8 10 

No. of Replicates (n) 

12 14 16 

Figure 14.5 Effect of CV and number of replicates on the power to detect a 20% decrease In treatment response refatlve to the 

control mean (alpha = 0.05 (one-Wed)). 

14.2.4.3 Standard Error of the #earn. The standard 

error of the mean (SE, or S/V%) estimates variation 

among sample means rather than among individual 

values. The SE is an estimate of the SD among means 

that would be obtained from several samples of n observations each. Most of the statistical tests in this manual 

compare means with other means (e.g., dredged sediment mean with reference mean) or with a fixed standard (e.g., FDA action level; Lee et al., 1994). Therefore, 

the “natural” or “random” variation of sample means 

(estimated by SE), rather than the variation among 

indivrdual observations (estimated by s), is required for 

the tests. 

14.2.4.4 Tests of Assumptions. In general, parametric 

statistical analyses such as t-tests and analysis of variance are appropriate only if (1) there are independent, 

replicate experimental units for each treatment, (2) the 

observations within each treatment follow a normal distribution, and (3) variances for both treatments are equal 

or similar. The first assumption is an essential component of experimental design. The second and third assumptions can be tested using the data obtained from 

the experiment. Therefore, before conducting statistical 

analyses, tests for normality and equality of variances 

should be performed. 

14.2.4.4.1 Outliers (extreme values) and systematic 

departures from a normal distribution (e.g., a log-normal 

distribution) are the most common causes of departures 

from normality or equality of variances. An outlier is an 

inconsistent or questionable data point that appears 

unrepresentative of the general trend exhibited by the 

majority of the data. Outliers may be detected by tabulation of the data, plotting, or by analysis of residuals. An 

explanation should be sought for any questionable data 

points. Without an explanation, data points should only 

be discarded with extreme caution. If there is no explanation, the analysis should be performed both with and 

without the outlier, and the results of both analyses 

should be reported. An appropriate transformation, such 

as the arc sine square root transformation, will normalize many distributions (USEPA, 1985). Problems with 

outliers can usually be solved only by using nonpara70 
18 

16 

6 

4 

0!::::::::::::::1:::::::4 

0.01 0.03 0.05 0.07 0.09 0.11 0.13 0.15 0.17 0.19 0.21 0.23 

Alpha (Beta = 0.25 - Alpha) 

Figure 14.6 Effect of alpha and beta on the number of replicates at various CVs (assuming combined alpha + beta = 0.25). 

metric tests, but careful laboratory practices can reduce 

the frequency of outliers. 

14.2.4.4.2 Tests for Normality. The most commonly 

used test for normality for small sample sizes (NeO) is 

the Shapiro-Wilk’s Test. This test determines if residuals 

are normally distributed. Residuals are the differences 

between individual observations and the treatment mean. 

Residuals, rather than raw observations, are tested 

because subtracting the treatment mean removes any 

differences among treatments. This scales the observations so that the mean of residuals for each treatment 

and overall treatments is zero. The Shapiro-Wilk’s Test 

provides a test statistic W, which is compared to values 

of W expected from a normal distribution. W will generally vary between 0.3 and 1 .O, with lower values indicating greater departure from normality. Because normality 

is desired, one looks for a high value of W with an 

associated probability greater than the pre-specified a 

level. 

14.2.4.4.3 Table 14.1 provides a levels to determine 

whether departures from normality are significant. Normality should be rejected when the probability associated with W (or other normality test statistic) is less than 

a for the appropriate total number of replicates (N) and 

design. A balanced design means that all treatments 

have an equal number (n) of replicate exposure chambers. A design is considered unbalanced when the 

treatment with the largest number of replicates (nmax) 

has at least twice as many replicates as the treatment 

with the fewest replicates (nmln). Note that higher a 

levels are used when the number of replicates is small, 

or when the design is unbalanced, because these are 

the cases in which departures from normality have the 

greatest effects on t-tests and other parametric comparisons. If data fail the test for normality, even after transformation, nonparametric tests should be used for additional analyses. 

14.2.4.4.4 Tables of quantiles of W can be found in 

Shapiro and Wilk (1965), Gill (1978), Conover (1980). 

USEPA (1989b) and other statistical texts. These references also provide methods of calculating W, although 

the calculations can be tedious. For that reason, commonly available computer programs or statistical packages are preferred for the calculation of W. 

14.2.4.4.5 Tests for Homogeneity of Variances. There 

are a number of tests for equality of variances. Some of 

these tests are sensitive to departures from normality, 

which is why a test for normality should be performed 

first. Bartlett’s Test or other tests such as Levene’s Test 

or Cochran’s Test (Winer, 1971; Snedecor and Cochran, 

1989) all have similar power for small, equal sample 

sizes (n=5) (Conover et al., 1981), and any one of these 

tests is adequate for the analyses in this section. Many 

software packages for t-tests and analysis of variance 

71 
Test for Normality 

Figure 14.7 Decision tree for analysis of survival and growth data subjected to hypothesis testing. 

(ANOVA) provide at least one of the tests. Bartlett’s Test 

is recommended for routine evaluation of homogeneity 

of variances (USEPA, 1985; USEPA, 1994b; 

USEPA,1994c). 

14.2.4.4.6 If no tests for equality of variances are 

included in the available statistical software, Hartley’s 

F,,, can easily be calculated: 

F max = ( larger of s: , si ) / ( smaller of s: , si ) 

Table 14.1 Suggested a Levels to Use for Tests of Assumptions 

Number of a When Desw Is 

Test Observations’ Balanced Unbalanced2 

Normallty N=2to9 0.10 0.25 

N = lOto 19 0.05 0.10 

N = 20 or more 0.01 0.05 

Eqdallty of va’cances n=2to9 0.10 0.25 

n = 10or more 0.05 0.10 

N = tctal number of observahons (repkates) In all treatments 

combu-ea n = nunber o! observations (replicates) In an Indwjual 

treatment 

: n -3. .r2 nWhen F is large, the hypothesis of equal variances is 

more li@y to be rejected. F is a two-tailed test 

because it does not matter whi% variance is expected 

to be larger. Some statistical texts provide critical values 

of F, (Winer, 1971; Gill, 1978; Rohlf and Sokal, 1981). 

14.2.4.4.7 Levels of a for tests of equality of variances 

are provided in Table 14.1. These levels depend upon 

number of replicates in a treatment (n) and allotment of 

replicates among treatments. Relatively high a’s (i.e., 2 

0.10) are recommended because the power of the above 

tests for equality of variances is rather low (about 0.3) 

when n is small. Equality of variances is rejected if the 

probability associated with the test statistic is less than 

the appropriate a. 

14.2.4.5 Transformations of the Data. When the 

assumptions of normality or homogeneity of variance 

are not met, transformations of the data may remedy the 

problem, so that the data can be analyzed by parametric 

procedures, rather than by a nonparametric technique. 

The first step in these analyses is to transform the 

responses, expressed as the proportion surviving, by 

the arc sine-square root transformation. The arc 

sine-square root transformation is commonly used on 

proportionality data to stabilize the variance and satisfy 

the normality requirement. If the data do not meet the 

72 
assumption of normality and there are four or more 

replicates per group, then the nonparametric test, 

Wilcoxon Rank Sum Test, can be used to analyze the 

data. If the data meet the assumption of normality, 

Bartlett’s Test or Hartley’s F test for equality of variances 

is used to test the homogeneity of variance assumption. 

Failure of the homogeneity of variance assumption leads 

to the use of a modified t test, and the degrees of 

freedom for the test are adjusted. 

14.2.4.5.1 The arc sine-square root transformation consists of determining the angle (in radians) represented 

by a sine value. In this transformation, the proportion 

surviving is taken as the sine value, the square root of 

the sine value is calculated, and the angle (in radians) 

for the square root of the sine value is determined. 

When the proportion surviving is 0 or 1, a special 

modification of the transformation should be used 

(Bartlett, 1937). An example of the arc sine-square root 

transformation and modification are provided below. 

1. Calculate the response proportion (RP) for each 

replicate within a group, where 

RP = (number of surviving organisms)/(number 

exposed) 

2. Transform each RP to arc sine, as follows: 

a. For RPs greater than zero or less than one: 

Angle (in radians) = arc sine &F) 

b. Modification of the arc sine when RP = 0. 

Angle (in radians) = arc sine d z 

where n = number of animals/treatment rep. 

c. Modification of the arc sine when RP = 1 .O. 

Angle = 1.5708 radians-(radians for RP = 0) 

14.2.4.6 Two Sample Comparisons (N&f). The true 

population mean (p) and standard deviation (CJ) are 

known only after sampling the entire population. In most 

cases samples are taken randomly from the population, 

and the s calculated from those samples is only an 

estimate of 0. Student’s I-values account for this uncertainty. The degrees of freedom for the test, which are 

defined as the sample size minus one (n-l), should be 

used to obtain the correct t-value. Student t-values 

decrease with increasing sample size because larger 

samples provide a more precise estimate of f~ and CJ. 

14.2.4.6.1 When using at table, it is crucial to determine 

whether the table is based on one-tailed probabilities or 

two-tailed probabilities. In formulating a statistical hypothesis, the alternative hypothesis can be one-sided 

(one-tailed test) or two-sided (two-tailed test). The null 

hypothesis (H,) is always that the two values being 

analyzed are equal. A one-sided alternative hypothesis 

(H,) is that there is a specified relationship between the 

two values (e.g., one value is greater than the other) 

versus a two-sided alternative hypothesis (H,J which is 

that the two values are simply different (i.e., erther larger 

or smaller). A one-failed test is used when there is an a 

priori reason to test for a specific relationship between 

two means such as the alternative hypothesis that the 

treatment mortality or tissue residue is greater than the 

control mortality or tissue residue. In contrast, the 

two-tailed test is used when the direction of the difference is not important or cannot be assumed before 

testing. 

14.2.4.6.2 Since control organism mortality or tissue 

residues and sediment contaminant concentrations are 

presumed lower than reference or treatment sediment 

values, conducting one-tailed tests is recommended in 

most cases. For the same number of replicates, one-tailed 

tests are more likely to detect statistically significant 

differences between treatments (e.g., have a greater 

power). This is a critical consideration when dealing with 

a small number of replicates (such as 8Itreatment). The 

other alternative for increasing statistical power IS to 

increase the number of replicates, which increases the 

cost of the test. 

14.2.4.6.3 There are cases when a one-tailed test is 

inappropriate. When no a priori assumption can be 

made as to how the values vary in relationship to one 

another, a two-tailed test should be used. An example of 

an alternative two-sided hypothesis is that the reference 

sediment total organic carbon (TOC) content is different 

(greater or lesser) from the control sediment TOC. A 

two-tailed test should also be used when comparing 

tissue residues among different species exposed to the 

same sediment and when comparing bioaccumulation 

factors (BAFs) or biota-sediment-accumulation-factors 

(BSAFs). 

14.2.4.6.4 The t-value for a one-tailed probability may 

be found in a two-tailed table by looking up t under the 

column for twice the desired one-tailed probability. For 

example, the one-tailed t-value for a = 0.05 and df = 20 

is 1.725, and is found in a two-tailed table using the 

column for a = 0.10. 

14.2.4.7 The usual statistical test for comparing two 

independent samples is the two-sample t-test (Snedecor 

and Cochran. 1989). The t-statistic for testing the equality of means X , and:, from two independent samples 

with n, and n2 replicates and unequal variances IS 

r T 

I = (T, - .i, I {.Fi ,I, +.ti ,I> 

- - 

where $ and s: are the sample variances of the two 

groups. Although the equation assumes that the variances of the two groups are unequal, it is equally useful 

for situations in which the variances of the two groups 

are equal. Thie statistic is compared with the Student t 

73 
distribution with degrees of freedom (df) given by 

Satterthwaite’s (1946) approximation: 

3 

‘/I = 

(5; PII +.Xi JlJ )’ 

bf u? In, -I,-.d,n,)2 (0,-l) -. - 

This formula can result in fractional degrees of freedom, 

in which case one should round the degree of freedom 

down to the nearest integer in order to use a t table. 

Using this approach, the degrees of freedom for this test 

will be less than the degrees of freedom for a t-test 

assuming equal variances. If there are unequal numbers 

of replicates in the treatments. the t-test with Bonferroni’s 

adjustment can be used for data analysis (USEPA, 

1993b; USEPA, 1993c). When variances are equal, an 

F test for equality is unnecessary. 

14.2.4.8 Nonparametric Tests. Tests such as the 

t-test, which analyze the original or transformed data 

and which rely on the properties of the normal distribution, are referred to as parametric tests. Nonparametric 

tests, which do not require normally distributed data, 

analyze the ranks of data and generally compare medians rather than means. The median of a sample is the 

middle or 50th percentile observation when the data are 

ranked from smallest to largest. In many cases, nonparametric tests can be performed simply by converting 

the data to ranks or normalized ranks (rankits) and 

conducting the usual parametric test procedures on the 

ranks or rankits. 

14.2.4.8.1 Nonparametric tests are useful because of 

their generality but have less statistical power than 

corresponding parametric tests when the parametric 

test assumptions are met. If parametric tests are not 

appropriate for comparisons because the normality assumption is not met, data should be converted to normalized ranks (rankits). Rankits are simply the z-scores 

expected for the rank in a normal distribution. Thus, 

using rankits imposes a normal distribution over all the 

data, although not necessarily within each treatment. 

Rankits can be obtained by ranking the data, then 

converting the ranks to rankits using the following formula: 

fankir = ‘[(rank - 0.375) I (N + 0.25)] 

where z is the normal deviate and N is the total number 

of observations. Alternatively, rankits may be obtained 

from standard statistical tables such as Rohlf and Sokal 

(1981). 

14.2.4.8.2 If normalized ranks are calculated, the ranks 

should be converted to rankits using the formula above. 

In comparisons involving only two treatments (N=2), 

there is no need to test assumptions on the rankits or 

ranks; simply proceed with a one-tailed t-test for unequal variances using the rankits or ranks. 

14.2.4.9 Analysis of Variance (N>2). Some experiments 

are set up to compare more than one treatment with a 

control while others may also be interested in comparing 

the treatments with one another. The basic design of 

these experiments is the same as for experiments evaluating pairwise comparisons. After the applicable comparisons are determined, the data must to be tested for 

normality to determine if parametric statistics are appropriate and whether the variances of the treatments are 

equal. If normality of the data and equal variances are 

established, then an analysis of variance (ANOVA) may 

be performed to address the hypothesis that all the 

treatments including the control are equal. If normality or 

equality of variance are not established, then transformations of the data may be appropriate or nonparametric statistics can be used to test for equal means. Tests 

for normality of the data should be performed on the 

treatment residuals. A residual is defined as the observed value minus the treatment mean, that is, r+ = o,, 

- (kth treatment mean). Pooling residuals provides an 

adequate sample size to test the data for normality. 

14.2.4.9.1 The variances of the treatments should also 

be tested for equality. Currently there is no easy way to 

test for equality of the treatment means using analysis of 

variance if the variances are not equal. In a toxicity test 

with several treatments, one treatment may have 100% 

mortality in all of its replicates, or the control treatment 

may have 100% survival in all of its replicates. These 

responses result in 0 variance for a treatment that 

results in a rejection of equality of variance in these 

cases. No transformation will change this outcome. In 

this case, the replicate responses for the treatment with 

0 variance should be removed before testing for equality 

of variances. Only those treatments that do not have 0 

replicate variance should be used in the ANOVA to get 

an estimate of the within treatment variance. After a 

variance estimate is obtained, the means of the treatments with 0 variance may be tested against the other 

treatment means using the appropriate mean comparison. Equality of variances among the treatments can be 

evaluated with the Hartley F, test or Bartlett’s test. The 

option of using nonparametric statistics on the entire set 

of data ia also an alternative. 

14.2.4.9.2 If the data are not normally distributed or the 

variances among treatments are not homogeneous, even 

after data transformation, nonparametric analyses are 

appropriate. If there are four or more replicates per 

treatment and the number of replicates per treatment is 

equal, the data can be analyzed with Steel’s Many-One 

Rank test. Unequal replication among treatments requires data analysis with the Wilcoxon Rank Sum test 

with Bonferroni’s adjustment. Steel’s Many-One Rank 

test is a nonparametric test for comparing treatments 

with a control. This test is an alternative to the Dunnett’s 

Procedure, and may be applied to data when the normality assumption has not been met. Steel’s test requires equal variances across treatments and the control but is thought to be fairly insensitive to deviations 

from this condition (USEPA, 1993a). Wilcoxon’s Rank 

Sum Test is a nonparametric test to be used as an 

alternative to the Steel’s test when the number of replicates are not the same within each treatment. A 

Bonferroni’s adjustment of the pairwise error rate for 

74 
comparison of each treatment versus the control is used 

to set an upper bound of alpha on the overall error rate. 

This is in contrast to the Steel’s test with a fixed overall 

error rate for alpha. Thus, Steel’s test is a more powerful 

test (USEPA, 1993a). 

14.2.4.9.3 Different mean comparison tests are used 

depending on whether an cx percent comparison-wise 

error rate or an a percent experiment-wise error rate is 

desired. The choice of a comparison-wise or 

experiment-wise error rate depends on whether a 

decision is based on a pairwise comparison 

(comparison-wise) or from a set of comparisons 

(experiment-wise). For example, a comparison-wise 

error rate would be used for deciding which stations 

along a gradient were acceptable or not acceptable, 

relative to a control or reference sediment. Each individual comparison is performed independently at a 

smaller a (than used in an experiment-wise comparison) 

such that the probability of making a Type I error in the 

entire series of comparisons is not greater than the 

chosen experiment-wise a level of the test. This results 

in a more conservative test when comparing any particular sample to the control or reference. However, if 

several samples were taken from the same area and the 

decision to accept or reject the area was based upon all 

comparisons with a reference then an experiment-wise 

error rate should be used. When an experiment-wise 

error rate is used, the power to detect real differences 

between any two means decreases as a function of the 

number of treatment means being compared to the 

control treatment. 

14.2.4.9.4 The recommended procedure for pairwise 

comparisons that have a comparison-wise a error rate 

and equal replication is to do an ANOVA followed by a 

one-sided Fisher’s Least Significant Difference (LSD) 

test (Steel and Torrie, 1980). A Duncan’s mean comparison test should give results similar to the LSD. If the 

treatments do not contain equal numbers of replicates, 

the appropriate analysis is the t-test with Bonferroni’s 

adjustment. For comparisons that maintain an 

experiment-wise a error rate Dunnett’s test is recommended for comparisons with the control. 

14.2.4.9.5 Dunnett’s test has an overall error rate of a, 

which accounts for the multiple comparisons with the 

control. Dunnett’s procedure uses a pooled estimate of 

the variance, which is equal to the error value calculated 

in an ANOVA. Dunnett’s procedure can only be used 

when the same number of replicate test chambers have 

been used at each treatment and the control. 

14.2.4.9.6 To perform the individual comparisons, calculate the t statistic for each treatment and control 

combination, as follows: 

t, = 

(v, -v, 

swJclq + (l/n,) 

where v, = mean for each treatment 

v, = mean for the control 

SW = square root of the within mean square 

n, = number of replicates in the control 

n, = number of replicates for treatment “i” 

To quantify the sensitivity of the Dunnett’s test, the 

minimum significant difference (MSD=MDD) may be 

calculated with the foflowing formula: 

MSD= d S,,‘o+(lln) 

where d = Critical value for the Dunnett’s Procedure 

SW = The square root of the within mean 

square 

n = The number of replicates per treatment. 

assuming an equal number of replicates 

at all treatment concentrations 

n, = Number of replicates in the control 

14.2.5 Methods for Calculating L CXOs, 

ECsOs, and ICps 

14.251 Figure 14.8 outlines a decision tree for analysis 

of point estimate data. USEPA manuals (USEPA, 1985; 

USEPA, 1989b; USEPA, 1993b; USEPA. 1993c) discuss in detail the mechanics of calculating LC50 (or 

EC50) or ICp values using the most current methods. 

Two or More Partial Mortalities 

t 

YeS No 

Significant Chi Square Test One Partial Mortahty 

tC50 and 95% Confidence Intervals 

Figure 14.8 Decision tree for analysis of point estimate dala. 

75 
The most commonly used methods are the Graphical, 

Probit, Trimmed Spearman-Karber and the Linear Interpolation Methods. In general, results from these methods should yield similar estimates. Each method is 

outlined below and recommendations are presented for 

the use of each method. 

14.2.5.2 Data for at least five test concentrations and 

the control should be available to calculate an LC50 

although each method can be used with fewer concentrations. Survival in the lowest concentration must be at 

least 50% and an LC50 should not be calculated unless 

at least 50% of the organisms die in at least one of the 

serial dilutions. When less than 50% mortality occurs in 

the highest test concentration, the LC50 is expressed as 

greater than the highest test concentration. 

14.2.5.3 Due to the intensive nature of the calculations 

for the estimated LC50 and associated 95% confidence 

interval using most of the following methods, it is recommended that the data be analyzed with the aid of computer software. Computer programs to estimate the 

LC50 or ICp values and associated 95% confidence 

intervals with the methods discussed below (except for 

the Graphical Method) were developed by USEPA and 

can be obtained by sending a diskette with a written 

request to USEPA, Environmental Monitoring Systems 

Laboratory (EMSL), 26 W. Martin Luther King Drive, 

Cincinnati, OH 45268 or call 513/569-7076. 

14.2.5.4 Graphical Method. This procedure estimates 

an LC50 (or EC50) by linearly interpolating between 

points of a plot of observed percentage mortality versus 

the base 10 logarithm (log,,) of treatment concentration. 

The only requirement for its use is that treatment mortalities bracket 50%. 

14.2.5.4.1 For an analysis using the Graphical Method 

the data should first be smoothed and adjusted for 

mortality in the control replicates. The procedure for 

smoothing and adjusting the data is detailed in the 

following steps: Let pO, p,, . . . . pk denote the observed 

proportion mortalities for the control and the k treatments. The first step is to smooth the p, if they do not 

satisfy p3 5 p, 5 . . . 5 pk. The smoothing process replaces 

any adjacent p,‘s that do not conform to p0 5 p,< . . . 5 pk 

with their average. For example, if p, is less than p,., then 

P; = (P;” -P$V(l-PZ) 

where p: = the smoothed observed proportion mortality for concentration i. 

Adjust the smoothed observed proportion mortality in 

each treatment for mortality in the control group using 

Abbott’s formula (Finney, 1971). The adjustment takes 

the form: 

PF = (P? - PZ)/(l -P!) 

where p; = the smoothed observed proportion mortality for the control 

p; = the smoothed observed proportion mortality for concentration i. 

14.2.5.5 The Probit Method. This method is a parametric statistical procedure for estimating the LC50 (or 

EC50) and the associated 95% confidence interval 

(Finney, 1971). The analysis consists of transforming 

the observed proportion mortalities with a Probit transformation, and transforming the treatment concentrations to log,,,. Given the assumption of normality for the 

log,, of the tolerances, the relationship between the 

transformed variables mentioned above is about linear. 

This relationship allows estimation of linear regression 

parameters, using an iterative approach. A Probit is the 

same as a z-score: for example, the Probit correspond- 

. . ing to 70% mortalrty 1s z.,~ or =.52. The LC50 is calculated from the regression and is the concentration associated with 50% mortality or ~0. To obtain a reasonably 

precise estimate of the LC50 with the Probit Method, the 

observed proportion mortalities must bracket 0.5 and 

the log,, of the tolerance should be normally distributed. 

To calculate the LC50 estimate and associated 95% 

confidence interval, two or more of the observed proportion mortalities must be between zero and one. The 

original percentage mortalities should be corrected for 

control mortality using Abbott’s formula (Section 

14.2.5.4.1; Finney, 1971) before the Probit transformation is applied to the data. 

14.2.5.5.1 A goodness-of-fit procedure with the 

chi-square statistic is used to determine if the data fit the 

Probit model. If many data sets are to be compared to 

one another, the Probit Method is not recommended 

because it may not be appropriate for many of the data 

sets. This method also is only appropriate for percent 

mortality data sets and should not be used for estimating 

endpoints that are a function of the control response, 

such as inhibition of growth. Most computer programs 

that generate Probit estimates also generate confidence 

interval estimates for the LC50. These confidence interval estimates on the LC50 may not be correct if replicate 

mortalities are pooled to obtain a mean treatment response (USEPA-USCOE, 1994). This can be avoided 

by entering the Probit-transformed replicate responses 

and doing a least squares regression on the transformed data. 

14.2.5.6 The Trimmed Spearman-Karber Method, 

The Trimmed Spearman-Karber Method is a modification of the Spearman-Karber, nonparametric statistical 

procedure for estimating the LC50 and the associated 

95% confidence interval (Hamilton et al., 1977). This 

procedure estimates the trimmed mean of the distribu76 
tion of the Iog,0 of the toferance. If the log tolerance 

distribution is symmetric, this estimate of the trimmed 

mean is equivalent to an estimate of the median of the 

fog tolerance distribution. Use of the Trimmed 

Spearman-Karber Method is only appropriate when the 

requirements for the Probit Method are not met (USEPA, 

1993b; USEPA, 1993c). This method is only appropriate 

for lethality data sets. 

14.2.5.6.1 To calculate the LC50 estimate with the 

Trimmed Spearman-Karber Method, the smoothed, adjusted, observed proportion mortalities must bracket 0.5. 

To calculate a confidence interval for the LC50 estimate, 

one or more of the smoothed, adjusted, observed proportion mortalities must be between zero and one. 

14.2.5.6.2 Smooth the observed proportion mortalities 

as described for the Probit Method. Adjust the smoothed 

observed proportion mortality in each concentration for 

mortality in the control group using Abbott’s formula (see 

Probit Method Section 14.2.5.4.1). Calculate the amount 

of trim to use in the estimation of the LC50 as follows: 

Trim = max(pe, 1- pt) 

where p: = the smoothed, adjusted proportion mortality for the lowest treatment concentration, exclusive of the control. 

p; = the smoothed, adjusted proportion mortality for the highest treatment concentration. 

k = the number of treatment concentrations, 

exclusive of the control. 

14.2.5.7 Linear Interpolation Method. This method 

calculates a toxicant concentration that causes a given 

percent reduction (e.g., 25%, 50%, etc.) in the endpoint 

of interest and is reported as an ICp value (IC = Inhibition Concentration; where p = the percent effect). The 

procedure was designed for general applicability in the 

analysis of data from chronic toxicity tests, and the 

generation of an endpornt from a continuous model that 

allows a traditional quantitative assessment of the precision of the endpoint, such as confidence limits for the 

endpoint of a single test, and a mean and coefficient of 

variation for the endpoints of multiple tests. 

14.2.5.7.1 As described in USEPA (1993b; 1993c), the 

Linear Interpolation Method of calculating an ICp assumes that the responses (1) are monotonically 

nonincreasing, where the mean response for each higher 

concentration is less than or equal to the mean response for the previous concentration, (2) follow a piecewise linear response function, and (3) are from a random, independent, and representative sample of test 

data. If the data are not monotonically nonincreasing, 

they are adjusted by smoothing (averaging). In cases 

where the responses at the low toxicant concentrations 

are much higher than in the controls, the smoothing 

process may result in a large upward adjustment in the 

control mean. In the Linear Interpolation Method, the 

smoothed response means are used to obtain the ICp 

estimate reported for the test. No assumption is made 

about the distribution of the data except that the data 

within a group being resampled are independent ana 

identically distributed. 

14.2.5.7.2 The Linear Interpolation Method assumes 

a linear response from one concentration to the next. 

Thus, the IC is estimated by iinear interpolation between 

two concentrations whose responses bracket the response of interest, the (p) percent reduction from the 

control. 

14.2.5.7.3 If the assumption of monotonicity of test 

results is met, the observed response means ( VI) should 

stay the same or decrease as the toxicant concentratron 

increases. If the means do not decrease monotonically. 

the responses are “smoothed” by averaging (pooling) 

adjacent means. Observed means at each concentration are considered in order of increasing concentration. 

starting with the control mean (5,). If the mean observed 

response at the lowest toxicant concentration (7,) is 

equal to or smaller than the control mean ( V,), it is used 

as the response. If it is larger than the control mean, it is 

averaged with the control, and this average is used for 

both the control response (M,) and the lowest toxicant 

concentration response (M,). This mean is then compared to the mean observed _response for the next 

higher toxicant concentration (Y,). Again, if the mean 

observed response for the next higher toxicant concentration is smaller than the mean of the control and the 

lowest toxicant concentration, it is used as the response. 

If it is higher than the mean of the first two, it is averaged 

with the first two, and the mean is used as the response 

for the control and two lowest concentrations of toxicant. 

This process is continued for data from the remaining 

toxicant concentrations. Unusual patterns in the deviations from monotonicity may require an additional step 

of smoothing. Where V, decrease monotonically. the 7, 

become M, without smoothing. 

14.2.5.7.4 To obtain the ICp estimate, determine the 

concentrations C and C which bracket the response 

M, (1 - p/100), where M,?s the smoothed control mean 

response and p is the percent reduction in response 

relative to the control response. These calculations can 

easily be done by hand or with a computer program as 

described below. The linear interpolation estimate is 

calculated as follows: 

IC~=C~+(M,(1-p/100)-M~] cc,., -C,) 

(MJ.1 -M,) 

where CJ = tested concentration whose observed 

mean response is greater than M, (1 - 

p/r 00). 

C = J. 1 tested concentration whose observed 

mean response is less than M.( 1 - pi 

100). 

77 
M. = smoothed mean response for the control. 

M, = smoothed mean response for concentration J. 

M = J.’ smoothed mean response for concentration J + 1. 

P = percent reduction in response relative 

to the control response. 

ICp = estimated concentration at which there 

is a percent reduction from the 

smoothed mean control response. 

14.2.5.7.5 Standard statistical methods for calculating 

confidence intervals are not applicable for the ICp. The 

bootstrap method, as proposed by Efron (1982), is used 

to obtain the 95% confidence interval for the true mean. 

In the bootstrap method, the test data Y,, is randomly 

resampled with replacement to produce a new set of 

data Y,,‘, that is statistically equivalent to the original 

data, but which produces a new and slightly different 

estimate of the ICp (ICp’). This process is repeated at 

least 80 times (Marcus and Holtzman, 1988) resulting in 

multiple “data” sets, each with an associated ICp’ estimate. The distribution of the ICp’ estimates derived from 

the sets of resampled data approximates the sampling 

distribution of the ICp estimate. The standard error of 

the ICp is estimated by the standard deviation of the 

individual ICp’ estimates. Empirical confidence intervals 

are derived from the quantiles of the ICp’ empirical 

distribution. For example, if the test data are resampled 

a minimum of 80 times, the empirical 2.5% and the 

97.5% confidence limits are about the second smallest 

and second largest ICp’ estimates (Marcus and 

Holtzman, 1988). The width of the confidence intervals 

calculated by the bootstrap method is related to the 

variability of the data. When confidence intervals are 

wide, the reliability of the IC estimate is in question. 

However, narrow intervals do not necessarily indicate 

that the estimate is highly reliable, because of undetected violations of assumptions and the fact that the 

confidence limits based on the empirical quantiles of a 

bootstrap distribution of 80 samples may be unstable. 

14.2.6 Analysis of Bioaccumulation Data. In some 

cases, body burdens will not approach steady-state 

body burdens In a 28-d test (Lee et al., 1994). Organic 

compounds exhibiting these kinetics will probably have 

a log Kow >5, be metabolically refractory (e.g., highly 

chlorinated PCBs, dioxins), or have low depuration rates. 

Additionally, tissue residues of several heavy metals 

may gradually increase over time so that 28 d is inadequate to approach steady-state. Depending on the 

goals of the study and the adaptability of the test species 

to long-term testing, it may be necessary to conduct an 

exposure longer than 28 d (or a kinetic study) to obtain a 

sufficiently accurate estimate of steady-state tissue residues of these compounds. 

14.2.6.1 Biotic Sampling. In the long-term studies, the 

exposure should continue until steady-state body burdens are attained. ASTM (1988b) recommends a minimum of five sampling periods (plus to) when conducting 

water exposures to generate bioconcentration factors 

(BCFs). Sampling in a geometric progression is also 

recommended with sampling times reasonably close to 

S/16, S/8, S/4, S/2, and S, where S is the time to 

steady-state. This sampling design assumes a fairly 

accurate estimate of time to steady-state, which is often 

not the case with sediment exposures. 

14.2.6.1 .l To document steady-state from sediment 

exposures, placing a greater number of samples at and 

beyond the predicted time to steady-state is recommended. With a contaminant expected to reach 

steady-state within 28 to 50 d, samples should be taken 

at Day 0, 7, 14, 21, 28, 42, 56, and 70. If the time to 

steady-state is much greater than 42 d, then additional 

sampling periods at two week intervals should be added 

(e.g., Day 84). Slight deviations from this schedule (e.g., 

Day 45 versus Day 42) are not critical, though for 

comparative purposes, samples should be taken at t,,. 

An estimate of time to steady-state may be obtained 

from the literature or estimated from structure-activity 

relationships, though these values should be considered 

the minimum times to steady-state. 

14.2.6.1.2 This schedule increases the likelihood of 

statistically documenting that steady-state has been obtained although it does not document the initial uptake 

phase as well. If an accurate estimate of the sediment 

uptake rate coefficient (KS) is required, additional sampling periods are necessary during the initial uptake 

phase (e.g., Day 0, 2, 4, 7, 10, 14). 

14.2.6.2 Abiotic Samples. The bioavailable fraction of 

the contaminants as well as the nutritional quality of the 

sediment are more prone to depletion in extended tests 

than during the 28-d exposures. To statistically document whether such depletions have occurred, replicate 

sediment samples should be collected for physical and 

chemical analysis from each sediment type at the beginning and the end of the exposure. Archiving sediment 

samples from every biological sampling period also is 

recommended. 

14.2.6.3 Short-Tern Uptake Tests. Compounds may 

attain steady-state in the oligochaete, Lumbriculus 

variegatus, in less than 28 d (Kukkonen and Landrum, 

1993). However, before a shorter test is used, it must be 

ascertained that the analytes of interest do indeed achieve 

steady-state in L. variegatus in ~28 d. Biotic and abiotic 

samples should be taken at Day 0 and 10 following the 

same procedure used for the 28-d tests. If time-series 

biotic samples are desired, sample on Day 0, 1, 3, 5, 7, 

and 10. 

14.2.6.4 Estimating Steady-State. In tests where 

steady-state cannot be documented, it may be possible 

to estimate steady-state concentrations. Several methods have been published that can be used to predict 

78 
steady-state contaminant levels from uptake and depuration kinetics (Spacie and Hamelink, 1982; Davies and 

Dobbs, 1984). Alf of these methods were derived from 

fish exposures and most use a linear uptake, first-order 

depuration model that can be modified for uptake of 

contaminants from sediment. To avoid confusing uptake 

from water versus sediment, KS, the sediment uptake 

rate coefficient, is used instead of Kl. The KS coefficient 

has also been referred to as the uptake clearance rate 

(Landrum et at., 1989). Following the recommendation 

of Stehly et al. (1990) the gram sediment and gram 

tissue units are retained in the formulation: 

Ct (t) = KS x Cs / K2 x (l-e-k2X*) 

where Ct = contaminant concentration in tissue 

at time t 

cs = contaminant concentration in sediment 

KS = uptake rate coefficient in tissue (g sed 

g-l day-‘) 

K2 = depuration constant (day-l) 

t = time (days) 

As time approaches infinity, the maximum or equilibrium 

contaminant concentration within the organism (Ct,a,) 

becomes 

ct man = Cs x KS / K2 

Correspondingly, the bioaccumulation factor (BAF) for a 

compound may be estimated from 

BAF= Ks/K2 

14.2.6.4.1 This model assumes that the sediment concentration and the kinetic coefficients are invariant. Depletion of the sediment concentrations in the vicinity of the 

organism would invalidate the model. Further, the rate 

coefficients are conditional on the environment and health 

of the test organisms. Thus, changes in environmental 

conditions such as temperature or changes in physiology such as reproduction will also invalidate the model. 

Despite these potential limitation, the model can provide 

estimates of steady-state tissue residues. 

14.2.6.4.2 The kinetic approach requires an estimate of 

KS and K2, which are determined from the changes in 

tissue residues during the uptake phase and depuration 

phase, respectively. The uptake experiment should be 

short enough that an estimate of KS is made during the 

linear portion of the uptake phase to avoid an unrealistltally low uptake rate due to depuration. The depuration 

phase should be of sufficient duration to smooth out any 

loss from a rapidly depurated compartment such as loss 

from the voiding of feces. Unless there is reason to 

suspect that the route of exposure will affect the depuration rate, it is acceptable to use a K2 derived from a 

water exposure. For further discussion of this method for 

bioconcentration studies in fish, see Davies and Dobbs 

(1984), Spacie and Hamelink (1982), and ASTM (1988b). 

For application of this procedure for sediment, see Lee 

et al. (1994). Recent studies of the accumulatron of 

sediment-associated contaminants by benthos suggest 

that the kinetics for freshly dosed sediments may require 

a more complex formulation to estimate the uptake 

clearance constant than that presented above (Landrum, 

1989). 

14.2.6.4.3 This model predicts that equilibrium would be 

reached only as time becomes infinite. Therefore, for 

practical reasons, apparent steady-state is defined here 

as 95% of the equilibrium tissue residue. The time to 

reach steady-state can be estimated by 

S = In[l / (l.OO-0.95)] / K2 = 3.0 / K2 

where S = time to apparent steady-state (days) 

Thus, the key information is the depuration rate of the 

compound of interest in the test species or phylogenetitally related species. Unfortunately, little of this data has 

been generated for benthic invertebrates. When no depuration rates are available, the depuration rate constant 

for organic compounds can then be estimated from the 

relationship between Kow and k2 for fish species (Spacie 

and Hamelink, 1982): 

K2 = antilog[l.47-0.414 x log(Kow)] 

The relationship between S and k2 and between k2 and 

Kow is summarized in Table 14.2. Estimated time (days) 

to reach 95% of contaminant steady-state tissue residue 

(S) and depuration rate constants (k2) are calculated 

from octanol-water partition coefficients using a linear 

uptake, first-order depuration model (Spacie and 

Hamelink, 1982). The k2 values are the amount depurated (decimal fraction of tissue residue lost per day). 

Table 14.2 may be used to make a rough estimate of the 

exposure time to reach steady-state tissue residues if a 

depuration rate constant for the compound of interest 

from a phylogenetically similar species is available. If no 

depuration rate is available, then the table may be used 

Table 14.2 Estimated lime to Obtain 95 Percent of Steady-State 

Tissue Residue 

LogKow K2 S (days) 

.~ -___ -~~ 

1 0.114 0.2 

2 0.44 0.5 

3 0.17 14 

4 0.0065 35 

5 0.0025 9.2 

6 0.00097 24 

7 000037 61 

6 0.00014 160 

9 0.00006 410 

.~ -___ -~~ 

79 
for estimating the S of organic compounds from the Kow 

value. However, as these data were developed from fish 

bioconcentration data, its applicability to the kinetics of 

uptake from sediment-associated contaminants is unknown. The portion of organics readily available for 

uptake may be small in comparison to the total sediment 

organic concentration (Landrum, 1989). Therefore S 

values generated by this model should be considered as 

minimum time periods. 

14.2.6.4.4 Using a linear uptake, first-order depuration 

model to estimate exposure time to reach steady-state 

body burden for metals is problematical for a number of 

reasons. The kinetics of uptake may be dependent upon 

a small fraction of the total sediment metal load that is 

bioavailable (Luoma and Bryan, 1982). Depuration rates 

may be more difficult to determine, as metals bound to 

proteins may have very low exchange rates (Bryan, 

1976). High exposure concentrations of some metals 

can lead to the induction of metal binding proteins, like 

metallothionein, which detoxify metals. These 

metal-protein complexes within the organism have extremely low exchange rates with the environment (Bryan, 

1976). Thus, the induction of metal binding proteins may 

result in decreased depuration rate constants in organisms exposed to the most polluted sediments. Additionally, structure-activity relationships that exist for organic 

contaminants (e.g., relationship between Kow and BCFs) 

are not well developed for metals. 

14.3 Data Interpretation 

14.3.1 Sediments spiked with known concentrations of 

contaminants can be used to establish cause and effect 

relationships between chemicals and biological responses. Results of toxicity tests with test materials 

spiked into sediments at different concentrations may be 

reported in terms of an LC50 (median lethal concentration), an EC50 (median effect concentration), an IC50 

(inhibition concentration), or as an NOEC (no observed 

effect concentration) or LOEC (lowest observed effect 

concentration; Section 3). Consistent spiking procedures 

should be followed in order to make interlaboratory 

compansons (Section 8.3). 

14.3.2 Evaluating effect concentrations for chemicals in 

sediment requires knowledge of factors controlling the 

broavailabllity. Similar concentrations of a chemical in 

units of mass of chemical per mass of sediment dry 

weight often exhibit a range in toxicity in different sediments (Di Toro et al., 1991; USEPA, 1992c). Effect 

concentrations of chemicals in sediment have been 

correlated to interstitial water concentrations, and effect 

concentrations in interstitial water are often similar to 

effect concentratrons in water-only exposures. The bioavailabillty of nonronic organic compounds are often 

inversely correlated with the organic carbon concentration of the sediment. Whatever the route of exposure, 

the correlations of effect concentrations to interstitial 

water concentrations indicate predicted or measured 

concentrations in interstitial water can be useful for 

quantifying the exposure concentration to an organism. 

Therefore, information on partitioning of chemicals between solid and liquid phases of sediment may be useful 

for establishing effect concentrations. 

14.3.3 Toxic units can be used to help interpret the 

response of organisms to multiple contaminants in sediment. A toxic unit is the concentration of a chemical 

divided by an effect concentration. For example, a toxic 

unit of exposure can be calculated by dividing the measured concentration of a chemical in pore water by the 

water-only LC50 for the same chemical (Ankley et al., 

199 1 a). Toxicity expressed as toxic units may be summed 

and this may provide information on the toxicity of 

chemical mixtures (Ankley et al., 1991a). 

14.3.4 Field surveys can be designed to provide either a 

qualitative reconnaissance of the distribution of sediment contamination or a quantitative statistical comparison of contamination among sites (Burton and Ingersoll, 

1994). Surveys of sediment toxicity are usually part of 

more comprehensive analyses of biological, chemical, 

geological, and hydrographic data. Statistical correlation 

can be improved and costs reduced if subsamples are 

taken simultaneously for sediment toxicity or bioaccumulation tests, chemical analyses, and benthic community structure. 

14.3.5 Descriptive methods such as toxicity tests with 

field-collected sediment should not be used alone to 

evaluate sediment contamination. An integration of several methods using the weight of evidence is needed to 

assess the effects of contaminants associated with sediment. Hazard evaluations integrating data from laboratory exposures, chemical analyses, and benthic community assessments provide strong complementary evidence of the degree of pollution-induced degradation in 

aquatic communities (Chapman et al., 1992; Burton, 

1991). 

14.3.6 Toxicity Identification Evaluation (TIE) procedures can be used to help provide insights as to specific 

contaminants responsible for toxicity in sediment 

(USEPA, 1991a; Ankley and Thomas, 1992). For example, the toxicity of contaminants such as metals, 

ammonia, hydrogen sulfide, and nonionic organic compounds can be identified using TIE procedures. 

14.3.7 hterpretafion of Comparisons of Tissue Residues. If the mean control tissue residues at Day 28 are 

not significantly greater than the Day 0 tissue residues, it 

can be concluded that there is no significant contamination from the exposure system or from the control sediment. If there is significant uptake, the exposure system 

or control sediment should be reevaluated as to suitability. Even if there is a significant uptake in the controls, it 

is still possible to compare the controls and treatments 

as long as the contaminant concentrations in the test 

tissue residues are substantially higher. However, if 

control values are high, the data should be discarded 

and the experiment conducted again after determining 

the source of contamination. 

80 
14.3.7.1 Comparisons of the 28-d control (or reference) 

tissue residues and 28d treatment tissue residues determines whether there was statistically significant bioaccumulation due to exposure to test sediments. Comparisons between control and reference tissue residues 

at Day 28 determine whether there was a statistically 

significant bioaccumulation due to exposure to the reference sediment. If no significant difference is detected 

when treatment tissue residues are compared to a set 

criterion value (e.g., FDA Action Limit) with a one-tailed 

test, the residues must be considered equivalent to the 

value even though numerically the mean treatment tissue residue may be smaller. 

14.3.7.2 BAFs and BSAFs. Statistical comparisons 

between ratios such as BAFs or BSAFs are difficult due 

to computation of error terms. Since all variables used to 

compute BAFs and BSAFs have errors associated with 

them, it is necessary to estimate the variance as a 

function of these errors. This can be accomplished using 

approximation techniques such as the propagation of 

error (Beers, 1957) or a Taylor series expansion method 

(Mood et al., 1974). BAFs and BSAFs can then be 

compared using these estimates of the variance. See 

Lee et al. (1994) provide examples of this approach. 

14.3.7.3 Comparing Tissue Residues of Different 

Compounds. In some cases, it is of interest to compare 

the tissue residues of different compounds. For example, Rubinstein et al. (1987) compared the uptake of 

thirteen different PCB congeners to test for differences 

in bioavailability. Because the values for the different 

compounds are derived from the same tissue samples, 

they are not independent and tend to be correlated, so 

standard t-tests and ANOVAs are inappropriate. A repeated measures technique (repeated testing of the 

same experimental unit) should be used where the 

experimental unit (individual) is considered as a random 

factor and the different compounds as a second factor. 

See Rubinstein et al. (1987) and Lake et al. (1990) for 

an example of the application of repeated measures to 

bioaccumulation data. 

14.4 Reporting 

14.4.1 The record of the results of an acceptable 

sediment test should include the following information 

either directly or by referencing available documents: 

14.4.1.1 Name of test and investigator(s), name and 

location of laboratory, and dates of start and end of test. 

14.4.1.2 Source of control or test sediment, method for 

collection, handling, shipping, storage and disposal of 

sediment. 

14.4.1.3 Source of test material, lot number if applicable, composition (identities and concentrations of major ingredients and impurities if known), known chemical 

and physical properties, and the identity and 

concentration(s) of any solvent used. 

14.4.1.4 Source and characteristics of overlying water, 

description of any pretreatment, and results of any demonstration of the ability of an organism to survive or grow 

in the water. 

14.4.1.5 Source, history, and age of test organisms; 

source, history, and age of brood stock, culture procedures; and source and date of collection of the test 

organisms, scientific name, name of person who identified the organisms and the taxonomic key used, age or 

life stage, means and ranges of weight or length. observed diseases or unusual appearance, treatments. 

holding procedures. 

14.4.1.6 Source and composition of food, concentrations of test material and other contaminants, procedure 

used to prepare food, feeding methods, frequency and 

ration. 

14.4.1.7 Description of the experimental design and test 

chambers, the depth and volume of sediment and overlying water in the chambers, lighting. number of test 

chambers and number of test organisms/treatment, date 

and time test starts and ends, temperature measurements, dissotved oxygen concentration (as percent saturation) and any aeration used before starting a test and 

during the conduct of a test. 

14.4.1.8 Methods used for physical and chemical characterization of sediment. 

14.4.1.9 Definition(s) of the effects used to calculate 

LC50 or ECSOs, biological endpoints for tests, and a 

summary of general observations of other effects. 

14.4.1 .lO A table of the biofogical data for each test 

chamber for each treatment including the control(s) in 

sufficient detail to allow independent statistical analysis. 

14.4.1 .l 1 Methods used for statistical analyses of data. 

14.4.1.12 Summary of general observations on other 

effects or symptoms. 

14.4.1.13 Anything unusual about the test, any deviation from these procedures, and any other relevant 

information. 

14.4.2 Published reports should contain enough information to clearly identify the methodology used and the 

quality of the results. 

81 
Section 15 

Precision and Accuracy 

15.1 Determining Precision and 

Accuracy 

15.1.1 Precision is a term that describes the degree to 

which data generated from replicate measurements differ and reflects the closeness of agreement between 

randomly selected test results. Accuracy is the difference between the value of the measured data and the 

true value and is the closeness of agreement between 

an observed value and an accepted reference value. 

Quantitative determination of precision and accuracy in 

sediment testing of aquatic organisms is difficult or may 

be impossible in some cases, as compared to analytical 

(chemical) determinations. This is due, in part. to the 

many unknown variables that affect organism response. 

Determining the accuracy of a sediment test using field 

samples is not possible since the true values are not 

known. Since there is no acceptable reference material 

suitable for determining the accuracy of sediment tests, 

the accuracy of the test methods has not been determined (Section 15.2). 

15.1.2 Sediment tests exhibit variability due to several 

factors (Section 9). Test variability can be described in 

terms of two types of precision either single laboratory 

(intralaboratory or repeatability; Section 15.5.1) precision or multi-laboratory (interlaboratory or reproducibility; Section 155.2) precision. lntralaboratory precision 

reflects the ability of trained laboratory personnel to 

obtain consistent results repeatedly when performing 

the same test on the same organism using the same 

toxicant. Interlaboratory precision (also referred to as 

round-robin or ring tests) is a measure of how reproducible a method is when conducted by a large number of 

laboratories using the same method, organism, and 

samples. Generally, intralaboratory results are less variable than interlaboratory results (USEPA, 1991b; USEPA, 

1993a; USEPA, 1994b; USEPA, 1994c; Hall et al., 

1989; Grothe and Kimerle, 1985). 

15.1.3 A measure of precision can be calculated using 

the mean and relative standard deviation (percent coefficient of variation. or CV% = standard deviation/mean x 

100) of the calculated endpoints from the replicated 

endpoints of a test. However, precision reported as the 

CV should not be the only approach used for evaluating 

precision of tests and should not be used for the NOEC 

effect levels derived from statistical analyses of hypothesis testing. The CVs may be very high when testing 

extremely toxic samples. For example, if there are multiple replicates with no survival and one with low survival, the CV may exceed 100%. yet the range of 

response is actually quite consistent. Therefore, additional estimates of precision should be used, such as 

range of responses, and minimum detectable differences (MDD) compared to control survival or growth. 

Several factors can affect the precision of the test, 

including test organism age, condition, sensitivity, handling and feeding of the test organisms, overlying water 

quality, and the experience of the investigators in conducting tests. For these reasons, it is recommended that 

trained laboratory personnel conduct the tests in accordance with the procedures outlined in Section 9. Quality 

assurance practices should include (1) single laboratory 

precision determinations using reference toxicants for 

each of the test organisms that are used to determine 

the ability of the laboratory personnel to obtain precise 

results. These determinations should be made before 

conducting a sediment test and should be routinely 

performed as long as whole sediment tests are being 

conducted; (2) control charts (Section 15.3) should be 

prepared for each reference toxicant and test organism 

to determine if the test results are within prescribed 

limits; and (3) tests must meet the minimum criteria of 

test acceptability specific for each test organism (Tables 

11.3. 12.3, 13.4; USEPA, 1991b). 

15.1.4 lntralaboratory precision data are routinely calculated for test organisms using water-only 96-h exposures to a reference toxicant, such as KCI. Intralaboratory precision data should be tracked using a control 

chart. Each laboratory’s reference toxicant data will 

reflect conditions unique to that facility, including dilution 

water, culturing, and other variables (Section 9). However, each laboratory’s reference toxicant CVs should 

reflect good repeatability. 

15.1.5 To date, interlaboratory precision (round-robin) 

tests have been completed with both Hydella azteca 

and Chironomus tentans using 4-d water-only and 10-d 

whole sediment tests. The results of these round-robin 

studies are described below. 

15.2 Accuracy 

15.2.1 The relative accuracy of toxicity tests cannot be 

determined since there is no acceptable reference material. The relative accuracy of the reference toxicity 

82 
tests can only be evaluated by comparing test responses 

to control charts. 

15.3 Replication and Test Sensitivity 

15.3.1 The sensitivity of sediment tests will depend in 

part on the number of replicates per concentration, the 

probability levels (alpha and beta) selected, and the type 

of statistical analysis. For a given level of variability 

remains constant, the sensitivity of the test will increase 

as the number of replicates is increased. The minimum 

recommended number of replicates varies with the objectives of the test and the statistical method used for 

analysis of the data (Section 14). 

15.4 Demonstrating Acceptable 

Laboratory Performance 

15.4.1 It is the responsibility of a laboratory to demonstrate its ability to obtain precise results with reference 

toxicants before it performs sediment tests (Section 

9.16). lntralaboratory precision, expressed as a coefficient of variation (CV), of the range for each type of test 

to be used in a laboratory should be determined by 

performing five or more tests with different batches of 

test organisms, using the same reference toxicant, at 

the same concentrations, with the same test conditions 

(e.g., the same test duration, type of water, age of test 

organisms, feeding), and same data analysis methods, 

A reference toxicant concentration series (0.5 or higher) 

should be selected that will consistently provide partial 

mortalities at two or more concentrations of the test 

chemical (Section 9.14, Table 9.1, 9.2). 

15.4.2 The quality of test organisms obtained from an 

outside source must be verified by conducting a 

reference-toxicity test concurrently with the sediment 

test. The supplier should provide data with the shipment 

describing the history of the sensitivity of organisms 

from the same source culture. If the supplier has not 

conducted five reference toxicity tests with the test 

organism, it is the responsibility of the testing laboratory 

to conduct five reference toxicity tests before starting a 

sediment test (Section 9.14.1). 

15.4.3 Before conducting tests with contaminated sediment, the laboratory should demonstrate its ability to 

conduct tests by conducting five exposures in control 

sediment as outlined in Table 11.1, 12.1, or 13.1. It is 

recommended that these five exposures with control 

sediment be conducted concurrently with the five reference toxicity tests described in Section 15.4.2. 

15.4.4 A control chart should be prepared for each 

combination of reference toxicant and test organism. 

Each control chart should include the most current data. 

Endpoints from five tests are adequate for establishing 

the control charts. In this technique, a running plot is 

maintained for the values (X,) from successive tests with 

a given reference toxicant (Figure 15.1), and the endpoints (LC50, NOEC, ICp) are examined to determine if 

Upper Control Limit A 

2 

P . 

Central Tendency 

Lower Control Limtt 

-----. 

rlllllrrll1111111111* 

0 5 10 15 a 

Upper Control Llmlt (Y + 2 S) B 

--- -------- 

I! 

‘i 

4 

Central Tendency 

hi 

Lower Control Limit (Y- 2 S) 

c11111111111111111111, 

0 5 10 15 a0 

Toxlclty Test with Reference ToxIcants 

where x, = Successive toxicity values of toxtcity tests. 

n = Number of tests. 

7 = Mean toxicity value. 

s = Standard deviation. 

Figure 15.1 Control (cusum) charts: (A) hypothesis testing 

and (8) point estimates (LC, EC, or IC). 

they are within prescribed limits. Control charts as described in USEPA (1993a) and USEPA (l993b) are 

used to evaluate the cumulative trend of results from a 

series of samples. The mean and upper and lower 

control limits (+2 SD) are recalculated with each successive test result. After two years of data collection, or a 

minimum of 20 data points, the control (cusum) chart 

should be maintained using only the 20 most recent data 

points. 

15.4.5 The outliers, which are values falling outside the 

upper and lower control limits, and trends of increasing 

or decreasing sensitivity, are readily identified using 

83 
control charts. With an alpha of 0.05, one in 20 tests 

would be expected to fall outside of the control limits by 

chance alone. During a 30 d period, if two reference 

toxicity tests out of a total of the previous 20 fall outside 

the control limits, the sediment toxicity tests conducted 

during the time in which the second reference toxicity 

test failed are suspect and should be considered as 

provisional and subject to careful review. 

15.4.5.1 A sediment test may be acceptable if specified 

conditions of a reference toxicity test fall outside the 

expected ranges (Section 9). Specifically, a sediment 

test should not automatically be judged unacceptable if 

the LC50 for a given reference toxicity test falls outside 

the expected range or if mortality in the control of the 

reference toxicity test exceeds 10%. All the performance 

criteria outlined in Tables 11.3, 12.3, and 13.4 must be 

considered when determining the acceptability of a sediment test. The acceptability of the sediment test would 

depend on the experience and judgment of the investigator and the regulatory authority. 

15.4.6 If the value from a given test with the reference 

toxicant falls more than two standard deviations (SD) 

outside the expected range, the sensitivity of the organisms and the overall credibility of the test system are 

suspect (USEPA, 1993a). In this case, the test procedure should be examined for defects and should be 

repeated with a different batch of test organisms. 

15.4.7 Performance should improve with experience, 

and the control limits for point estimates should gradually narrow. However, control limits of k2 SD, by definition, will be exceeded 5% of the time, regardless of how 

well a laboratory performs. Highly proficient laboratories 

that develop a very narrow control limit may be unfairly 

penalized if a test that falls just outside the control limits 

is rejected de facto. For this reason, the width of the 

control limits should be considered in determining whether 

or not an outlier is to be rejected. This determination 

may be made by the regulatory authority evaluating the 

data. 

15.4.8 The recommended reference toxicity test consists of a control and five or more concentrations in 

which the endpoint is an estimate of the toxicant concentration that is lethal to 50% of the test organisms in 

the time period prescribed by the test. The LC50 is 

determined by an appropriate procedure, such as the 

Trimmed Spearman-Karber Method, or Probit Method, 

Graphical Method, or the Linear interpolation Method 

(Section 14). 

15.4.9 The point estimation analysis methods recommended in this manual have been chosen primarily 

because they are well-tested, well-documented, and are 

applicable to most types of test data. Many other methods were considered in the selection process, and it is 

recognized that the methods selected are not the only 

possible methods of analysis of toxicity data. 

15.5 Precision of Sediment Toxicity 

Test Methods 

75.5. I Intraiaboratory Precision 

15.5.1 .l lntralaboratory precision of the Hyalella azteca 

and Chironomus tentans 10-d tests (as described in 

Tables 11.1 and 12.1) was evaluated at ERL-Duluth 

using one control sediment sample in June 1993. In this 

study, five individuals simultaneously conducted the 1 Od 

whole sediment toxicity tests as described in Tables 

11 .l and 12.1 with the exception of the feeding rate of 

1 .O mL rather than 1.5 mL for C. tentans. The results of 

the study are presented in Table 15.1. The mean survival for H. azteca was 90.4% with a CV of 7.2% and the 

mean survival for C. tentans was 93.0% with a CV of 

5.7%. All of the individuals met the survival performance 

criteria of 80% for H. azteca (Table 11.3) or 70% for C. 

tentans (Table 12.3). 

15.52 lnterlabofatory Precision 

15.5.2.1 Interlaboratory precision using reference toxicity tests and 1 O-d whole sediment toxicity tests using the 

methods described in this manual (Tables 9.1,9.2, 11 .l , 

and 12.1) were conducted by federal government laboratories, contract laboratories, and academic laboratories that had demonstrated experience in sediment toxicity testing (Table 15.2). The only exception to the 

methods outlined in Table 9.1 and 9.2 was that 80% 

rather than the current recommendation of 90% survival 

was used to judge the acceptability of the reference 

toxicity tests. The round robin study was conducted in 

two phases for each test organism. The experimental 

design for the round robin study required each laboratory to conduct 96-h water-only reference toxicity tests in 

Phase 1 and 10-d whole sediment tests in Phase 2 with 

Hyalella azteca or Chironomus tenfans over a period of 

six months. Criterta for selection of participants in the 

Tabk 15.1 

Individual 

lntralaboratory Precision for Survlvsl of HyaMla 

azteca and Chlronomus tentans in 10-d WholeSediment Toxicity Tests, June 1993’ 

Percent Surwval 

H. azteca C. tentans 

A 85 85 

B 93 93 

C 90 93 

0 84 94 

E 100 100 

N 5 5 

Mean 90.4 93.0 

cv 7.2% 5.7% 

1 Test sample was from a control sediment (T.J. Norberg-King, 

USEPA, Duluth, MN. personal communication.) The test was 

conducted at the same time by five individuals at ERL-Duluth. The 

source of overlying water was from Lake Superior. 
round-robin study were that the laboratories had (1) 

existing cultures of the test organisms, (2) experience 

conducting tests with the organisms, and (3) would 

participate voluntarily. The test methods for the reference toxicity tests and the whole sediment toxicity tests 

were similar among laboratories. Standard operating 

procedures detailing the test methods were provided to 

all participants. Culture methods were not specified and 

were not identical across laboratories. 

15.5.2.2 In Phase 1, water-only reference toxicity (KCI) 

tests were conducted with H. azteca for 96-h and LCSOs 

were calculated. In these tests, H. azteca were placed in 

reconstituted hard water in 250-mL beakers containing a 

small piece of plastic mesh substrate. Ten organisms 

were randomly added to each of four replicates at five 

concentrations of KCI and a control. The organisms 

were fed 0.5 mL of a 1800 mg/L stock solution of YCT on 

Day 0 and Day 2. Mortality was monitored at 24 h 

intervals and the test was ended at 96 h (Table 9.2). In 

Phase 2, the variability of the 10-d whole sediment test 

procedure for H. azteca was evaluated using an automated water renewal exposure system (Table 11.1 and 

Section 6.3). This system consisted of eight replicate 

300-mL beakers per treatment with each containing 10 

organisms each. Each beaker contained a lOO-mL aliquot of sediment and the overlying water was replaced 

twice a day (Table 11.1). The test sediments that were 

previously tested at ERL-Duluth to ascertain their toxicity included a control sediment (RR 3), a moderately 

Table 15.2 Participants in Round Robin Studies’ 

Chironomus tentans Hyalella azteca 

96 h 96 h 10-d 96 h 10-d 

KC1 KC1 Sediment KCI Sediment 

Test Test Test Test Test 

Laboratory Dee 92 May 93 May 93 Ott 92 Mar 93 

~--- 

Lab A Y N N Y N 

Lab 0 Y Y Y Y Y 

Lab C Y N Y Y Y 

Lab D Y Y Y N N 

Lab E Y Y Y Y Y 

Lab F Y Y Y Y Y 

Lab G Y Y Y Y Y 

Lab H Y N N Y N 

Lab I Y Y Y 2 Y 

Lab J Y Y Y Y Y 

Lab K 3 -3 -3 Y Y - 

Lab L 4 -1 -4 Y Y 

___. 

N 9 7 8 10 9 

’ Y = Laboratory participated II-I testing sediment samples. 

2 Test in January 1993. 

J Participated using C. riparius only. 

’ Did not intend to parilcipate with C. tentans. 

contaminated sediment (RR 2), and a heavily contaminated test sediment (RR 1). Sediments RR 2 and RR 3 

were contaminated primarily with copper. An additional 

sediment heavily contaminated with polycyclic aromatic 

hydrocarbons (RR 4) was tested by five laboratories. At 

the end of a test, the sediment from each replicate was 

sieved and surviving organisms were counted. 

15.5.2.3 Ten laboratories participated in the H. azteca 

reference toxicity test (Table 15.2). The results from the 

tests with KCI are summarized in Table 15.3. The test 

performance criteria of 280% control survival was met 

by 90% of the laboratories resulting in a mean control 

survival of 98.8% (CV = 2.1%). The mean LC50 was 305 

mg/L (CV = 14.2%) and the LC5Os ranged from 232 to 

372 mg/L KCI. 

15.5.2.4 In the 10-d whole sediment tests with H. 

azteca, nine laboratories tested the three sediments 

described above and five laboratories tested a fourth 

sediment from a heavily contaminated site (Table 15.4). 

All laboratories completed the tests; however, laboratory 

C had 75% survival, which was below the acceptable 

test criteria for survival (Table 11.3). For these tests, the 

CV was calculated using the mean percent survival for 

Table 15.3 Interlaboratory Precision for Hyalella azfeca 96-h 

LC5Os from Water-Only Static Acute Toxicity Tests 

Using a Reference Toxicant (KCI) (October 1992) 

KCI Percent 

LC50 Confidence Intervals Control 

Laboratory OWLI Lower Upper Survival 

Lab A 372 352 395 100 

Lab B 321 294 350 98 

Lab C 232 205 262 100 

LabD -1 -1 -1 -1 

Lab E 325 282 374 100 

Lab F 276 240 316 98 

Lab G 297 267 331 73 

Lab H 336 317 356 100 

Lab I 1 422 101 200 93 

Lab J 337 286 398 100 

Lab L 250 222 282 100 

N 10 10 

Mean 289.03 96.2% 

cv 23.0%? 8.3% 

N 9 9 

Mean 305.04 98.8 

cv 14.2%’ 2.1% 

‘Laboratory did not parttctpate in 14. azteca test In October. 

*Results are from a retest In January using three concentrations only; 

results excluded from analysis. 

3Mean 1 and CV 1 include all data points. 

‘Mean 2 and CV 2 exclude data points for all sediment samples from 

laboratorles that did not meet minlmum control survival of 280%. 

85 
the eight laboratories that met the performance criteria 

for the test. The CV for the control sediment (RR 3) was 

5.8% with a mean survival of 94.5% with survival ranging from 86% to 100%. For sediments RR 2 and RR 4, 

the mean survival was 3.3% and 4.3%, respectively 

(Table 15.4). For RR 2, survival ranged from 0% to 24% 

(CV = 253%) and for RR 4 the survival ranged from 0% 

to 11% (CV = 114%). Survival in the moderately contaminated sediment (RR 1) was 54.2% with survival 

ranging from 23% to 76% (CV = 38.9%). When the RR 1 

data for each laboratory were compared to the control 

for that laboratory, the range for the minimum detectable 

difference between the test sediments and the control 

sediment ranged from 5 to 24% with a mean of 11% (SD 

= 6). 

15.5.2.5 The Phase 1 C. tentans reference toxicity test 

was conducted with KCI on two occasions (Tables 15.5 

and 15.6). Both tests were conducted in 20 mL of test 

solution in 30-mL beakers using 10 replicates per treatment with 1 organism per beaker. Animals were fed 0.25 

mL of a 4 g/L solution of Tetrafin@ on Day 0 and Day 2 

(Table 9.1). For the first reference toxicity test comparison, 10 laboratories participated, and eight laboratories 

met the survival criteria of the round robin, which was 

80% survival (Table 15.5). The mean LC50 for the eight 

laboratories that met the survival criterion was 4.25 g/L 

(CV of 51.8%). The LC5Os ranged from 1.25 to 6.83 g/L. 

Length and instar were determined for a subset of 

organisms at the start of the tests for some of the 

laboratories. When length was correlated with the LC50, 

the larger animals were less sensitive than the smaller 

animals. The effect level was significantly correlated (r2 

= 0.78) with the organism size, which ranged from 1.56 

mm to 10.87 mm (ages of animals ranged from 7- to 

13-d postdeposition). The majority of these animals 

were the third instar, with the smallest animals in their 

first instar and the largest animals a mix of third and 

fourth instar (Table 15.5) as determined by head capsule width. 

15.5.2.6 For the second Phase 1 KCI reference toxicity 

tests with C. fentans, seven laboratones participated 

(Table 15.6). The test conditions were identical to those 

in the previous reference toxicity test except that a 

minimum size was specified rather than using initial age 

of the animals. Each laboratory was instructed to start 

the test when larvae were at least 0.4 to 0.6 mm long. 

Therefore, a more consistent size of test organisms was 

used in this test. Six out of the seven laboratories met 

the 280% control survival criterion with a mean LC50 of 

5.37 g/L (CV = 19.6%). The LC5Os ranged from 3.61 to 

6.65 g/L. 

15.5.2.7 In the 10-d whole sediment test with C. tentans 

eight laboratories participated. The same three sediments used in the H. azteca whole sediment test were 

used for this test (Table 15.7). All test conditions were 

those as described in Table 12.1 with the exception of 

the feeding rate of 1.0 mL rather than 1.5 mL for C. 

tenfans. Three laboratories did not meet the control 

criteria for acceptable tests of 170% survival in the 

control (RR 3) sediment; Table 12.3). For the five laboratories that successfully completed the tests, the mean 

survival in the control sediment (RR 3) was 92.0% (CV 

of 8.3%) and survival ranged from 81.2% to 98.8%. For 

Table 15.4 Interlaboratory Precision for Survival of Hyde//a azfeca in 10-d Whole Sediment Toxicity Tests Using Four Sediments 

(March 1993) 

Mean Percent Survival (SD) in Sedrment Samples 

Laboratory RR 1 RR2 RR 3 (Control) RR4 

-.-- --__ -.__~ .__- 

Lab A -I -1 -1 -1 

Lab B 76.2 

Lab C 57.522 

Lab D -’ 

Lab E 46.2 

Lab F 72.5 

Lab G 50.0 

Lab f-f -’ 

Lab I 73.7 

Lab J 65.0 

Lab K 22.5 

Lab L 27.5 

(20.7) 

(14.9) 

(17.7) 

(12.8) 

(28.3) 

(32.0) 

(9.3) 

(18.3) 

(16.7) 

2.5 (7.1) 

t .2* (0) 

-1 

0 (0) 

23.7 (18.5) 

0 (0) 

-1 

0 (0) 

0 (0) 

0 (0) 

0 (0) 

97.5 

75.02 

-1 

97.5 

98.7 

100 

-’ 

86.2 

96.2 

95.0 

86.2 

(4.6) 

(17.7) 

(7.1) 

(3.5) 

(0) 

(10.6) 

(5.2) 

(5.3) 

(18.5) 

11.2 (13.6) 

1.22 (0) 

-1 

- 

0 (0) 

3.3 (5.2) 

-: 

- 

2.5 17.1) 

- 

- 

.__ 

N 9 9 9 5 

Mean l3 54.6 3.0 93.0 3.6 

cv 1 36.2% 256% 9.0% 12 1% 

N 8 8 9i.5 4 

Mean 2’ 54.2 3.3 4.3 

cv2 38.9% 253% 5.8% 114% 

.-~ .~. -- .-._ .-.. -- 

. Laboratory drd not particrpate in H. azteca test in March. 

;’ Survival in control sediment (RR 3) below minimum acceptable level. 

3 Mean 1 and CV 1 include all data points. 

’ Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that dd not meet mrmmum control survwal ol ‘80%. 

86 
Table 15.5 Interlaboratory Precision for Chirononws lentans 96-h LC5Os from Water-only Static Acute Toxicity Tests Using a 

Reference Toxicant (KCI) (December 1992) 

Laboratory 

KCI 

LC50 

W) 

Confidence Interval 

Lower Upper 

Control Mean 

Survival length 

(%I (mm) 

It-star 

SZl 

of Test 

Age at 

Start 

of Test 

(day) 

Lab A 6.19 5.37 

Lab 0 6.83 6.38 

Lab C 5.00 4.16 

Lab D 3.17 2.29 

Lab E 2.002 -2 

Lab F 1.25 -3 

Lab G 6.28 5.26 

Lab H 2.89 2.39 

Lab I 6.66 6.01 

Lab J 1.77 0.59 

7.13 

7.31 

6.01 

4.40 

- 

- 

7.50 

3.50 

7.24 

5.26 

75’ 10.87 

100 10.43 

100 5.78 

100 5.86 

80 6.07 

80 1.56 

95 7.84 

95 6.07 

100 -’ 

65’ 4.42 

3.4 

3 

3 

3 

3 

3 

3 

-’ 

2.3 

10 

13 

11 

11 

11 

12 

11 

7 

10 

7 

N 10 10 8 10 

Mean I5 4.20 89.0 6.6 10.3 

cv 1 52.7% 14.5% 46.6% 17.9% 

N 8 

Mean 26 4.25 

9i.8 7 8 

6.2 10.75 

cv2 51.8% 9.3% 39.5% 15.2% 

Control survival below minimum acceptable level. 

2 Unable to calculate LC50 with tnmmed Spearman Karber; no confidence interval could be calculated. 

3 Confidence intervals cannot be calculated as no partial mortalities occurred. 

4 No animals were measured. 

5 Mean 1 and CV 1 mclude all data pomts. 

6 Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that did not meet minimum control survival of 280%. 

Table 15.6 Interlaboratory Precision for Chironomus lentans 96-h LC5Os from Water-only Static Acute Toxicity Tests Using a 

Reference Toxicant (KCI) (May 1993) 

Laboratory 

KCI 

LC50 

(gju 

Age at 

Control Start 

Confidence Interval Survival of Test 

Lower Upper VW (day) 

Lab A -1 - - - - 

Lab I3 6.65 -2 - 90 12 

Lab C -1 - - - - 

Lab D 5.30 4.33 6.50 553 10 

Lab E 5.11 4.18 6.24 100 11 

Lab F 3.61 2.95 4.42 90 10 

Lab G 5.36 4.43 6.49 93 12 

Lab H -1 - - - 

Lab I 5.30 4.33 6.52 95 10-11 

Lab J 6.20 4.80 7.89 100 13 

n 7 7 7 

Mean 1’ 5.36 89 11.1 

cv 1 17.9% 17.5% 9.46% 

Zlean 2 5 6 5.37 94.7 6 11.2 6 

cv2 19.6% 4.8% 9.13% 

’ Did not participate In reference toxicity test in April. 

7 Confidence Intervals cannot be calculated as no partial mortalities occurred. 

3 Control survtval below mimmum acceptable level. 

q Mean 1 and CV 1 include all data points. 

c Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that did not meet minimum control survival of 270%. 

87 
the RR 2 sediment sample, the mean survival among 

the five laboratories was 3.0% (CV = 181%) and for the 

RR 1 sediment sample, the mean survival was 86.8% 

(CV = 13.5%). A significant effect on survival was not 

evident for the RR 1 sample, but growth was affected 

(Table 15.8). When the RR 1 data for each laboratory 

were compared to the control for that laboratory, the 

minimum detectable difference for survival among laboratories ranged from 2.3 to 12.1 with a mean of 8% (SD 

= 4). 

1552.8 For C. tentans, growth is a sensitive indicator 

of sediment toxicity (Ankley et al., 1993) and growth was 

also measured in the round-robin comparison (Table 

15.8). Using the data from five laboratories with acceptable control survival in the control sediment (RR 3), the 

mean weight of C. tentans for the control sediment (RR 

3) was 1.254 mg (CV = 26.6%). The C. tentans in the 

moderately contaminated sediment (RR 1) had a mean 

weight of 0.546 mg (CV = 31.9%). No growth measurements were obtained for C. fentans in sediment RR 2 

because of the high mortality. The mean minimum detectable difference for growth among laboratories meeting the survival performance criteria was 11% (SD = 5) 

and the MDD ranged from 4.8 to 23.6% when the RR 1 

data were compared to the RR 3 data. 

15.5.2.9 These tests exhibited similar or better precision than many chemical analyses and effluent toxicity 

test methods (USEPA, 1991 b; USEPA, 1993a). The 

success rate for test initiation and completion of the 

EPA’s round-robin evaluations is a good indication that 

a well equipped and trained staff will be able to successfully conduct these tests. This is an important consideration for any test performed routinely in any regulatory 

program. 

Table 15.7 Interlaboratory Precision for Survival of Chironomus tentans in 19-d WholeSedIment Toxicity Tests Using Three 

Sediments (May 1993) 

Laboratory RR 1 

Mean Percent Survival (SD) in Sediment Samples 

RR2 RR 3 (Control) 

Lab A 

Lab B 

Lab C 

LabD 

LabE 

Lab F 

Lab G 

Lab H 

Lab I 

Lab J 

-1 

67.5 (14.9) 

15.02 (12.0) 

tm.ff (20.0) 

85.0 (11.9) 

87.52 (12.5) 

90.0 (13.1) 

-1 

97.5 (4.6) 

93.8 (11.8) 

-1 

2.5 

02 

02 

0 

02 

12.5 

-1 

0 

0 

(7.1) 

(0) 

(0) 

(0) 

(0) 

(3.5) 

(0) 

(0) 

-1 

98.8 (3.5) 

62.52 (26.0) 

66.32 (27.7) 

93.8 (9.2) 

43.a2 (30.2) 

87.5 (10.3) 

-1 

98.8 (3.5) 

81.2 (8.3) 

N 8 a 8 

Mean l3 74.5 1.89 79.1 

cv 1 36.7% 233% 25.1% 

N 5 5 5 

Mean 2’ 86.8 3.0 92.0 

cv2 135% 181% 8.3% 

’ Old not participate in C. tentans test in May. 

* Survival in control sediment (RR 3) below minimum acceptable level. 

3 Mean 1 and CV 1 rnclude all data points. 

’ Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that did not meet minimum control survrval of >70%. 

88 
Table 15.8 Interlaboratory Precision for Growth of Chlronomus tentans in 10-d WholeSediment Toxicity Tests Using Three 

Sediments (May 1993) 

Laboratory RR 1 

Growth-Dry Weight in mg (SD) in Sediment Samples 

RR2 RR 3 (Control) 

Lab A -1 -’ -1 

Lab B 0.370 (0.090) 0 (0) 1.300 (0.060) 

Lab C 0.8832 (0.890) 0 (0) 0.504 (0.212) 

Lab D 0.2152 (0.052) 0 (0) 1.070 (0.107) 

Lab E 0.657 (0.198) 0 (0) 0.778 (0.169) 

Lab F 0.2102 (0.120) 0 (0) 0.610 (0.390) 

Lab G 0.718 (0.114) 0 (0) 1.710 (0.250) 

Lab H -1 -1 -1 

Lab I 0.639 (0.149) 0 (0) 1.300 (0.~1 

Lab J 0.347 (0.050) 0 (0) 1.180 (0.123) 

iean l3 

8 8 8 

0.505 - 1.056 

cv 1 49.% - 38.3% 

Lean 2’ 

5 5 5 

0.546 - 1.254 

cv2 31.9% - 26.6% 

’ Did not participate in testing in May. 

* Survival in control sediment (RR 3) below minimum acceptable level. 

3 Mean 1 and CV 1 include all data points. 

’ Mean 2 and CV 2 exclude data points for all sediment samples from laboratories that did not meet minimum control survival of 270%. 

09 
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100 
Appendix A 

Summary USEPA Workshop on 

Development of Standard Sediment Test Methods 

A.1 The USEPA Office of Water, Office of Science and 

Technology, and Office of Research and Development 

held a workshop September 16-18, 1992, in Washington, DC, to provide an opportunity for experts in the field 

of sediment toxicology and staff from USEPA regional 

and Headquarters program offices to discuss the development of standard freshwater and marine sediment 

testing procedures (USEPA, 1992a). As part of USEPA’s 

Contaminated Sediment Strategy, the Agency’s program offices agreed to develop and use consistent tests 

for the assessment of sediment contamination. USEPA 

sponsored research to address uncertainties associated 

with the use of sediment tests discussed at the workshop. The results of discussions held at the workshop 

were used to identify research issues and develop manuals for conducting sediment toxicity and bioaccumulation tests. The following test organisms were selected 

for sediment test method development in 1993: (1) 

Freshwater toxicity tests: Hyalella azteca and Chironomus 

tentans, (2) Freshwater bioaccumulation tests: 

Lumbriculus variegatus, (3) Marine toxicity tests: 

Ampelisca abdita, Rhepoxynius abronius, Eohaustorius 

estuarius, and Leptocheirus plumulosus, (4) Marine bioaccumulation tests: Macoma nasuta and Nereis spp. 

A.2 If funds are available in future years, additional work 

will be started on developing chronic toxicity tests, toxicity identification evaluation (TIE), and test development 

for other organisms. USEPA plans to develop two other 

methods documents: (1) one on sediment spiking and 

(2) one on sediment collection, handling, and storage. 

Parts of the document on collection, handling, and storage methods are already under development for a Quality Assurance and Quality Control guidance document 

that will supplement both the Inland and Ocean Testing 

Manuals for disposal of dredged material 

(USEPA-USCOE, 1991; 1994). 

A.3 Before the workshop on July 13, 1992, a questionnaire was sent out to freshwater workshop invitees and 

other selected researchers. Information was requested 

on culture methods for and testing of H. azteca, C. 

tentans, and L. variegatus and any additional organisms 

used for sediment tests. The discussion topics for culturing and testing of the three species were ranked in order 

of importance for developing standard methods based 

on the similarity of the issues across all tests. The 

following section summarize results of the questionnaire 

for each test organism. 

1. Development of a Standard Testing Method for 

Hyalella azteca. Twenty-one responses to the survey were received and eighteen laboratories reported information on H. azteca (USEPA, 1992a 

and Table A.1). The summary of the survey responses follow. The most common response is underlined and when no item is underlined it indicates 

no single most common response. Published procedures for conducting sediment toxicity tests with H. 

azteca are also listed in Table A.2. 

Table A.1 List of Laboratories Responding to the Survey 

Laboratory Hyalella Chronomus Lumbriculus 

azteca tentans variegatus 

Dept. of Fish. & Oceans. x 

Canada 

Environ. Canada, X X 

Burlington, ON 

EPA-Duluth, Duluth, MN x X x 

EPA Region :, Lexington, x X 

MA 

EPA Region 8. Denver, CO x 

EPA-Newtown. Cincinnati. x X 

OH 

EVS Consultants, X X 

Vancouver, BC 

MD Dept. Environ., X 

Baltimore, MD 

Miami Univ., Oxford, OH x 

Mich. State Univ., X x 

E. Lansing. MI 

NBS-Athens, Athens. GA x 

NBS-Columbia, X 

Columbia, MO 

NOAA-Ann Arbor, X 

Ann Arbor, MI 

Old Dominion. Norfolk. VA x 

State of WA, X 

Manchester, WA 

Univ. of MS, University, x 

MS 

Univ. of WI-Superior. X X x 

Superior. WI 

Wright State Univ., X X 

Dayton, OH 

101 
Table A.2 Summary of Testing Procedures Used to Evaluate the Toxicity of Whole Sediments with Hyalella azteca 

Citation 

Condition [1] [2] [3] [4] [5] [6] 

Temperature (°C) 20 20 22 20-25 20-23 20-22 

Light intensity 

(foot candles) 

NR 25-50 NR 50-100 25-50 NR 

Photoperiod NR 16-8 16-8 16-8 16-8 16-8 

Test chamber (mL) 1000 1000 300 300 30-300 2500 

Sediment volume (mL) 200 200 100 40-50 5-100 -150 

Overlying water 

volume (mL) 

800 100 175 160-200 20-150 -1350 

Renewal rate of) 

overlying water 

(additions/day; 

0 1-4 1-4 variable 0-2 0 

Age of organisms 

(days) 

Size of organisms (mm) 

juvenile juvenile 7-14 juvenile 7-14 0-7 

NR 1-2 NR NR 1-2 NR 

Number of organisms/ 

chamber 

15 20 10 10 3-10 20 

Number of replicate 

chambers/treatment 

NR 4 3 4 5-10 2 

Food AC RC YCT RC YCT, mL TM 

Aeration Yes None None DO<3 None Yes 

Overlying water Natural Natural Natural Natural Reconst. Natural 

Test duration (d) 10 10-28 10 

S 

80 

7 28 

Endpoints S S.G,M S S.G 

Test acceptability 

(survival %) 

NR 80 80 

10 

S 

80 NR 

Citations Conditions 

[1] Nebeker et al. (1984a) Food: YCT = yeast-cerophyll-trout chow, RC = Rabbit chow, 

[2] Ingersoll and Nelson (1990) TM = Tetramin®, ML = maple leaves. 

[3] Ankley et al. (1993a) Endpoints: S = survival, G = growth (length or weight), M = maturation, 

[4] Burton et al. (1989) NR = Not reported 

[5] Winger and Lasier (1993) 

[6] Borgmann and Munawar (1989) 

102 
A. Survey Summary of Culture Methods for H. azteca 

Flow: 

Temperature: 

Light: 

Chamber: 

Age of organisms: 

Frequency restart: 

Water Quality: 

Source of Strains: 

Aeration: 

Feeding: 

Substrate: 

Static vs. renewal 

19 to 25°C (23°C) 

16L:8D photoperiod; about 500 to 1000 lux 

1 L to 80 L 

Known age vs. mixed age 

Monthly, every two months 

Natural vs. reconstituted 

ERL-Duluth, ERL-Corvallis. Burlington, Michigan State (most cultured in moderately-hard 

or hard water) 

Moderate 

leaves, Tetramin®, rabbit chow, diatoms, yeast, wheat grass, Chlorella, alfalfa, Nutrafin® 

YCT, paper towels, Selenastrum, Ankistrodesmus, brine shrimp, aquatic plants, sediment. Feed 2 to 3 times/week typical. 

Leaves, nylon mesh, cotton gauze, 3-M web plastic, paper towels 

Reference toxicants: Cd, Cu, KCI, Zn, NaCI, Cr (water-only exposures) 

B. Survey Summary of Testing Procedures for H. azteca 

Flow: Static vs. renewal 

Aeration: None or moderate 

Temperature: 20 to 25°C (20°C) 

Light: 16L:8D photoperiod; about 250 to 1000 lux 

Chamber: 30 mL to 1 L (250 to 300 ml ) 

Sediment ratio: 1:1 to 1:4 ratio sediment:water 

Age of organisms: Known age (0 to 7 d, 7 to 14 d) vs. mixed age (size about 7 to 14 d) (sieved) 

Number of organisms:5 to 20/chamber (10/chamber) 

Number replicates: 2 to 10/treatment (3 to 5/treatment) 

Duration: 2- to 28d (10-d) 

Feeding: None, Rabbit Chow, YCT, maple leaves, Tetramin® 

Endpoints: Survival, length, weight, sexual maturation (males), young production, bioaccumulation 

Acceptability: Survival (80%), length, weight 

103 
2. Development of a Standard Testing Method for Chironomus 

fentans. Twenty-one responses to the survey were received 

and twelve laboratories reported information on C. teentans 

(USEPA, 1992a and Table A.l). The summary of the survey 

responses follow. The most common response is underlined 

and when no item is underlined it indicates no single most 

common response. Published procedures for conducting sediment toxicity tests with C. tenfans are also listed in Table A.3. 

Table A.3 Summary ot Testing Procedures Usad to Evaluate the Toxicity of Whole Sediments with Chifonomus tentans 

Citation 

Condltlon 111 VI PI I41 Fl 

Temperature (‘C) 

Light lntenslty 

(foot candles) 

20 

NR 

22 

-100 

22 

NR 

23 

NR 

22 

NR 

Photopenod 16-8 16-8 NR NR 

Test chamber (mL) 

Sediment volume (mL) 

Overlymg water 

volume (mL) 

NR 

lOoa 

200 

8co 

3rm 

-250 

2000 

300 

100 

175 

50 

-7.5 

47 

2ooo 

1500 

-200 

Renewal rate of 

overlymg water 

(addltlons/day) 

0 o-5 1-4 0 0 

Age of orgamsms 

(nstar) 

2nd 2nd 2nd 2nd 2nd 

Size of orgamsms NR 0.15 mg NR 

Number of organisms/ 

chamber 

15 25 10 

0.5 g 

1 

6-8 mm 

20 

Number of replrcate 

chambers/treatment 

NR 2 NR 15 NR 

Food TM.CP TM TF TF None 

Aeration Yes None None Yes Yes 

OverlyIng water Natural Natural Natural Natural Natural 

Test duration (d) 10 14 10 17 

EndpoInts S.G S,G G S.G 

Test acceptabtllty 

(survlval %) 

NR NR 

10 

S,G 

70 NR NR 

Clfahons: Conditions 

Ill Nebeker et al. (1984a) Food: CP = cerophyll, RC = Rabbit chow, TM = TetraminB. 

I21 Adams et al. (1985) TF = TetrafM 

I:; 

Ankley et al. (1993a) Endpoints: S = survival, G = growth (length or weight), M = maturation, 

Glesy et al. (1988) NR = not reported 

I51 Wentsel et al. (1977) 

104 
A. Survey Summary of Culture Methods for C. tentans 

Flow: 

Temperature: 

Light: 

Chamber: 

Age of organisms: 

Frequency restart: 

Age restart: 

Water Quality: 

Aeration: 

Feeding: 

Substrate: 

&& vs. renewal 

19 to 25°C (W 

16L:8D photoperiod; about 500 to 1300 lux 

1 L to 80 L 

Known age vs. fpixed u 

2x/week to every 6 months 

Egg masses to 224-h old larvae 

Natural vs. reconstituted 

Moderate 

Tetramin@, Nutrafin@, YCT and algae, alfalfa and Tettafin@, feed daily to ax/week 

Paper towels (bleached or unbleached); sand 

Reference toxicants: Cu, NaCI, Cd, KCI (water-only exposures) 

B. Survey Summary of Testing Procedures for C. tentans 

Flow: static, vs. renewal 

Aeration: None or moderate 

Temperature: 20 to 25°C (w 

Light: 16L:BD photoperiod; about 250 to 1300 Iux 

Chamber: 50mLto2 L 

Sediment ratio: 1: 1 to 1;4 ratio sediment:water 

Age of organisms: Knownage(Oto16d;jOto14d) 

Number of organisms:10 to 80/chamber (10 to 15/cNumber replicates: 2-15 (3 to 4) 

Duration: 2 to 14-d (m 

Feeding: Trout chow, Tetrafin@. YCT 

Endpoints: Survival, weight 

Acceptability: Survival (m, weight (dry weight) 

105 
3. Development of a Standard Testing Procedure for Lumbriculus 

variegatus. Twenty-one responses to the survey were received and five laboratories reported information on 

L. variegatus (USEPA. 1992a and Table A.l). The summary 

of the survey responses follow. The most common response 

is underlined and when no item is underlined it indicates no 

single most common response. Published procedures for 

conducting sediment bioaccumulation tests with L. variegatus 

are also listed in Table A.4. 

Table A.4 Summary of Testing Procedures Used to Conduct Whole Sediment Bioaccumulation Tests with Lumbriculus variegatus 

Condition 

Temperature (“C) 

Ill 

20 

(21 

no 

Lrght Intensity 

(fool candles) 

NR 

LJ 

NR 

Citation 

I31 141 

n- L.J 20 

25-50 NR 

Photoperiod NR Various NR 

Test chamber (L) 3-5 0.15-0.6 3-3.8 

Sedrment volume (L) 1.5-2 

Overlying water 

volume (L) 

1.5-3 

30-1809 

0.1-0.45 

16-8 

4 

1 

3 

0.3-0.35 

2.7-3 

Renewal rate of 

overlymg water 

(additrons/day) 

2-6 0.5-l 1 0 

Age of organisms Adult Adult Adult Adult 

Loadrng (@chamber) 1 O.l-0.39/L 

Number of replicate 

chambers/treatment 

NR 

1:50’ 

3-4 3-5 3 

Food None None None Yes 

Aeration None Yes Yes Yes 

OJerlyrng waler NaturaU 

Reconst. 

Natural Natural Natural 

Test duratron fd) 1 O-60 10-60 56 28-44 

lest acceptability NR Biomass 

lrpld 

Bomass NR 

’ ’ -50 g dry ieIght orgamsm:sedrment organic carbon 

NR = not reported 

Crfaffons: 

Ill Phrpps et al (1993) 

PI Kukkonen and Landrum (1993) 

PI E L. Brilnson, NBS, Columbra, MO. unpublished data 

141 Schuytema et al. (1988) 

106 
A. Survey Summary of Culture Methods for L. variegatus 

Flow: 

Temperature: 

Light: 

Chamber: 

Age of organisms: 

Frequency restart: 

Water Quality: 

Aeration: 

Feeding: 

Substrate: 

Reference toxicants: 

Static vs. renewd 

22 to 24°C 

16L:8D photoperiod; illuminance unspecified 

1 L to 80 L 

Mixed-age adults 

Monthly, B 

Natu vs. reconstituted 

Moderate 

Frozen silver cup trout chow, salmon starter, sediment, Tetramin@, yeast, wheat grass. 

Chlorella, alfalfa, NutrafinQ YCT, paper towels. Feed 2 to 3 times/week typical. 

.&per towels, sediment 

No reference toxicants specified 

6. Survey Summary of Testing Procedures for L. variegates 

Flow: Static vs. renewal 

Aeration: None or moderate 

Temperature: 10 to 23°C 

Light: 16L:8D photoperiod; illuminance unspecified 

Chamber: 1 to6L 

Sediment Ratio: 1 :l to 1;4 ratio sediment: water (sediment volumes should be adequate to allow feeding 

and burrowing) 

Age of organisms: Adults, 3.8 cm. 

Number of organisms:Adequate number to provide tissue mass for analysis of residue of concern 

Number replicates: 4 to S/treatment 

Duration: lOto28d 

Endpoints: Bioaccumufation 

Feeding: None 

Acceptability: Adequate tissue mass for residue analysis 

107 
A.4 Workgroup participants arrived at a consensus on 

several culturing and testing procedures. Where it was 

not possible to make a decision because of lack of 

information, the group identified research items that 

need further consideration before a specific decisions 

could be made. 

A.5 In developing guidance for culturing test organisms 

to be included in the methods manual for sediment tests, 

it was generally agreed that no one method had to be 

used to culture organisms. The success of the tests 

should rely on the health of the culture from which the 

organisms were taken for testing. That is, having healthy 

organisms of known quality and age for testing was 

deemed to be the key consideration relative to culture 

procedures. Therefore, performance-based criteria were 

selected as the preferred approach laboratories should 

use to evaluate cultures rather than using control-based 

criteria. Performance criteria were chosen to allow each 

laboratory to optimize culture techniques and meet quality control monitoring requirements. 

A.6 The selection of test organisms for standardization 

were based on (1) current and historical acceptance, (2) 

logistical considerations, and (3) availability of testing 

methods. Major differences between freshwater and 

marine tests were discussed: (1) freshwater organisms 

are cultured in the laboratory and marine organisms are 

collected from the field, (2) freshwater organisms are 

smaller and younger, (3) freshwater organisms are generally epibenthic and marine test organisms are generally infaunal, (4) freshwater test conditions are sensitive 

to organic carbon, sediment oxygen demand, ammonia, 

and the buffering capacity of the overlying water. The 

overlying water is more stable in saltwater tests compared to freshwater tests. 

A.7 Performance-based culturing and testing criteria 

were outlined for freshwater species at the workshop 

(Table 11.3. 12.3, 13.4). Consensus was reached on 

critena that must be met and criteria that should be 

considered. Factors that must be met include reference 

toxicants with short-term water only exposures, survival 

of control organisms, age or size of organisms at the 

beginning or end of tests, and consistent water quality. 

Factors that should be considered include parental survival, food quakty, frequency of restarting cultures, and 

time to emergence (midges). In addition, a 

performance-based criterion for culture of L. variegatus 

to monitor is population doubling time (number of organisms). 

A.8 The following topics were discussed related to the 

use of Hyalella azteca and Chirunomus spp. in sediment 

toxicity tests: (1) Age of organisms used to start a test: 

Hyalella azteca: Organisms of age 0- to 14-d old are 

typically used to start a test. It may be best to test 

organisms 7- to 14-d old, since organisms 0- to 7-d old 

are difficult to recover from sediment. Chironomus 

tentans: 9- to 1 l-d old organisms should be tested. 

Chironomus riparius: 6- to 8-d old organisms should be 

tested; (2) Length of test: The length of the test agreed 

upon was 10 d with survival as the endpoint. Additional 

research is ongoing to evaluate growth as an endpoint; 

(3) Feeding: A minimal amount of food is required to 

consistently achieve adequate control survival and 

growth. Additional research is ongoing to evaluate the 

influence of feeding on sediment toxicity; (4) Water 

renewal: Limited renewal of overlying water was recommended (about 1 to 2 volume/d): (5) Sediment volumes: 

Sediment volume up to in the 200 mL have been routinely tested, but smaller volumes would be acceptable; 

(6) Grain size: There does not seem to be a substantial 

effect of sediment grain size in the 10-d exposures with 

H. azteca. Additional research is ongoing to evaluate 

the influence of grain size on the response of amphipods 

and midges; (7) Strains of organisms: Different strains of 

H. azteca have been used for testing. Reference toxicant comparisons of the strains are needed. 

A.9 The following topics were discussed related to the 

use of L. variegatus sediment bioaccumulation tests: (1) 

Age of test organisms: Adults should be tested; (2) 

Length of test: 28 d. Additional research is ongoing to 

evaluate duration of the exposure; (3) Feeding: No 

feeding is required during the test; (4) Water renewal: 

Limited renewal of overlying water was recommended 

(about 1 to 2 volume/d); (5) Sediment volumes: Sediment volume up to in the 200 mL have been routinely 

tested, but smaller volumes would be acceptable; (6) 

Grain size: There does not seem to be a substantial 

effect of sediment grain size in 1Od exposures with 

L. variegatus; (7) Additional discussion topics: Standard 

lipid content should be addressed in the document, 

sediment avoidance may be important, and rigorous 

techniques have been developed to purge the gut in 

clean water. Research is needed to see if purging is 

necessary. 

108 
Appendix B 

Exposure Systems 

B.1 Renewal of overlying water is recommended during 

sediment tests (Section 11.3, 12.3, 13.3). The overlying 

water can be replaced manually (e.g., siphoning) or 

automatically. Automated systems require more equipment and initially take more time to build, but manual 

addition of water takes more time during a test. In 

addition, automated systems generally result in less 

suspension of sediment compared to manual renewal of 

water. 

B.2 At any particular time during the test, flow rates 

through any two test chambers should not differ by more 

than 10%. Mount and Brungs (1967) diluters have been 

modified for sediment testing, and other diluter systems 

have also been used (Maki, 1977; Ingersoll and Nelson, 

1990; Benoit et al., 1993; Zumwalt et al., 1994). The 

water-delivery system should be calibrated before a test 

is started to verify that the system is functioning properly. Renewal of overlying water is started on Day -1 

before the addition of test organisms or food on Day 0. 

Water-delivery systems are described by Benoit et al. 

(1993) in Section B-3 and by Zumwalt et al. (1994) in 

Section 6.4. A 60-mL syringe with a mesh screen over 

the end can be used to manually remove and replace 

overlying water (J. Lazorchak, USEPA, Cincinnati, OH, 

personal communication). 

8.3 Benoit et al. (1993) describe a sediment testing 

intermittent-renewal (STIR) system (stationary or portable) for invertebrate toxicity testing with sediment. 

Either stationary or portable systems enable the maintenance of acceptable water quality (e.g., dissolved oxygen) by automatically renewing overlying water in sediment tests at rates ranging from 1 to 21 volume renewals/d. The STIR system not only reduces the labor 

associated with renewal of overlying water but also 

affords a gentle exchange of water that results in virtually no sediment suspension. Both gravity-operated systems can be installed in a compact vented enclosure. 

The STIR system has been used for conducting 10-d 

whole-sediment tests with Chironomus tentans, Hyalella 

azteca and Lumbriculus variegatus. 

8.3.1 STIR systems described in Benoit et al. (1982) 

can be modified to conduct sediment tests and at the 

same time maintain their original capacity to deliver 

varying concentrations of toxicants for water-only toxicity tests. A STIR system (stationary or portable) solely 

for sediment toxicity tests was designed, which offers a 

simple, inexpensive approach for the automated renewal of variable amounts of overlying water (Figures 

B.1 and 8.2). This system is described below. The 

system can be built as a two-unit system (Section 8.3.2) 

or with more exposure treatments (Section 8.3.4). All 

exposure systems consist of exposure holding tanks, 

head tanks, head tank support stands, and a water bath 

(Section B.3.2 and B.3.3). The automated delivery system includes design descriptions for a support stand, 

water renewal supply, and water delivery apparatus 

(Section 8.3.4). 

8.3.2 Two Unit Portable STIR System Construction 

(Figure B.1 and B.2) 

8.3.2.1 Exposure Holding Tanks (2) (Figure 8.3). 

1. Outer diameter: 15.8 cm wide x 29.3 cm long x 11.7 

cm high 

2. Cutting dimensions: (double strength glass, 3 mm) 

2 Bottoms: 15.8 cm x 29.3 cm 

4 Sides: 11.4 cm x 28.7 cm 

4 Ends: 11.4cm x 15.8cm 

3. Hole: 1.6 cm centered between sides and 7.2 cm 

from bottom edge of 11.4 cm high end piece. 

4. Standpipe Height: 10.3 cm above inside of tank 

bottom. 

8.3.2.2 Head Tanks (2) (4-L capacity; Figure 8.3) 

1. Outer diameter: 15.8 cm wide x 24 cm long x 14.5 

cm high 

2. Cutting dimensions: (acrylic plastic. 6 mm) 

2 Bottoms: 15.8 cm x 24 cm 

4 Sides: 13.9 cm x 22.8 cm 

4 Ends: 13.9 cm x 15.8 cm 

3. Acrylic plastic sheets should be cut with a smooth 

cutting fine toothed table saw blade. Dimension cut 

pieces can most easily be glued together with 

Weld-On® #16 clear-thickened cement for acrylic 

109 
Figure B.1 Portable table top STIR system described in Benoit et al. (1993). 

110 
Calibrated Volume Sight Tube 

(1.3cm Clear Tube) 

Head Box 

b y (30.5 x 30.5 x 3&m High 

d= A - Water Inlet 

1 1 ‘Timer Controlled Solenoid Val&s 

1 

Optional 

Automated z 

Water 

Delivery 

g 

Apparatus ,” n

Water Distribution p 

II II II II I 

Manifold with Open Ends 

s 6mm Pipe to Hose Adaptor 

(1.3cm plastic pipe) 

Water Bath 

u LJ IJ u u u 

f 

Thermostat All tanks and water bath drain to common 19L jug with air 

vent and optional hose from jug to floor drain. 

Figure 8.2 Portable table top STJR system with several additional options as described in Benoit et al. (1993). 

111 
Width (end) Width (end) 

Exposure Holding Tank Head Tank 

I A _..--- 3.2cm 

1 ,\;/f Thermostat 

Length (side) 

Basic Water Bath 

Basic Water Bath with Optional Holes for Water Bath 

Width (end) 

Add on Water Bath for One Additional Unit 

Figure 8.3 Tanks for the STIR system in Benolt et al. (1993). 

112 
plastic (Industrial Polychemical Service, P.O. Box 

471, Gardena, CA, 90247). 

4. Hole: 1.6 cm centered between sides and 2 cm from 

front edge of 24 cm long bottom piece. Holes can 

most easily be drilled in acrylic plastic by using a 

wood spade bit and drill press. 

5. Flow Tubes: IO-mL pipet tip initially cut off at the 6 

mL mark and inserted flush with top of #0 stopper. 

Top of stopper should be inserted nearly flush with 

head tank bottom. With 2 L of water in head tank, 

calibrate flow tube to deliver 32 mlimin. 

8.3.2.3 Head Tank Support Stand (1) (Figure B.3) 

1. Outer diameter: 16.7 cm wide x 33.7 cm long x 17.8 

cm high 

2. Cutting dimensions: (acrylic plastic, 6 mm) 

1 Bottom: 16.7 cm x 33.7 cm 

2 Sides: 17.2 cm x 32.5 cm 

2 Ends: 17.2 cm x 16.7 cm 

3. Size is such that both head tanks fit into support 

stand for storage and transport. 

8.3.2.4 Water Bath (1) (Figure B.3) 

1. Outer diameter: 33 cm wide x 40.6 cm long x 7.4 cm 

high 

2. Cutting dimensions: (acrylic plastic, 6 mm) 

1 Bottom: 33 cm x 55.9 cm 

2 Ends: 33 cm x 6.8 cm 

2 Sides: 39.4 cm x 6.8 cm 

3. Holes: 

a. Overflow drain; 1.6 cm centered 2.9 cm from 

bottom edge of 39.4 cm long side piece and 

17.8 cm from right edge. 

b. Thermostat; 3.2 cm centered 2.5 cm from bottom edge of 39.4 cm long side piece and 3.2 cm 

from left edge. 

c. Water pump outlet; 2.5 cm centered 2.5 cm 

from bottom edge of 33 cm long end piece and 

8.3 cm from back edge. 

d. Water pump inlet; 2.5 cm centered 2.5 cm from 

bottom edge of 33 cm long end piece and 

2.0 cm from back edge. 

4. A small 90” elbow made of glass or plastic is attached to the water pump inlet tube and turned 

downward so the circulator pump will not pick up air 

at the water surface. 

5. The bottom piece for the water bath includes 15.3 

cm extension for motor mount and the thermostat 

electrical junction box. 

6. Motor Mount: 5.1 cm wide x 11.4 cm long x 3.8 cm 

thick mount made from 6 pieces of 6-mm acrylic 

plastic. Four of these pieces are glued together. The 

other two pieces are glued together, motor attached 

to the edge with two screws and the two pieces (with 

motor attached) are then screwed to the top of the 

four pieces. The entire unit is then glued to water 

bath extension after 6-mm PVC piping is attached 

and secured with stoppers to the inlet and outlet 

water bath holes. 

7. Thermostat Conduit Junction Box: (1.3.cm small left 

back (SLB)) is attached to the water bath extension 

by screwing a 1.3.cm PVC plug into junction box 

and securing this plug with a screw, countersunk up 

through the bottom and into the PVC plug. 

8.3.2.5 Latex Rubber Mold: If you plan to construct a 

substantial number of exposure test beakers, as described in Benoit et al. (1993), then it would be to your 

advantage to make a latex rubber mold to give support 

to the underside of the glass when drilling holes. It 

significantly reduces the number of broken beakers. 

Liquid latex, with hardener that can be purchased from 

the local hardware store is commonly used to coat the 

handles of tools- The rubber mold is constructed as 

follows: 

1. Mix latex with hardener as per instructions. 

2. Fill one exposure test beaker with the mixture. 

3. Suspend one 5 cm eye bolt (5 mm diameter) with 

nut on end so that the eye is protruding just above 

the top of the mixture. 

4. Allow the latex plenty of time to “set up.” 

5. With proper eye protection and wearing heavy gloves, 

gently break the beaker with a small hammer and 

remove all of the glass from the mold. 

6. Using a long drill bit for wood, drill an air vent hole 

through the mold from top through bottom. 

7. When using the mold, wet the mold and the beaker 

with water before inserting. Place the beaker, with 

pre-marked location of holes, on its side in a 3.5-L 

stainless steel pan filled with coolant water so that 

the beaker is just below the surface. The beaker is 

then held in position with one hand while the other 

hand operates the drill press. Operator should wear 

proper eye protection. 

8. After the two holes are drilled, the mold can be 

easily removed, with some effort, by inserting the 

eye bolt into the handle of a securely attached “C” 

113 
clamp and physically pulling the beaker from the 

mold. 

8.33 Suggested options for more exposure treatments 

(examples given are for a three unit treatment system) 

8.3.3.1 Exposure Holding Tanks and Head Tanks: 

Same dimensions as for two unit system except that 

three (3) of each should be made. 

B.3.3.2 Head Tank Support Stand (1) (Figure 8.3) 

1. Outer diameter: 16.7 cm wide x 49.5 cm long x 17.8 

cm high 

2. Cutting dimensions:(acrylic plastic, 6 mm) 

1 Bottom: 16.7 cm x 49.5 cm 

2 Sides: 17.2 cm x 48.3 cm 

2 Ends: 17.2 cm x 16.7 cm 

3. Size is such that the three head tanks will fit into the 

support stand for storage and transport. 

8.3.3.3 Water Bath (1) (Figure 8.3) 

1. Outer diameter: 33 cm wide x 56.4 cm long x 7.4 cm 

high 

2. Cutting dimensions:(acrylic plastic, 6 mm) 

1 Bottom: 33 cm x 71.7 cm 

2 Ends: 33 cm x 6.8 cm 

2 Sides: 55 cm x 6.8 cm 

3. Holes: All hole sizes and locations are the same as 

for the two unit system except that overflow drain is 

located 25.7 cm from right edge of 55-cm side. Also, 

two optional 1.6-cm holes centered 2.5 cm from 

bottom edge of 33cm long end piece and 1.8 cm 

from corner edges are shown in the drawing for 

future additions of “add-on” water baths. 

4. Motor mount and junction box installation are the 

same as for two unit system. 

B.3.3.4 “Add-On” Water Bath (example given is for one 

additional unit treatment system; Figure 8.3) 

1. Outer diameter: 18.5 cm wide x 33 cm long x 8 cm 

high 

2. Cutting dimensions: (acrylic plastic, 6 mm) 

1 Bottom: 18.5cmx33cm 

2 Ends: 17.3 cm x 7.4 cm 

2 Sides: 33 cm x 7.4 cm 

3. Holes: Inlet and outlet holes (1.6 cm) are centered 

2.5 cm from bottom edge of 33-cm long side piece 

and 1.8 cm from corner edges. 

4. The above holes will match the previously drilled 

holes in the main water bath. The “add-on” water 

bath is connected using #2 stoppers and 6.4 cm 

lengths of clear plastic tubing (1.3 cm diameter). 

The circulator pump outlet tubing (Tygon@) in the 

main water bath is extended through the inlet connection as shown in Figure 8.2. Circulating water is 

then forced into the “add-on” bath and flows back to 

the main water bath by gravity. 

5. Note that the walls of the “add-on” bath are 6 mm 

higher than the main water bath to accommodate 

the small head of water that builds up. 

6. “Add-on” water baths tend to run a little warmer 

(0.2%) than main water bath test temperatures. 

B-3.4 Optional Automated Water Delivery Apparatus 

For Table Top STIR Systems (examples given are for a 

three unit treatment system) 

B.3.4.1 Support Stand: A stand to support the automated water delivery apparatus, shown in Figure 8.2, 

can be made from bolted slotted angle iron bolted with 

corner braces. A convenient size to construct is 30 cm 

wide x 85 cm long x 43 cm high. The head box in 

Figure 8.2 sits on top of the stand, and the water distribution manifold as shown in Figure B.2 is placed directly 

under the top of the stand with two 1.3 cm conduit 

hangers. A small portion of each angle iron cross piece 

is cut away to allow the pipe to be clamped into the 

conduit hanger. This also keeps the manifold up high 

enough for sufficient clearance between the head tanks 

and the 6-mm pipe to hose adapters as shown in 

Figure 8.2. 

8.3.4.2 Water Renewal Supply: If tests will be conducted in the local water supply, then the head box 

water inlet shown in Figure B.2 is simply plumbed into 

the supply line. However, if the tests are conducted with 

transported water or with reconstituted water, the head 

box water inlet can be connected to a Nalgene” drum 

with flexible Tygon@ tubing. With a four volume test 

beaker water renewal flow rate per day, both 114-L and 

208-L Nalgene* drums will hold a 5-d supply for a 3-unit 

treatment system and a 5-unit treatment system, respectively. If the water supply drum is located below the 

head box, then an open air water pump such as a 

March@ model MDXT pump (PFC Equipment Corp., 

Minneapolis, MN 55440) can be used between the drum 

and head box. 

8.3.4.3 Operation of Water Delivery Apparatus: The 

head box water inlet solenoid valve (Figure 8.2) and the 

open air water pump (if needed) are connected to the 

same timer control switch. The head box water outlet 

solenoid valve is connected to another separate timer 

control switch. With four test beaker renewals/d and a 

3-unit treatment system, the head box toilet float valve is 

pre-adjusted to allow the head box to fill to the 12-L mark 

on the sight tube (Figure 8.2). 

114 
B-3.4.3.1 With head box filled, the renewal cycle begins 

when the first timer opens the head box outlet solenoid 

valve. The distribution manifold is quickly flooded and 

the 12 L of renewal water divided equally to each of the 

three 4-L head tanks. Since the timers have a minimum 

setting of one hour on-off periods, the first timer is set to 

shut off the head box outlet solenoid valve one hour 

after it opens. 

B.3.4.3.2 About 30 min later, the second timer is set to 

open the head box water inlet solenoid valve (and pump 

if needed). As head box water volume reaches the 12-L 

mark, the pre-adjusted toilet tank valve stops the water 

ffow. One hour after they come on, the second timer will 

shut off the solenoid valve inlet and water pump. 

B.3.4.3.3 The automated system is then ready for the 

next renewal cycle that is set to begin 12 h after the first 

cycle. Head box volume dimensions are such that up to 

five unit treatment systems can be tested simultaneously 

as shown in Figure B.2. 

B.3.5 A criticism of the system described by Benoit et al. 

(1993) is that the (up to) 8 beakers placed in each 

holding tank are not true replicates because of the 

potential for exchange of water overlying the sediments 

among the beakers. However, this concern is largely 

semantic with regard to actual test results. The rationale 

for this position is described below. The data described 

below are unpublished data from ERL-Duluth (G.T. 

Ankley, USEPA, Duluth, MN, personal communication). 

B.3.5.1 Beakers within a test tank should contain an 

aliquot of the same homogenized sediment and the 

same test species. The replication is intended to reflect 

variability in the biology (e.g., health) of the organism, as 

well as placement and recovery of the animals from the 

test sediments (i.e., operator variability). To treat even 

completely separate tanks containing homogenized sediment from the same source as true replicates (of the 

sediment ‘treatment”) is inaccurate and is pseudoreplication. Hence, because the same sediment is tested in 

each beaker in a particular tank, and because the replication is focused on defining variability in the biology of 

the organism (and the operator), this is essentially a 

non-issue from a theoretical standpoint. 

8.3.5.2 From a practical standpoint, it is important to 

determine the potential influence of one beaker on another over the course of a test. To determine this, a 

study was designed (which is not advocated) in which 

treatments were mixed within a tank. In the first experiment, four beakers of highly metal-contaminated sediment from the Keweenaw Waterway, MI, were placed in 

the same tank as four beakers containing clean sediment from West Bearskin Lake, MN. This was done in 

two tanks; in one tank, 10 amphipods (Hyaleila azteca) 

were added to each beaker, while in the other tank, 10 

midges (Chironomus tentans) were placed in each beaker. Controls for the experiment consisted of the West 

Bearskin sediments assayed in separate “clean” tanks. 

The four contaminated beakers were placed “upstream” 

Table 8.1 Sediment Copper Concentrations and Organism 

Survival and Growth at the End of a 10-d Test with 

West Bearskin Sediment in an Individual Tank 

Versus 10-d Cu Concentrations and Organism 

Survival and Growth In West Bearskin Sediment 

Tested in the Same Tank as Keweenaw Waterway 

Sediment’ 

Survival Dry wt 

Sediment Tank Species W) (mg/organism) ($g) 

WB2 1 Amphipod 90 ND3 22.4 

WB 2 Amphipod 100 ND 13.6 

KW4 2 Amphlpod 20 ND 9397.0 

WB 3 Midge 95 1.34 12.3 

WB 4 Mtctge 100 1.33 15.6 

KW 4 Midge 5 ND 9167.0 

’ All values are the mean of duplicate observations (G.T. Ankley. 

USEPA, Duluth, MN. unpublished data) 

2 West Bearskin 

3 Not determrned 

4 Keweenaw Waterway 

of the four clean beakers to attempt to maximize possible exchange of contaminant. At the end of the test, 

organism survival (and growth in for C. tenfans) was 

measured in two of the beakers from each site and 

sediment Cu concentrations were determined in the 

other two beakers from each site. The Keweenaw sediments contained concentrations of Cu in excess of 

9,000 pg/g (dry wt), and were toxic to both test species 

(Table B-1). Conversely, survival of both C. tentans and 

H. azteca was high in the West Bearskin sediments from 

the Keweenaw tank, and was similar to survival in West 

Bearskin sediments held in separate tanks. Most important, there was no apparent increase in Cu concentrations in the West Bearskin sediments held in the 

Keweenaw tank (Table 8.1). 

B.3.5.3 A similar design was used to determine transfer 

of contaminants among beakers containing sediments 

spiked with the organochlorine pesticide dieldrin. In this 

experiment, sediment from Airport Pond, MN, was spiked 

with dieldrin and placed in the same tank as clean 

unspiked Airport Pond sediments. Two different concentrations were assayed: (1) in the midge test sediment 

concentrations were about 150 j@ dieldrin/g (dry weight) 

and (2) in the amphipod test sediments contained in 

excess of 450 rrg dieldrin/g sediment. The control for the 

experiment again consisted of clean Airport Pond sediment held in a separate tank. The spiked sediments 

were toxic to both test species, and survival of organisms held in the clean Airport Pond sediments was 

similar in the two different tanks. However, there was an 

effect on the growth of C. tenfans from the clean Airport 

Pond sediment assayed in the tank containing the spiked 

sediment. This corresponded to the presence of measurable dieldrin concentrations in unspiked Airport Pond 

sediments in the tank with the mixed treatments 

115 
Table 8.2 

Sediment 

-- - 

AP2 

AP 

DAP4 

AP 

AP 

DAP 

- 

Sediment Dietdrin Concentrations and Organism 

Survival and Growth at the End of a 10-d Test with 

Airport Pond Sediment in an individual Tank Versus 

t O-d DLeldrin Concentrations and Organism Sun&al 

and Growth in Airport Pond Sediment Tested in the 

Same Tank as Dieldren-spiked Airport Pond 

Sediment’ 

Surwval Dry WI Dieldrin 

Tank Species (“4 (m@orgamsm) Wcl) 

1 Amphlpod 75 ND3 CO.01 

2 Amphlpod 80 ND 0.07 

2 Amphlpod 20 ND 446.4 

3 Midge 85 1.71 cu.01 

4 Mtdge 85 0.13 0.04 

4 Mtdge 0 ND 151.9 

1 All values are the mean of duplicate observations (G.T. Ankley, 

USEPA. Duluth. MN, unpublished data) 

2 Aqorl Pond 

3 Not determined 

’ Dleldren-spiked Airport Pond 

(Table B.2). The concentrations of dieldrin in the unspiked sediment, although detectable, were on the order 

of 5,000-fold lower than the spiked sediments, indicating 

relatively minimal transfer of pesticide. 

8.3.5.4 Using a similar design, an investigation was 

made to evaluate if extremely low dissolved oxygen 

(DO) concentrations, due to sediment oxygen demand, 

in four beakers in a test system would result in a 

decrease in DO in other beakers in the tank. In this 

experiment, trout chow was added to each of four beakers containing clean Pequaywan Lake sediment, and 

placed in a test tank with four beakers containing 

Pequaywan Lake sediment without exogenous organic 

carbon. Again, the control consisted of Pequaywan Lake 

sediment held in a separate tank under otherwise identical test conditions. Assays were conducted, without 

organisms, for 10 d. At this time, DO concentrations 

were very low in the beakers containing trout chowamended sediment (ca., 1 mg/L, n = 4). However, overlying water DO concentration in the “untreated” vs. the 

“treated” beakers in a separate tank, i.e., 6.8 vs. 6.9 mg/ 

L. respectively. This indicates, that from a practical 

standpoint, even under extreme conditions of mixed 

treatments (which again, is not recommended), interaction between beakers within a tank is minimal. 

8.3.5.5 One final observation germane to this issue is 

worth noting. If indeed beakers of homogenized sediment within a test tank do not serve as suitable replicates, this should be manifested by a lack of variability 

among beakers with regard to biological assay results. 

This has not proven to be the case. For example, in a 

recent amphipod test with a homogenized sediment 

from the Keweenaw Waterway in which all eight replicates were held in the same tank; mean survival for the 

test was 76%; however, survival in the various beakers 

ranged from 30 to lOO%, with a standard deviation of 

21%. Clearly, if the test system were biased so as to 

reduce variability (i.e., result in unsuitable replicates due 

to common overlying water), this type of result would not 

be expected. 

B.3.5.6 In summary, in both a theoretical and practical 

sense, use of the system described by Benoit et al. 

(1993) results in valid replicates that enable the evaluation of variability due to factors related to differences in 

organism biology and operator effects. To achieve this, 

it is important that treatments not be mixed within a tank; 

rather, the replicates should be generated from the 

same sediment sample. Given this, and the fact that it is 

difficult to document interaction between beakers using 

even unrealistic (and unrecommended) designs, leads 

to the conclusion that variability of replicates from the 

test system can be validly used for hypothesis testing. 

8.4 Zumwalt et al. (1994) also describe a water-delivery 

system that can accurately deliver small volumes of 

water (50 ml/cycle) to eight 300-mL beakers to conduct 

sediment tests. The system was designed to be comparable with the system described by Benoit et al. (1993). 

8.4.1 Eight 35-mL polypropylene syringes equipped 

with 18-gauge needles are suspended from a splitting 

chamber (Figure 8.4). The system is suspended above 

eight beakers and about 1 L of water/cycle is delivered 

manually or automatically to the splitting chamber. Each 

syringe fills and empties 50 mL into each beaker and the 

600 mL of excess water empties out an overflow in the 

splitting chamber (Section 8.4.3.1). The volume of water 

delivered per day can be adjusted by changing either 

the cycling rate or the size of the syringes. The system 

has been used to renew overlying water in 

whole-sediment toxicity tests with H. azteca and 

C. tentans. Variation in delivery of water among 24 

beakers was less than 5%. The system is inexpensive 

(41 00), easy to build (4 h), and easy to calibrate (~15 

min). 

8.4.2 Water Splitting Chamber 

8.4.2.1 The glass water-splitting chamber is 14.5 cm 

wide, 30 cm long, and 6.5 cm high (inner diameter). 

Eight 3.8-cm holes and one 2.5-cm hole are drilled in a 

15.5 cm x 30.5 cm glass bottom before assembly 

(Figure 8.4 and Table B-3). The glass bottom is made 

from 4.8- (3/16 inch) or 6.4 mm (l/4 inch) plate glass. An 

easy way to position the 3.8-cm holes is to place the 

eight 300- mL beakers (2 wide x 4 long) under the 

bottom plate and mark the center of each beaker. The 

2.5cm hole for overflow is centered at one end of the 

bottom plate between the last two holes and endplate 

(Figure 6.4). After drilling the holes in the bottom plate, 

the side (6.5 x 30.5 cm) and end (6.5 x 14.5 cm) plates 

are cut from 3.2-mm (l/8 inch) double-strength glass 

and the splitting box is assembled using silicone adhesive. Sharp glass edges should be sanded smooth using 

a whetstone or a piece of Carborundum wheel. After the 

116 
Splitter Chamber 

I 

Overview 

Sditter Chamber 

Wing Nut 

Waterbath (Optional) 

Figure 6.4 Water splitting chamber described in fumwalt et al. (1994). 

117 
Table 6.3 Materials Needed for Constructing a Zumwalt et al. 

(1994) Delivery System 

Equipment 

Drill press 

Glass drill bits (2.54 cm [l Inch] and 3.8 cm 11.5 Inch]) 

Cork bonng set 

Table-top saw equrpped with a Carborundum wheel 

Small level (about 30 cm long) 

Supplies 

300-mL beakers (Itpless. tall form; e.g.. Pyrex Model 1040) 

Stainless-steel screen (50- x 50-mesh) 

9.5 mm (3/8 Inch x 16) stainless-steel threaded rod 

9.5 mm (3/8 Inch x 16) nylon wingnuts 

9.5 mm (3/8 inch x 16) nylon nuts 

35 mL Mono-ject syringes (Sherwood Medical, St. Louis, MO) 

18-gauge Mono-ject stainless-steel hypodermic needles 

Srlrcone stoppers (#O. 5. and 7) 

Plate glass (6.4 mm [l/4 inch], 4.8 mm [3/l 6 inch], 3.2 mm [l/8 inch]) 

Glass tubmg (8 mm outer diameter) 

Stainless-steel tubing (12 mm outer diameter) 

SlItcone adhesive (without fungicide) 

Sway stainless-steel gang valves and 

Pasteur pipets (14.5 cm 15.75 inch]) 

splitting chamber has dried for 24 h, four 12-mm (outer 

diameter) stainless-steel tubes (7 cm long) are glued to 

each corner of the splitting chamber (the surface of the 

steel tubes is scored with rough emery paper to allow 

better adhesion of the silicone). These tubes are used 

as sleeves for attaching the legs to the splitting chamber. The legs of the splitting chamber are threaded 

stainless-steel rods (9.5 mm [3/8 inch] diameter, 36 cm 

long). The location of the tubes depends on the way that 

the beakers are to be accessed in the waterbath. If the 

tubes are placed on the side of the splitting chamber, a 

3.2-mm-thick x 2cm-wide x 7cm-long spacer is required so beakers and the optional waterbath can be slid 

out the ends (Figure 8.4). If the sleeves and legs are 

attached to the ends of the splitting chamber, the beakers and waterbath can be removed from the side. The 

legs are inserted into the 12-mm tubes and secured 

using nylon nuts or wingnuts. The distance between the 

tips of the needles to the surface of the water in the 

300-mL beakers is about 2 cm. Four 1-L beakers could 

also be placed under the splitting chamber. 

B.4.2.2 A #7 silicone stopper drilled with a 21-mm (outer 

diameter) core borer is used to hold each 35mL polypropylene syringe (45 mL total capacity) in place. Glass 

syringes could be used if adsorption of contaminants on 

the surface of the syringe is of concern. A dilute soap 

solution can be used to help slide the syringe into the #7 

stopper (until the end of the syringe is flush with the top 

of stopper). Stoppers and syringes are inserted into 

3.8-cm holes and are visually leveled. A #5 silicone 

stopper drilled with an 8 mm (outer diameter) core borer 

is placed in the 2.5 cm overflow hole. An 8-mm (outer 

diameter) glass tube (7.5 cm long) is inserted into the 

stopper. Only 3 mm of the overflow tube should be left 

exposed above the stopper. This overflow drain is placed 

about 3 mm lower than the top of the syringes. A short 

piece of 6.4-mm (l/4 inch; inner diameter) tubing can be 

placed on the lower end of drain to collect excess water 

from the overflow. 

8.4.2.3 The splitting chamber is leveled by placing a 

level on top of the chamber and adjusting the nylon nuts. 

Eighteen-gauge needles are attached to the syringes. 

About 6 mm of the needle should remain after the sharp 

tip has been cut off using a carborundum wheel. Jagged 

edges left in the bore of the needle can be smoothed 

using a small sewing needle or stainless-steel wire. 

8.4.2.4 When about 1 L of water is delivered to the 

splitting chamber, the top of each syringe should be 

quickly covered with water. The overflow tube will quickly 

drain excess water to a level just below the tops of the 

syringes. The syringes should empty completely in about 

4 min. If water remains in a syringe, the needle should 

be checked to ensure that it is clean and does not have 

any jagged edges. 

8.4.3 Calibration and Delivery of Water to the Splitting 

Chamber. Flow adjustments can be made by sliding 

either the stoppers or syringes up or down to deliver 

more or less water. A splitting chamber with eight syringes can be calibrated in less than 15 min. Delivery of 

water to the splitting chamber can be as simple as 

manually adding about 1 L of water/cycle. Water can be 

added automatically to the splitting chamber using a 

single cell or a Mount and Brungs (1967) diluter that 

delivers about 1 L/cycle on a time delay. About 50 mL 

will be delivered to each of the 8 beakers/cycle and 

600 mL will flow out the overflow. A minimum of about 1 

L/cycle should be dumped into the splitting chamber to 

ensure each syringe fills to the top. If the quantity of 

water is limited at a laboratory, the excess water that 

drains through the overflow can be collected and recycled. 

B.4.4 Waterbath and Exposure Beakers. The optional 

waterbath surrounding the beakers is made from 3.2-mm 

(l/&inch) double-strength glass and is 15.8 cm wide x 

29.5 cm long x 11.7 cm high (Figure B.4 [Figure 8.3 in 

the Benoit et al., 1993 system]). Before the pieces are 

assembled, a 1.4-cm hole is drilled in one of the end 

pieces. The hole is 7.2 cm from the bottom and centered 

between each side of the end piece. A glass tube 

inserted through a ##O silicone stopper can be used to 

drain water from the waterbath. A notch is made in each 

300-mL beaker by making two cuts with a Carborundum 

wheel 1.9 cm apart to the 275 mL level. The beaker is 

etched across the bottom of the cuts, gently tapped to 

remove the cut section, and the notch is covered with 

50- x XI-mesh stainless-steel screen using silicone adhesive. The waterbath illustrated in Figure B.4 is optional if the splitting chambers and beakers are placed in 

a larger waterbath to collect waste water. This smaller 

waterbath could be used to collect waste water and a 

surrounding larger waterbath could be used for temperature control. 

118 
B.4.4.5 Operation and Maintenance 

B.4.4.5.1 Maintenance of the system is minimal. The 

syringes should be checked daily to make sure that all of 

the water is emptying with each cycle. As long as the 

syringe empties completely, the rate of flow out of the 

syringes is not important because a set volume of water 

is delivered from each syringe. If the syringe does not 

empty completely with each cycle, the needle tip should 

be replaced or cleaned with a thin wire or sewing 

needle. If the screens on the beakers need to be cleaned, 

a toothbrush can be used to brush the outside of screens. 

B.4.4.5.2 Overlying water can be aerated by suspending Pasteur pipets (e.g., Pyrex disposable 14.5cm I5.75 

inch) length) about 3 cm above the sediment surface in 

the beakers. Five-way stainless steel gang valves are 

suspended from the splitting chamber using stainless 

steel hooks. Latex tubing (3.2-mm [l/8 inch] inner diameter) is used to connect valves and pipets. Flow rate of 

air should be maintained at about 2 to 3 bubbles/s and 

the pipets can be placed on the outside of the beakers 

when samples of overlying water are taken during a test. 

B-4.4.5.3 The splitting chambers were used to deliver 

water in a toxicity test with the midge Chironomus 

tentans exposed to metal-contaminated sediments 

(Zumwalt et al., 1994). Ten third-instar midges were 

exposed in 300-mL beakers containing 100 mL of sediment and 175 mL of overlying water at 23°C. Midges in 

each beaker received a daily suspension of 4 mg 

Tetrafin@ flake food and survival and growth were measured after 10 d. Splitting chambers delivered 50 mL/ 

cycle of overlying water to each of the eight replicate 

beakers/sediment sample. One liter of water was delivered with a single-cell diluter to each splitting chamber 4 

times/d. This cycle rate resulted in 1 .l volume additions 

of overlying water/d to each beaker ([4 cycles/d x 50-mL 

volume/cycle]/1 75 mL of overlying water). The variation 

in delivery of water between 24 beakers was less than 

5%. 

8.4.4.5.4 Hardness, alkalinity, and conductivity in water 

overlying the sediments averaged about 20% higher 

than inflowing water. These water-quality characteristics 

tended to be more similar to inflowing water at the end of 

the exposure compared with the beginning of the exposure. The average pH was about 0.3 units lower than 

inflowing water. Ammonia in overlying water ranged 

from 0.20 to 0.83 mg/L. The dissolved oxygen content 

was about 1 mg/L lower than inflowing water at the 

beginning of the exposure and was about 2 to 3 mg/L 

lower than inflowing water by the end of the exposure. 

Survival and growth of midges were reduced with exposure to metal-contaminated sediments. Water delivered 

at a similar rate to a second set of beakers using a 

system described by Benoit et al. (1993) resulted in 

similar overlying water quality and similar toxic effects 

on midges. 

8.4.4.5.5 The system has been used to deliver 33 %O 

salt water to exposure chambers for 10 d. Precipitation 

of salts on the tips of the needles reduced flow from the 

syringes. Use of a larger bore needle (1 &gauge) reduced clogging problems; however, daily brushing of the 

needle tips is required. Use of larger bore needles with 

300-mL beakers containing 100 mL of sediment and 

175 mL of overlying water results in some suspension of 

sediment in the overlying water. This suspension of 

sediment can be eliminated if the stream of water from 

the larger bore needle falls on a baffle (e.g., a piece of 

glass) at the surface of the water in the beaker. 

119 
Appendix C 

Food Preparation 

C.1 Yeast, Cerophyl® and Trout Chow (YCT) for 

feeding the cultures and Hyalella azteca. Food should 

be stored at 4°C and used within two weeks from 

preparation; however, once prepared, YCT can be frozen until use. 

C.1.1 Digested trout chow is prepared as follows: 

1. Preparation of trout chow requires one week. 

Use starter or No. 1 pellets prepared according 

to current U.S. Fish and Wildlife Service specifications. Suppliers of trout chow include Zeigler 

Bros., Inc., P.O. Box 95, Gardners, PA, 17324 

(717/780-9009); Glencoe Mills, 1011 Elliott, 

Glencoe, MN, 55336 (612/864-3181); and Murray 

Elevators 118 West 4800 South, Murray, UT 

84107 (800/521-9092). 

2. Add 5.0 g of trout chow pellets to 1 L of deionized water. Mix well in a blender and pour into a 

2-L separatory funnel or similar container. Digest before use by aerating continuously from 

the bottom of the vessel for one week at ambient laboratory temperature. Water lost due to 

evaporation is replaced during digestion. Because of the offensive odor usually produced 

during digestion, the vessel should be placed in 

a ventilated area. 

3. At the end of digestion period allow material to 

settle for a minimum of 1 h. Filter the supernatant through a fine mesh screen (e.g., Nitex® 

110 mesh). Combine with equal volumes of the 

supernatant from Cerophyl® and yeast preparation (below). The supernatant can be used fresh, 

or it can be frozen until use. Discard the remaining particulate material. 

C.1.2 Yeast is prepared as follows: 

1. Add 5.0 g of dry yeast, such as Fleishmann’s® 

Yeast, Lake State Kosher Certified Yeast, or 

equivalent, to 1 L of deionized water. 

2. Stir with a magnetic stirrer, shake vigorously by 

hand, or mix with a blender at low speed, until 

the yeast is well dispersed. 

3. Combine the yeast suspension immediately (do 

not allow to settle) with equal volumes of supernatant from the trout chow (above) and 

CerophyI® preparations (below). Discard excess material. 

C.1.3 Cerophyl® is prepared as follows: 

1. Place 5.0 g of dried, powdered, cereal or alfalfa 

leaves, or rabbit pellets, in a blender. Cereal 

leaves are available as “Cereal Leaves,” from 

Sigma Chemical Company, P.O. Box 14508, St. 

Louis, MO, 63178 (8001325-3010); or as 

Cerophyl® from Ward’s Natural Science Establishment, Inc., P.O. Box 92912, Rochester, NY, 

14692-9012 (716/359-2502). Dried, powdered, 

alfalfa leaves may be obtained from health food 

stores, and rabbit pellets are available at pet 

shops. 

2. Add 1 L of deionized water. 

3. Mix in a blender at high speed for 5 min, or stir 

overnight at medium speed on a magnetic stir 

plate. 

4. If a blender is used to suspend the material, 

place in a refrigerator overnight to settle. If a 

magnetic stirrer is used, allow to settle for 1 h. 

Decant the supernatant and combine with equal 

volumes of supernatant from trout chow and 

yeast preparations (above). Discard excess 

material. 

C.1.4 Combined yeast-cerophyl-trout chow (YCT) is 

mixed as follows: 

1. Thoroughly mix equal (e.g., 300 mL) volumes of 

the three foods as described above. 

2. Place aliquots of the mixture in small (50 mL to 

100 mL) screw-cap plastic bottles. 

3. Freshly prepared food can be used immediately, or it can be frozen until needed. Thawed 

food is stored in the refrigerator between feedings and is used for a maximum of two weeks. 

Do not store YCT frozen over three months. 

120 
4. It is advisable to measure the dry weight of 

solids in each batch of YCT before use. The 

food should contain 1.7 to 1.9 g solids/L. Cultures are fed IO mUL on Monday and 5 mL/L on 

Wednesday and Friday (USEPA, 1993). 

C.2 Algal Food: Starter cultures of the green algae, 

Selenastrum capricomufum and the diatom Navicula (or 

Synedra) are available from the following sources: American Type Culture Collection (Culture No. ATCC 22662), 

12301 Parklawn Drive, Rockville, MD 10852, or Culture 

Collection of Algae, Botany Department, University of 

Texas, Austin, TX 78712. 

C.2.1 Algal Culture Medium for the green algae and 

diatoms (Navicula or Synedra) prepared as follows 

(USEPA, 1993a): 

Table C.l Nutrient Stock Solutions for Maintaining Algal Stock 

Cultures 

Stock Compound Amount dissolved in 

solutron 500 mL deionized water 

1. Macronutnents 

A. MgCI,*6H,O 6.08 g 

CaCl *2H,O 

NaN& 

2.20 g 

12.75 g 

a. MgSO,*7H,O 7.35 g 

C. K,H PO, 0.522 g 

0. NaHCO, 7.50 g 

2. Micronutrients 

HP% 92.8 mg 

MnCI,*4H,O 268.0 mg 

ZnCI, 1.64 mgl 

FeCI,.GH,O 79.9 mg 

CoCI,.GH,O 0.714 mg2 

Naj~Io0,.2H,O 3.63 mg3 

CuCI,.PH,O 0.006 mg4 

N~QEDTA*PH,O 150.0 mg 

Na$eO, 1.196 mg5 

rZnCI,-Weigh out 164 mg and dilute to 100 mL. Add 1 mL of this 

solution to micronutrient stock. 

*CoCI,~GH,O-Weighout .4mganddrluteto 100 mL.Add 1 mLofthis 

solution to micronutrient stock. 

3N%Mo04.2H,0-Wergh out 36.6 mg and dilute to 10 mL. Add 1 mL of 

this solutton to micronutnent stock. 

4CuCI,*2H,0-Weigh out 60.0 mg and dilute to 1000 mL. Take 1 mL of 

this solution and dilute to 10 mL. Take 1 mL of the second dilution and 

add to micronutrient stock. 

5N a2 GO,-Weigh out 119.6 mg and dilute to 100 mL. Add 1 mLof this 

solution to micronutrient stock. 

Table C.2 Final Concentration of Macronutrients and Micronutrients in the Algal Culture Medium 

Macronutnent Concentratron Element Concentration 

VwN-1 OWL) 

NaNO, 25.5 N 4.20 

MgCI,*6H,O 12.2 Mg 2.90 

CaCI,*2H,O 4.41 Ca 1.20 

MgS04*7H20 14.7 S 1.91 

K,HPO, 1.04 P 0.186 

NaHCO, 15.0 Na 11.0 

K 0.469 

C 2.14 

Micronutrient Concentratron Element Concentration 

bw) wu 

HP4 185 B 32.5 

MnCI,*4H,O 416 Mn 115 

ZnCI, 3.27 Zn 1.57 

CoCI,~GH,O 1.43 co 0.354 

CUCI,~PH,O 0.012 cu 0.004 

N%MoO,*2H,O 7.26 MO 2.88 

FeCI,.GH,O 160 Fe 33.1 

N%EDTA*2H,O 300 - - 

es~4 2.39 Se 0.91 

1. 

2. 

3. 

4. 

Prepare stock nutrient solutions using reagent 

grade chemicals as described in Table C.l. 

Add 1 mL of each stock solution, in the order 

listed in Table C.1, to about 900 mL of deionized water. Mix well after the addition of each 

solution. Dilute to 1 L, mix well. The final concentration of macronutrients and micronutrients 

in the culture medium is listed in Table C.2. 

Immediately filter the medium through a 0.45 

pm pore diameter membrane at a vacuum of not 

more than 380 mm (15 in.) mercury, or at a 

pressure of not more than one-half atmosphere 

(8 psi). Wash the filter with 500 mL deionized 

water before use. 

If the filtration is carried out with sterile apparatus, filtered medium can be used immediately, 

and no further sterilization steps are required 

before the inoculation of the medium. The medium can also be sterilized by autoclaving after 

it is placed in the culture vessels. Unused sterile 

medium should not be stored more than one 

week before use, because there may be substantial loss of water by evaporation. 

121 
C.2.2 Algal Cultures: Two types of algal cultures are 

maintained: (1) stock cultures and (2) “food” cultures. 

C.2.2.1 Establishing and Maintaining Stock Cultures of 

Algae: 

1. Upon receipt of the “starter” culture of S. 

capricornutum (usually about IO mL), a stock 

culture is started by aseptically transferring 1 

mL to each of several 250-mL culture flasks 

containing 100 mL algal culture medium (prepared as described above). The remainder of 

the starter culture can be held in reserve for up 

to six months in a refrigerator (in the dark) at 

4°C. 

2. The stock cultures are used as a source of 

algae to initiate “food” cultures. The volume of 

stock culture maintained at any one time will 

depend on the amount of algal food required for 

culture. Stock culture volume may be rapidly 

“scaled up” to several liters using 4-L serum 

bottles or similar vessels containing 3 L of growth 

medium. 

3. Culture temperature is not critical. Stock cultures may be maintained at 25°C in environmental chambers with cultures of other organisms if the illumination is adequate (continuous 

“cool-white” fluorescent lighting of about 

4300 lux). 

4. Cultures are mixed twice daily by hand. 

5. Stock cultures can be held in the refrigerator 

until used to start “food” cultures, or can be 

transferred to new medium weekly. One to 3 mL 

of 7-d old algal stock culture, containing about 

1.5 X 106 cells/ml are transferred to each 100 mL 

of fresh culture medium. The inoculum should 

provide an initial cell density of about 10,000 to 

30,000 cells/ml in the new stock cultures. Aseptic techniques should be used in maintaining the 

stock algal cultures, and care should be exercised to avoid contamination by other microorganisms. 

6. Stock cultures should be examined microscopically weekly at transfer for microbial contamination. Reserve quantities of culture organisms 

can be maintained for 6 to 12 months if stored in 

the dark at 4°C. It is advisable to prepare new 

stock cultures from “starter” cultures obtained 

from established outside sources of organisms 

every four to six months. 

C-2.2.2 Establishing and Maintaining ‘S. capricornutum 

food’ Cultures: 

1. ‘S. capricornutum food” cultures are started 7 d 

before use. About 20 mL of 7-d-old algal stock 

culture (described in the previous paragraph), 

containing 1.5 X 1 O6 cells/ml are added to each 

liter of fresh algal culture medium (e.g., 3 L of 

medium in a 4-L bottle or 18 L in a 20-L bottle). 

The inoculum should provide an initial cell density of about 30,000 cells/ml. Aseptic techniques 

should be used in preparing and maintaining the 

cultures, and care should be exercised to avoid 

contamination by other microorganisms. However, sterility of food cultures is not as critical as 

in stock cultures because the food cultures are 

used in 7 to 10 d. A one-month supply of algal 

food can be grown at one time and stored in the 

refrigerator. 

2. Food cultures may be maintained at 25°C in 

environmental chambers with the algal stock 

cultures or cultures of other organisms if the 

illumination is adequate (continuous “cool-white” 

fluorescent lighting of about 4300 lux). 

3. Cultures are mixed continuously on a magnetic 

stir plate (with a medium size stir bar), in a 

moderately aerated separatory funnel, or are 

manually mixed twice daily. If the cultures are 

placed on a magnetic stir plate, heat generated 

by the stirrer might elevate the culture temperature several degrees. Caution should be taken 

to prevent the culture temperature from rising 

more than 2 to 3°C. 

C.2.2.3 Preparing Algal Concentrate of S. capricornutum 

for Use as Food: 

1. 

2. 

3. 

4. 

5. 

An algal concentrate of S. capricornutum containing 3.0 to 3.5 X 10’ cells/ml is prepared 

from food cultures by centrifuging the algae with 

a plankton or bucket-type centrifuge, or by allowing the cultures to settle in a refrigerator for 

at least three weeks and siphoning off the supernatant. 

The cell density (cells/mL) in the concentrate is 

measured with an electronic particle counter, 

microscope and hemocytometer, fluorometer, 

or spectrophotometer and used to determine 

the dilution (or further concentration) required to 

achieve a final cell count of 3.0 to 3.5 X lo7 

cells/ml. 

Assuming a cell density of about 1.5 X 1 O6 cells/ 

mL in the algal food cultures at 7 d, and 100% 

recovery in the concentration process, a 3-L 

culture at 7 to 10 d will provide 4.5 X 1 Og algal 

cells. 

Algal concentrate can be stored in the refrigerator for one month. 

Cultures of Hyalella azteca are fed 10 mUL on 

Monday and 5 mL/t on Wednesday and Friday 

(ERL- Duluth, 1993). 

122 
C.2.2.4 Establishing and Maintaining Stock Cultures of 

Diatoms (Navicula sp. or Synedra sp.): 

1. Upon receipt of the “diatom starter’culture (usually about 10 mL), a stock culture is started by 

aseptically transferring 1 mL to each of several 

250-mL culture flasks containing 100 mL algal 

culture medium (prepared as described above). 

The remainder of the starter culture can be held 

in reserve for up to six months in a refrigerator 

(in the dark) at 4°C. 

2. The stock cultures are used as a source of 

diatoms to initiate “diatom food” cultures. The 

volume of stock culture maintained at any one 

time will depend on the amount of food required 

for culture. Stock culture volume may be rapidly 

“scaled up” to several liters using 4-L serum 

bottles or similar vessels containing 3 L of growth 

medium. 

3. Culture temperature is not critical. Stock cultures may be maintained at 23°C in environmental chambers with cultures of other organisms if the illumination is adequate (continuous 

“cool-white” fluorescent lighting of about 

1075 lux). 

4. Cultures are mixed twice daily by hand. 

5. Stock cultures can be held in the refrigerator 

until used to start “diatom food” cultures or can 

be transferred to new medium weekly. One to 3 

mL of 7-d old algal stock culture, containing 

about 1.5 X 1 OS cells/mL are transferred to each 

100 mL of fresh culture medium. The inoculum 

should provide an initial cell density of about 

10,000 to 30,000 cells/ml in the new stock cultures. Aseptic techniques should be used in 

maintaining the stock algal cultures, and care 

should be exercised to avoid contamination by 

other microorganisms. 

6. Stock cultures should be examined microscopically weekly at transfer for microbial contamination. Reserve quantities of culture organisms 

can be maintained for 6 to 12 months if stored in 

the dark at 4°C. It is advisable to prepare new 

stock cultures from “starter” cultures obtained 

from established outside sources of organisms 

every four to six months. 

C-2.2.5 Establishing and Maintaining “Diatom food” 

Cultures: 

1. “Diatom food” cultures are started about 10 d 

before use. About 20 mL of 7-d-old algal stock 

culture (described in the previous paragraph) 

are added to each liter of fresh culture medium 

(e.g., 3 L of medium in a 4-L bottle). Aseptic 

techniques should be used in preparing and 

maintaining the cultures, and care should be 

exercised to avoid contamination by other microorganisms. However, sterility of food cultures is not as critical as in stock cultures because the food cultures are used in 7 to 10 d. A 

one-month supply of diatom food can be grown 

at one time and stored in the refrigerator. 

2. Food cultures may be maintained at 23°C in 

environmental chambers, but not with the algae 

stock cultures. The illumination is continuous 

“cool-white” fluorescent lighting of about 

1075 fux). Higher temperatures can be problematic for diatom cultures. 

3. Cultures are mixed continuously on a magnetic 

stir plate (with a medium size stir bar) in a 

moderately aerated separatory funnel, or are 

manually mixed twice daily. Cultures become 

very brown before harvesting. If the cultures are 

placed on a magnetic stir plate, heat generated 

by the stirrer might elevate the culture temperature several degrees. Caution should be taken 

to prevent the culture temperature from rising 

more than 2 to 3°C. 

C.2.2.6 Preparing Concentrate of Diatoms for Use as 

Food: 

1. A diatom concentrate containing 1 X 1 Og cells/ 

mL is prepared from food cultures by centrifuging the algae with a plankton or bucket-type 

centrifuge, or by allowing the cultures to settle in 

a refrigerator for at least three weeks and siphoning off the supernatant. 

2. The cell density (cells/mL) in the concentrate is 

measured with an electronic particle counter, 

microscope and hemocytometer, fluorometer, 

or spectrophotometer and used to determine 

the dilution (or further concentration) required to 

achieve a final cell count of 1 X 1 Og cells/ml. 

3. Algal concentrate can be stored in the refrigerator for one month. 

4. Cultures of Hyalelia azteca are fed 10 mL/L on 

Monday and 5 mL/L on Wednesday and Friday 

(ERL- Duluth, 1993). 

C.2.2.7 Cell counts: 

1. Several types of automatic electronic and optical particle counters are available to rapidly 

count cell number (cells/mL) and mean cell 

volume (MCV; f.rm3/cell). The Coulter Counter is 

widely used and is discussed in detail in USEPA 

(1978). When the Coulter Counter is used, an 

aliquot (usually 1 mL) of the test culture is 

diluted 10X to 20X with a 1% sodium chloride 

electrolyte solution, such as Coulter ISOTONB, 

to facilitate counting. The resulting dilution is 

counted using an aperture tube with a 100~pm 

123 
diameter aperture. Each cell (particle) passing 

through the aperture causes a voltage drop 

proportional to its volume. Depending on the 

model, the instrument stores the information on 

per replicate are counted to obtain ?lO% precision at the 95% confidence level. This method 

has the advantage of allowing for the direct 

examination of the condition of the cells. 

the number of particles and the volume of each, 

and calculates the mean cell volume. The fol- C.3 Tetrafin@ food for culturing and testing C. tentans. 

lowing procedure is used: Food should be stored at 4°C and used within two 

weeks from preparation or can be frozen until use. 

A. 

8. 

C. 

D. 

Mix the algal culture in the flask thoroughly 

by swirling the contents of the flask about 

six times in a clockwise direction, and then 

six times in the reverse direction; repeat the 

two-step process at least once. 

1. 

2. 

At the end of the mixing process, stop the 

motion of the liquid in the flask with a strong 

brief reverse mixing action, and quickly remove 1 mL of cell culture from the flask with 

a sterile pipet. 

Place the aliquot in a counting beaker, and 

add 9 mL (or 19 mL) of electrolyte solution 

(such as Coulter ISOTONQ. 

3. 

Determine the cell density (and MCV, if 

desired). 

2. Manual microscope counting methods for cell 

counts are determined using a Sedgwick-Rafter, 

Palmer-Maloney, hemocytometer, inverted microscope, or similar methods. For details on 

microscope counting methods, see APHA (1992) 

and USEPA (1973). Whenever feasible, 400 cells 

4. 

Blend the Tetrafin@ food in deionized water for 

1 to 3 min or until very finely ground. 

Filter slurry through an #l 10 Nitex screen to 

remove large particles. Place aliquot of food in 

lOO- to 500-mL screw-top plastic bottles. It is 

desirable to determine dry weight of solids in 

each batch of food before use. Food should be 

held for no longer than two weeks at 4°C. Food 

can be frozen before use, but it is desirable to 

use fresh food. 

Tetrafiti food is added to each culture chamber to provide about 0.04 mg dry solids/mL of 

culture water. A stock suspension of the solids 

is prepared in culture water such that a total 

volume of 5.0 mL of food suspension is added 

daily to each culture chamber. For example, if a 

culture chamber volume is 8 L, 300 mg of food 

would be added daily by adding 5 mL of a 56 g/ 

L stock suspension (USEPA, 1993). 

In a sediment test, Tetrafim food (4 g/L) is 

added at 1.5 mL daily to each test chamber. 

124 
Appendix D 

Sample Data Sheets 

125 
Culture 

Aquarium 

A 

B 

C 

D 

E 

F 

Date of Egg 

Mass 

Deposition 

Date 4th 

lnstar 

Larvae 

Were 

Weighed 

Age of 

Weighed 

4th Instar 

Larvae 

Mean Dry 

Weight of 

4th lnstar 

Larvae 

(n = 10) 

Figure D.1 Data sheet for the evaluation of a Chironomus tentans culture. 

126 

Date of 

Observed 

First 

Emergent 

Adult 

Total 

Number of 

Egg 

Masses 

Produced 

General 

Comments 

Initials of 

Culturist 
Brood Stock Source 

Test Type (circle one)‘: SU SM RU RM FU FM 

No. of Animals Tested Per Replicate 

No. of Replicates 

Method of LWJ Estimate 

Reference Toxicant (CuSO, or KCI) 

Reference Toxicant Supplier and Lot No. 

Reference Toxicant Purity 

Test Initiation Date 

Toxicologist 

Exposure Duration (t-l) 

0 

24 

46 

R 

96 

COfltr01 Control Exp. 1 

A B A B 

Number of Mortalltles 

Exp. 2 Exp. 3 

A B A 0 

Current Test 96 h LC50 = 

Number of Reference Toxicant Test Used 

to Determine Cumulative Mean 96 h LC50 

Mean 96 h LC50 for All Tests to Date 

Acceptability of Current Test2 Yes NoExp. 4 Exp. 5 

A 0 A 0 

1 SU = Static unmeasured 

SM = Static measured 

RU = Renewal unmeasured 

RM = Renewal measured 

FU = Flow-through unmeasured 

FM = Flow-through measured 

2 Based on two standard deviations around the cumulative mean 96 h LC50 

Flgure 0.2 Data sheet for performing reference toxicant tests. 

127 
S+rrer! Sample Source 

Da:e i:‘ Test Irlt,at:on 

T,:x cc! ;q,st Con3uctirq Test 

-55: c3; 

I_ 

‘es: Dissolved Specific Total 

R?yllcafe Temperature Cxygen Hardness Alkalmlty Conductance Ammonia 

Saw ed i-c) (m@Li PH (mwU (mwU (umhos!cm) img/L) 

Figure D.3 Data sheet for temperature and overlying water chemistry measurements. 

128 
Daily Checklist for Sediment Tests 

Study Code 

Study Name 

Bulldmg Diluter Waterbath Target temperature -“C 

Dissolved Oxygen 

Minimum Acceptable Concentration 

(40% of Saturation at Target Temp) 

Study Director 

Lead Technician Month 

Acceptable Range “C to -“C 

= -mW 

Day of Month 1 1 1 2 1 3 1 4 I 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 a0 21 22 23 24 25 1 26 1 27 

Day of Study 

Diluter 

Operation 

Number of 

Cycles 

Tune of Day 

Temperature 

Aw Pressure 

Aeration 

Brush 

Screens 

____--. 

Clean 

Needles 

Feedlng 

Total Water 

Oualty 

Partial Water 

Comments 

Figure D.4 Data sheet for daily checklist for sediment tests. 

129 
Study Code 

Study NameDate 

Water Quality Data Sheet 

Test Day Study Dtrector lnvestlgator 

Volume of 

Sample = 

Sample Code 

Meter # 

Initials 

Comments 

DlSSolved ConductMy 

Oxygen Temp. s~~$y (umhos or pH 

(mgiL) “c US) 

Alkallnlty 

(pm 

CaCO,) 

m 

I ml of Other 

IsA ~~ 

ml - -ml -ml -ml -ml -ml -ml -ml -ml 

I -1 -1 

~ I ~ I -1 

-I -.I -1 

___ 

I I - -1 

-1-I -I 

___ 

I __ I -- 

I -I -I 

I -I -I 

I I - - ---I 

(ml Tltrant x mult. (ml Tltrant x mult. 3 

factor =) factor =) s I- 

..~ I __ 

I x I I - : ~~~_. I px-= ___ I _..m_-I -.- 1 -1 

I I-x- z ___ __ -1 pxp=-p I I -1 -1 ---I .- __I 

I I x-= -- -x-=-- I I --I -I -1 --I 

IpIpx - zz _ I -X-= I I I I -1 --I 

1-J-x ; - -1 -x- = _- I I -1 -I -I -I 

I I -x - =-I -x-= _- I I -1 -I -I -I 

1 I-x- = ___ __ -1 -x-= J-1 1 --I --I -I 

I-1 -x - --I -x-= I I ---I -I --I -1 

1-I-x - = -1 -x-=-- I I ---I ---I -I -I 

l-I--.2 - = -I -x- = _- I I --I --I -I -I 

-1-1 -x - = -I -x- _- I I -I ----I ---I -I 

I-1-x - = -1 -x- = 1-l -I -I -I -I 

----I -I --I I I -- ---I _I ---I ---I 

- - _---___- 

Approved by Date 

Figure D.5 Data sheet for water quality parameters. 

130 
Chemistries 

Test Type 

Organism 

Test 

Dates 

Sample Info Water Type 

Experimenter 

Test 

System 

PH 

DO (mg/L) 

Temp (“C) 

Hard/Alk 

PH 

DO VW-1 

Temp (“C) 

Hard/Alk 

PH 

DO (mgiL) 

Temp !*C) 

HardlAlk 

?H 

DO (mgiL) 

Temp (“C) 

Hard:Alk 

Figure 0.6 Chemistry data sheet. 

131 
Study Director 

Study Code 

Study Name 

Day 

Daily Comment Sheet 

Data - - Initials 

Day Data - - Initials 

Day Date - - Jnitials 

Day Data - - lnitkds 

Day Date - - Initials 

Figure D.7 Dally comment data sheet. 

132 
Weight Data Form 

est Dates 

est Material 

ocation 

nalyst 

Sample Replicate 

Species 

Weighing Date Food 

Oven Temp (“C) Age Organisms 

Drying Time (h) .- Initial No/Rep 

Wt of (mg) wt. of Owl 

oven dried pan + f&tan dry OW 

pan organism Total (mg) Survival (#) wt./- Mean/Sample 

Figure D.8 Weight data sheet. 

133 
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