EPA/600/6-91/007
September, 1991
Sediment Toxicity Identification Evaluation:
Phase I (Characterization), Phase n (Identification)
and Phase ffl (Confirmation) Modifications of Effluent Procedures.
by
Gerald T. Ankley
U.S. Environmental Protection Agency
Environmental Research Laboratory-Duluth
6201 Congdon Blvd.
Duluth, MN 55804
Mary K. Schubauer-Berigan
AScI Corporation
6201 Congdon Blvd.
Duluth, MN 55804
Joseph R. Dierkes
and
Marta T. Lukasewycz
AScI Corporation
6201 Congdon Blvd.
Duluth, MN 55804
National Effluent Toxicity
Assessment Center
Technical Report 08-91
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EPA/600/6-91/007
September, 1991
Sediment Toxicity Identification Evaluation:
Phase I (Characterization), Phase n (Identification)
and Phase HI (Confirmation) Modifications of Effluent Procedures.
by
Gerald T. Ankley
U.S. Environmental Protection Agency
Environmental Research Laboratory-Duluth
6201 Congdon Blvd.
Duluth, MN 55804
Mary K. Schubauer-Berigan
AScI Corporation
6201 Congdon Blvd.
Duluth, MN 55804
Joseph R. Dierkes
and
Marta T. Lukasewycz
AScI Corporation
6201 Congdon Blvd.
Duluth, MN 55804
National Effluent Toxicity
Assessment Center
Technical Report 08-91
-------
NOTICE
Mention of trade names or commercial products does not constitute endorsement or recom-
mendation for use.
u
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Acknowledgements
The work described herein has been performed, since 1988 at the National Effluent Toxicity
Assessment Center (NETAQ. Current or former NETAC scientists Teresa Norberg-King,
Don Mount, Joe Amato, Larry Burkhard, Liz Durban, Steve Baker, Lara Anderson, Jim
Jenson, Greg Peterson, Jo Thompson, Doug Jensen, Shaneen Murphy, Linda Eisenschank,
Nola Englehom and An Fenstad all contributed to the research presented here. Jeff Denny,
Scott Coilyard, and Kurt Mead provided fathead minnows (Pimephales promelas, Hyalella
azteca, and Chironomus tentans, and Gary Phipps provided Lumbriculus variegatus for this
work. Debra Williams assisted in coordinating the document Finally, of great importance to
this work was the financial and programmatic support given by W.R. Brandes, Office of
Water, Permits Division, and Nelson Thomas, Senior Advisor for National Programs.
111
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CONTENTS
Page
Notice u
Acknowledgements L11
Contents -IV
Figures vii
Tables
I. Introduction I-1
TIE Overview H-l
n.l Phase I H-l
0.2 Phase 0 H-7
n.3 Phase ffl H-10
EEL Special Consideration for Sediment TIE EH-1
III.1 Aqueous Fraction Selection ffl-1
EQ.2 Aqueous Fraction Preparation IH-4
IEI.2.1 Saginaw River Sediment Pore Water Characterization ffl-6
HL2.2 Keweenaw Waterway Sediment Pore Water Characterization HI-9
HL2.3 Recommended Pore Water Preparation Method EH-12
En.3 Use of Benthic Species for Aqueous Testing HI-13
HL3.1 Selection of TIE Species 01-16
ffl.4 Test Volume Consideration ffl-20
in.5 Common Sediment Contaminants: Ammonia, Metals,
and Hydrogen Sulfide ffl-22
EH.5.1 The Graduated pH Test ffl-23
HL5.1.1 Methods of pH Control ffl-26
HL5.2 Alternative Species Testing ffl-31
HL5.3 Toxicant Dilution Testing ffl-32
EEL5.4 Recovering Volatile and Filterable Contaminants ffl-32
HL5.4.1 Volatile Toxicant Transfer Experiment ffl-33
EEL5.4.2 Recovering Filterable Toxicity ffl-34
HL6 Clt Fractionation Consideration ffl-35
IV
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CONTENTS (continued)
Page
IV. Sample Collection, Preparation and Initial Toxicity Tests IV-1
FV.l Shipping jv-1
FV.2 Arrival and Storage rV-1
FV.3 Test Fraction Preparation JV-2
IV.3.1 Pore Water Preparation TV-2
IV.3.2 Elutriate Preparation FV-4
FV.4 Toxicity Tests IV-5
V. Methods for Phase I Sediment TIE V-1
V.I Initial Test V-l
V.2 Baseline Test V-l
V.3 TIE Toxicity Tests V-2
V.4 pH Adjustments V-3
V.5 Filtration V-6
V.6 Aeration V-7
V.7 Qg Solid Phase Extration V-7
V.8 Readjustment of Samples to pH i/Toxicity Testing V-l 1
V.9 EDTA Chelation Test V-ll
V.10 Sodium Thiosulfate Test V-l2
V. 11 Graduated pH Test V-14
V.I 1.1 Graduated pH Test: Closed-cup Method V-l5
V.I 1.2 Graduated pH Test: COj Method V-17
V.I 1.3 Graduated pH Test: Buffer Method V-l9
VI. Methods for Phase 0 Sediment TIE VI-1
VI. 1 Filter-Removable Toxicants: Metals and Nonpolar
Organic Compounds VI-2
VI. 1.1 Nonpolar Organic Compounds: General Overview VI-2
VI. 1.1.1 Nonpolar Organic Compounds: Filter Extraction VI-5
VI.1.1.2 Nonpolar Organic Compounds: High-Speed Centrifugation VI-7
VI. 1.1.3 CltSPE Fractionation VI-7
VI. 1.2 Metals: General Overview VI-13
VI. 1.2.1 Filter Extraction VI-13
VI.2 Use of Multiple Manipulation in Phase n VI-14
VI.3 Volatile Toxicants: Hydrogen Sulfide VI-15
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CONTENTS (continued)
Page
VIL Methods for Phase ED Sediment TIE VH-1
VH.l Correlation VII-1
VH.2 Species Sensitivity VH-5
VH.3 Mass Balance VH-7
vn.4 Deletion Approach VH-9
VH.5 Symptoms VH-10
VH.6 Spiking VH-10
VH.7 Matrix Changes vn-12
VH.8 Summary vn-13
VIIL Literature Cited VIH-l
VI
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FIGURES
Page
II-1 Overview of Phase I pore water characterization tests 0-2
m-1 Relative sensitivities of Pimephales promelas,
Ceriodaphnia dubia, Hyalella azteca, and
Lwnbriculus variegams to sediment pore water
and sediment elutriate. ffl-17
VH-1 Correlation of concentrations of ammonia in sediment
pore waters from the lower Fox River/Green Bay watershed
with toxicity of the samples to fathead minnows and C. dubia. Vfl-4
vu
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TABLES
Page
n-1 Summary of analytical methods currently used or proposed
for Phase n TIEs. n-9
HI-1 Summary of toxicity data for pore water versus bulk sediment,
or elutriate versus bulk sediment to Pimephales promelas,
Hyalella azteca and Lumbricuius variegatus. HI-3
LH-2 Results of pore water characterization studies of Saginaw
River (MD sediments. ffl-7
HI-3 Results of pore water characterization studies of Keweenaw
Waterway (MI) sediments. HI-10
ffl-4 Trends in metal and ammonia toxicity with respect to test pH. ffl-18
ffl-5 Sensitivities of C. dubia, fathead minnow, H. azteca, and
L. variegatus to the pH-control buffers, Mes, Mops, and Popso. ffi-29
IH-6 Sensitivity of C. dubia to certain metals (tested using
different pH-adjustment/control techniques), and ability
of EDTA to chelate metal toxicity in the presence and
absence of pH-control buffers. ffl-30
V-l Species sensitivity to Phase I additives. V-5
VI-1 Composition of 11 recommended solvents for eluting the
Cu column in Phase 0 sediment TIE. VI-9
Vn-1 Comparison of the sensitivities of C. dubia, fathead
minnow, and Photobacterium phosphoreum to pore water
to Green Bay/Fox River sediment pore water. VTI-6
VTI-2 Comparison of the sensitivies of C. dubia, fathead
minnows and H, azteca to sediment pore water from five
sites along Turkey Creek, Joplin, MO. VTI-8
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I. Introduction
The extent of sediment contamination in the United States has been amply documented, and it
is apparent that in order to comply with the Qean Water Act the issue of contaminated
sediments must be addressed by the U.S. Environmental Protection Agency. Studies in
conjunction with regulatory/remedial activities with contaminated sediments at a great number
of freshwater and marine sites have demonstrated that the sediments are acutely and/or
chronically toxic to a variety of test species. Although the presence of toxicity clearly is a
matter of concern with regard to existing or potential impacts of sediment-associated
contaminants on benthic, epibenthic or pelagic organisms, toxicity alone does not provide a
useful or logical basis for regulatory activities focused upon identifying remedial options such
as point-source control. It clearly would be desirable to be able to identify those compounds
responsible for sediment toxicity.
Attempts to use chemical screening (e.g., priority pollutant analyses) and correlation tech-
niques to identify specific contaminants responsible for sediment toxicity generally have not
been successful for a number of reasons. First, there are thousands of contaminants present at
detectable concentrations in most contaminated sediments; therefore, it is impossible to
ascertain whether chemicals responsible for toxicity are even measured. Second, even if all
possible contaminants of concern could be measured, there are virtually no reliable techniques
for assessing the biological availability of each component of the complex mixture of
compounds in the sediments. Finally, even if bioavailability issues could be resolved, it is
1-1
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difficult to predict the additive, antagonistic and/or synergistic interactions that may occur
among the contaminants. These same types of problems were encountered in the NPDES
permitting program focused upon utilizing toxicity to aquatic organisms in effluent permit
limits; i.e., it was difficult to use chemical-specific approaches to define those compounds
responsible for toxicity of complex effluents (Burkhard and Ankley 1989). In response to
this, researchers at the Environmental Research Laboratory in Duluth developed a set of
toxicity-based guidelines for identifying toxic compounds in complex effluents (U.S. EPA
1988; 1989a; 1989b; 1991a). These toxicity identification evaluation (TIE) procedures utilize
toxicity-based fractionation schemes to characterize (Phase I), identify (Phase IT) and confirm
(Phase HI) compounds responsible for sample toxicity. Initial studies focused on the use of
these TIE procedures for identifying toxicants in effluents (e.g., Amato et aL 1992; Ankley
and Burkhard 1992); further work, however, demonstrated that TIE also could be used to
successfully identify acutely toxic compounds in ambient waters (Norberg-King et al. 1991),
hazardous waste leachates (Kuehl et al. 1990) and aqueous fractions of sediments (Ankley et
al. 1990a; 1991a; Schubauer-Berigan and Ankley 1991).
The identification of compounds responsible for toxicity of contaminated sediments has a
broad application in a number of EPA programs. For example, the ability to link sediment
toxicity to a specific discharger could be used to develop discharge permit limits protective of
aquatic species associated with sediments. Along these lines, the results of sediment TEE
procedures may be useful in ascribing responsibility to parties involved in ongoing remedia-
tion activities at contaminated locations, such as Superfund sites. The identification of
1-2
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specific compounds responsible for toxicity also could be useful in designing effective
remediation. For example, sediment toxicity due to ammonia would be dealt with in quite a
different manner than toxicity caused by metals or pesticides. It also may be possible to use
sediment TIE procedures in permitting programs for dredged materials in order to identify
environmentally protective options for disposal (Ankley et al. 1991b). Finally, the identifica-
tion of specific problem contaminants in sediments could prove to be very useful to EPA
programs involved in the development of water or sediment quality criteria, and the registra-
tion of compounds such as pesticides.
The following document was developed to provide guidance for performing sediment TIE
analyses. This guidance does not include recommendations for the implementation of
sediment TIE in a regulatory context. The document is divided into eight sections: I.
Introduction; EL TEE Overview; EH. Special Considerations for Sediment TIE; IV, Sample
Collection, Preparation and Initial Toxicity Tests, V. Methods for Phase I Sediment TIE; VI.
Methods for Phase 0 Sediment TIE; VH. Methods for Phase EH Sediment TIE, and VUL
Literature Cited. Section n consists of a brief overview of existing TIE procedures. Section
in presents the conceptual and technical basis for several aspects of sediment TEE procedures,
in particular those that differ from effluent TEE methods. Sections FV, V, VI and VH present
specific procedural details for collecting and preparing samples and for performing sediment
TEE. Emphasis in these sections is given to those manipulations and interpretations in Phases
I, El, and EQ that differ from existing guidance for effluents. Thus, it is imperative that the
1-3
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E
snecificallv U.S. EPA 1988: 1989* 1989b: and 199 la.
1-4
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n. TIE Overview
HI Phase I
Phase I characterizes the physical/chemical properties of sample toxicants through the use of
manipulations designed to alter or render biologically unavailable generic classes of com-
pounds with similar properties (U.S. EPA 1988; 199la). Toxicity tests, performed in
conjunction with the manipulations, provide information on the nature of the toxicant(s).
Successful completion of Phase I occurs when both the nature of the components causing
toxicity, as well as their consistency among different samples, can be established. After
Phase L, the toxicant(s) can be tentatively categorized as having chemical characteristics of
cationic metals, non-polar organics, volatiles, oxidants, substances whose toxicity is pH
dependent and/or substances whose toxicity is not influenced by Phase I methods, e.g.,
possibly a polar organic and/or anionic inorganic.
Fig. El-1 shows an overview of the sample manipulations employed in Phase I. Not shown in
Fig. II-1, but performed initially on all samples are routine water chemistry measurements
including pH, hardness, conductivity and dissolved oxygen. These routine measurements are
needed for designing sample manipulations and interpreting test data. The manipulations
shown are usually sufficient to characterize toxicity caused by a single chemical and some
combinations of toxicants (e.g., nonpolar organics, ammonia). When other toxicants are
n-i
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Figure 11-1. Overview of Phase 1 pure water characterization tests (note: pH i represents initial pH).
*
Toxic Effluent Sample
Initial Toxicity Test
(Day 1)
A
J
Baseline Toxicity
Test (Day 2)
Aeration Tests
(Day 2)
t
Filtration Tests
(Day 2)
I
t
*
I
EDTA
Chelation
Test (Day 2)
pH Adjustment
Tests (Day 2)
f
I
Base
C1H Solid Phase
Extraction Tests
(Day 2)
*
*
Oxidant
Reduction
Test (Day 2)
1
Graduated pH
Tests (Day 2)
f
pH7
pH8
II 2
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present simultaneously, various sequential combinations of the Phase I manipulations will be
required for toxicant characterization.
Many of the manipulations in Phase I require that samples be pH adjusted. The adjustment
of pH is a powerful tool for detecting cationic and anionic toxicants because their behavior is
strongly influenced by pH. By changing the pH of a sample, the ratio of ionized to un-
ionized species in solution for an anionic chemical is changed significantly. The ionized and
un-ionized species have different physical/chemical properties as well as toxicities. In Phase
I, two types of pH manipulations are used to examine different questions. First, "Is the
toxiciry of the sample different at various pHs?", and second, "Does changing the pH,
performing a sample manipulation, and then readjusting it to ambient pH affect toxiciry?".
The graduated pH test examines the first question, and the pH adjustment and subsequent
aeration, filtration and solid phase extraction (SPE) manipulations examine the second
question.
In the graduated pH test, the pH of a sample is adjusted within a physiologically tolerable
range, e.g., pHs 6.5, 7.5, and 8.5, before toxiciry testing. For some chemicals (e.g., ammonia,
ionizable organic pesticides), the un-ionized form of a compound is able to cross biological
membranes more readily than the ionized form and thus, is more toxic. In other instances
(e.g., for metals) the more soluble form of a chemical, a property also affected by pH, tends
to be more toxic. Thus although the graduated pH test originally was designed primarily for
ammonia, a relatively common toxicant in sediments and effluents whose toxiciry can be
n-3
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extremely pH-dependent, the test also is useful for implicating the presence of toxic concen-
trations of some pesticides and certain canonic metals (U.S. EPA 1985; Campbell and Stokes
1985; Doe et al. 1988; Schubauer-Berigan et aL 1992). Differences in test pH also influence
the toxicity of hydrogen sulfide, another relatively common sediment contaminant (Broderius
et al. 1977).
Aeration tests are designed to determine whether toxicity is attributable to volatile, oxidizable
or sublatable compounds. Samples at pH i (the pH of the sample under laboratory condi-
tions), pH 3, and pH 11 are sparged with air for one hour, readjusted to pH i, and tested for
toxicity. The different pHs affect the ionization state of polar toxicants, thus making them
more or less volatile, and also affect the redox potential of the system. If toxicity is reduced
by air sparging at any of the pHs, the presence of volatile or oxidizable compounds is
suggested. To distinguish the former from the latter situation, further experiments are
performed using nitrogen to sparge the sample(s), rather than air. If toxicity remains the
same as in the baseline toxicity test, oxidizable materials are implicated; if toxicity is again
reduced, volatile compounds are suspected. The pH at which toxicity is reduced is also
important If nitrogen sparging decreases toxicity at pH i, neutral volatiles are present;
whereas, if toxicity decreases at pH 11.0 or pH 3.0, basic and acidic volatiles, respectively,
are implicated. An additional mechanism through which toxicants can be removed from a
sample by aeration is sublation, which is movement of die compound through the aqueous
phase on the surface of the air bubbles, followed by deposition as a solid on the aeration
glassware at the air/water interface. If sublation were the mechanism through which sample
0-4
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toxicants were removed, it might be possible to recover this toxicity by rinsing the aeration
glassware (Ankley et aL 1990b). Compounds possessing both polar and nonpolar characteris-
tics, such as surfactants or resin acids, are particularly prone to sublation from aqueous
samples.
In the filtration test, samples at pH /, pH 3,0 and pH 11.0 are passed through 1 um glass fiber
filters, readjusted to pH i, and tested for toxicity. Filtration provides information concerning
the amount of toxicity associated with filterable components; however, filtration of the sample
also may remove toxicants through adsorption of compounds to the filter or substances on the
filter. Thus, of all of the Phase I manipulations, filtration is probably the least useful for
identifying specific classes of toxicants.
Reverse-phase solid phase chromatography is designed to determine die extent of sample
toxicity due to compounds that are relatively nonpolar at pH i, pH 3.0 or pH 9.0. This test,
in conjunction with associated Phase n analytical procedures, is an extremely powerful TIE
tool. In this procedure, filtered sample aliquots at pH i, pH 3.0 and pH 9.0 are passed
through separate small columns packed with an octadecyl (Cts) sorbent At pH i, the C,,
solid phase exchange (SPE) column will remove neutral nonpolar compounds, such as certain
pesticides (Junk and Richard 1988) and some metals (unpublished data). By shifting
ionization equilibria at the low and high pHs, the SPE column also can be used to extract
organic acids and bases (Wells and Michael 1987). During extraction, the resulting post-
column sample is collected and tested for toxicity in order to determine whether or not the
0-5
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manipulation removed toxicity and/or whether the capacity of the column was exceeded. If
sample toxicity is decreased, a nonpolar toxicant is suspected. Because the Q, SPE column
may remove metals and metal complexes, observation of toxicity recovery in a methanol
elution provides stronger evidence for a nonpolar organic toxicant than SPE toxicity removal
alone (U.S. EPA 1991a).
The presence of toxicity due to canonic metals is tested through additions of ethylenediamine-
tetraacetic acid (EDTA), a strong chelating agent which produces non-toxic complexes with
many metals. As with SPE chromatography, the specificity of the EDTA test for a class of
ubiquitous toxicants makes it a powerful TIE tool. Cations chelated by EDTA include certain
forms of aluminum, barium, cadmium, cobalt, copper, iron, lead, manganese, nickel, strontium
and zinc (Stumm and Morgan 1981). EDTA does not complex anionic forms of metals, and
only weakly chelates certain canonic metals (e.g., silver, chromium, thallium) (Stumm and
Morgan 1981). EDTA appears to preferentially bind the aforementioned transition metals
over calcium and magnesium (hardness ions), and studies at ERL-Duluth suggest the
equilibration time for heavy metal cheiation by EDTA is relatively brief for samples of
various hardnesses (J. Thompson, NETAC, personal communication).
The sodium thiosulfate addition test is designed to determine the presence of toxicity
associated with chemicals reduced or chelated by thiosulfate. Oxidants such as chlorine,
bromine, iodine and manganous ions can be neutralized by this treatment Sodium thiosulfate
also will chelate, and reduce the biological availability of, a number of canonic metals
0-6
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including copper, cadmium, mercury and silver (Hockett and Mount 1990). Sodium thiosul-
fate, like EDTA, has low toxicity to most aquatic organisms; therefore a relatively wide range
of concentrations can be tested.
H.2 Phase H
The information obtained in Phase I provides the analytical roadmarks for performing the
fractionation and identification tasks in Phase El. For example, if the addition of EDTA
reduced sample toxicity in Phase I then analytical techniques appropriate for metal analyses
(e.g., atomic adsorption spectroscopy) would be utilized in Phase II of the TIE. Similarly, if
Ctl SPE reduced sample toxicity, which was recoverable in a methanol elution of the column
in Phase I, then different concentration, separation and detection techniques suitable for
nonpolar organics would be utilized in Phase n (e.g., see Durhan et al. 1990; Burkhard et al.
1991). An important component of all of the Phase n procedures is the concurrent use of
toxicity tests with the test species of concern from Phase I, both to "track" toxicity through
various sample fractionations (e.g., in the case of nonpolar organic s), and just as importantly,
in single chemical exposures to help evaluate whether measured concentrations of suspect
toxicants in the unknown sample are sufficient to result in observed toxicity.
The types of analytical techniques that theoretically could be used in Phase n of the TIE are
quite variable and dependent not only upon the types and concentrations of toxic compounds
present in the test sample, but to a certain extent also upon the number of compounds that
appear to be contributing to sample toxicity. A full discussion of the types of Phase I results
0-7
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that could be encountered, and the specific Phase U analyses that may be used is beyond the
scope of this guidance document; however, changes/additions to the original Phase n
document (U.S. EPA 1989a) are being m^, and a revised version of the document is
anticipated by mid-1992 (E. Durban, U.S. EPA, ERL-D, personal communication).
Table 0-1 indicates several examples of Phase n analytical methods that have been or could
be used, depending upon the classes of toxicants indicated in Phase I. Most of the specific
rractionation/identification/quantiflcation methods indicated in Table £1-1 can be used either
on whole samples, or where appropriate, on some fraction of the sample (e.g., solvent
fractions from a Clt column or filter extract, sample fractions after being passed over a C,, or
cation exchange column, etc.). Some of the techniques listed, although theoretically feasible
(e.g., liquid chromatography/mass spectrometry for polar organics) have not been used
extensively in TIE studies at ERL-Duluth.
Upon successful completion of Phase n, one or more compounds will have been identified as
suspect toxicants, based both upon their presence in the test sample and their concentration
relative to the concentration expected to result in toxicity to the organism used in the toxicity
tests. Depending upon the results of the Phase ffi confirmation process (described below), it
may be necessary to revisit the Phase I and n portions of the lit.
n-8
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Table II-1. Summary of analytical methods currently used or proposed for Phase II TIEs.
Compound Class
Analytical Methods
Nonpolar Organics
Metals
Polar Organics
Ammonia, Hydrogen sulfide
High Pressure Liquid Chromatography (HPLC)
Gas Chromatography (GC)-Mass Spectroscopy (MS)
MS MS
Inductively-Coupled Plasma Emission Spectroscopy (ICAP)
Atomic Absorption Spectroscopy (AAS)
LCMS
Colohmethc Methods
Specific Ion Electrodes
119
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0.3 Phase HI
After Phase n identification procedures implicate suspect toxicants, Phase HI is initiated to
confirm that the suspects are indeed the true toxicants. Confirmation is perhaps the most
critical step of the TIE process because procedures used in Phases I and n may create
artifacts which could lead to erroneous conclusions about the toxicants. Furthermore, there is
a possibility that substances causing toxicity change from sample to sample. Phase HI
enables both situations to be addressed. The tools used in Phase EQ include correlation,
evaluation of relative species sensitivity, observation of symptoms, spiking and mass balance
techniques, as well as approaches that feature specific changes in water quality. In most
instances no single Phase ffl test is adequate to confirm suspects as the true toxicants;
therefore multiple confirmation procedures are necessary to develop a "weight of the
evidence" argument.
In the correlation approach, observed toxicity is regressed against expected toxicity due to
measured concentrations of the suspect toxicant(s). For the correlation approach to succeed,
sample variation must be wide enough to provide a range of values adequate for meaningful
analyses. In the case of effluents, this variation generally is achieved by collecting samples
over rime (e.gM Amato et al. 1992); however, for sediments among-sample variation in
concentrations of toxicants may have to be maximized by collecting samples from a number
of locations (e.g., Ankley et al. 1990a), The number and types of sediment samples needed to
conduct a meaningful correlation analysis is very site-specific; considerations in performing
this type of confirmation are discussed in greater detail in Section VTI below.
H-10
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By utilizing the toxicity correlation technique described by U.S. EPA (1989b), the analyst has
a statistical basis for reliably attributing sample toxicity to a specific compound (or com-
pounds). In order to use the correlation approach effectively when there are multiple suspect
toxicants, data must be generated concerning the additive, antagonistic and synergistic effects
of the toxicants in ratios similar to those found in the samples. These data also are needed
for the spiking and mass balance techniques described below.
The relative sensitivity of different test species can be used to evaluate suspect toxicants. If
there are two (or more) species that exhibit markedly different sensitivities to a suspect
toxicant in pure chemical toxicity tests, and the same patterns in sensitivity are seen with the
test sample, this provides evidence for the validity of the suspect being the true toxicant
Another Phase in procedure is observation of symptoms in test animals. Although this
approach does not necessarily provide support for a given suspect, it can be used to provide
evidence against a suspect toxicant If the symptoms observed in a pure chemical toxicity
test with a suspect toxicant are much different from those observed with the test sample
(which contains similar concentrations of the suspect toxicant), this is strong evidence for a
miside ntification.
Confirmatory evidence can be obtained by spiking samples with the suspect toxicants. While
not conclusive, if toxicity is increased by the same proportion that the concentration of the
suspect toxicant is increased in the sample, this suggests that the suspect is correct To get a
n-n
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proportional increase in toxicity from the addition of a suspect toxicant when in fact it is not
the true toxicant, both the true and suspect toxicants would have to have very similar toxicity
levels and would presumably have to be additive.
Confirmatory evidence also can be obtained by spiking samples from which the suspect
toxicant has been removed (e.g., via SPE or cation exchange resins, aeration, filtration, etc.)
to the original ambient concentrations of the toxicant Theoretically, the spiked sample
should have the same toxicity as the unaltered sample; this manipulation is particularly
attractive in that, as opposed to using laboratory control/dilution water for spiking and toxicity
comparisons, data may be derived for the suspect toxicants in a matrix that is similar to that
of the original test sample.
Mass balance calculations can be used as confirmation steps when toxicity can be (at least
partially) removed from the sample, and subsequently recovered. This approach can be useful
in instances when SPE chromatography or filtration removes toxicity. The solvent fractions
eluted from the SPE column are evaluated individually for toxicity; these toxicities are
summed and then are compared to the total amount of toxicity lost from the sample.
The alteration of water quality characteristics (generally pH) in a manner designed to affect
the toxicity of specific compounds also can provide very powerful confirmatory evidence.
This approach has been especially useful for sediment and effluent samples in which
ammonia and/or metals were sample toxicants.
n-i2
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Note that, of the Phase I, Phase II and Phase 03 portions of the TIE, we have had the least
actual experience with the latter, particularly in the case of sediments. Thus, the guidance
presented below for Phase m of the TIE is necessarily somewhat less well defined than
guidance for Phases I and H
H-13
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HL Special Considerations for Sediment TIE.
Because sediments are solid phase samples, TIE procedures designed for complex effluents
require certain modifications to be used for sediment evaluations. The appropriateness of
using an aqueous fraction to represent bulk sediment toxicity, the selection and preparation of
a particular aqueous fraction, and the use of benthic species in aqueous phase tests all must
be addressed when considering the use of TEE procedures with contaminated sediments.
Before beginning sediment TIE studies, consideration also must be given to appropriate
sampling of the contaminated area. A representative spatial sampling scheme, including tests
with a reference (uncontaminated) site, conducted over time is likely to give a more complete
assessment of sediment toxicity than single sites sampled only once. The representativeness
of sampling, however, is site-specific and, other than general recommendations, cannot be
addressed adequately in this document
HI.1 Aqueous Fraction Selection
Because TEE procedures were designed for use with aqueous samples, the extraction of some
aqueous fraction from the bulk sediment is required. Two aqueous fractions that have
commonly been used to assess sediment toxicity include interstitial (pore) water (extracted
directly from the sediments) and elutriates (the supernatant from a water/sediment mixture).
Pore water has been implicated for benthic organisms as an important route of exposure for
several nonpolar organic compounds (Eadie et al. 1982; Adams et al. 1985; Knezovich et al.
ffl-1
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1987; Connell et al. 1988; Swam et aL 1990), and metals (Swanz et al. 1985; DiToro et al.
199O, Ankley et aL 1992), and has been used in recent studies to evaluate in-place sediment
toxicity (Giesy et al. 1989; Swartz et al. 1989). Elutriates were developed initially to
simulate the dispersal of toxicants to the water column during suspension events such as
open-water disposal of dredged materials (U.S. Army Corps of Engineers/U.S. Environmental
Protection Agency 1977). Despite the limited intent of design, elutriates also have been used
to evaluate the toxicity of in-place sediments (Chapman and Fink 1984; Burton et al. 1989;
Long et al. 1990).
Recent work performed in conjunction with toxicity studies of 29 contaminated sediments
from throughout the U.S. showed that pore water (prepared by centrifugation) was a more
conservative predictor of sediment toxicity than elutriate (Ankley et al. 1991c; Schubauer-
Berigan and Ankley 1991). To ascertain this, Ankley et al. (199lc) compared the toxicity of
pore water, elutriate and bulk sediment to each of three species [Hyalella azteca, fathead
minnow (Pimephales promelas), and Lumbriculus varieganu] in 96 h acute exposures.
Elutriates were not particularly accurate in predicting bulk sediment toxicity, especially with
sensitive species such as H. azteca (Table EM). The percentage of "Type H" errors (i.e.,
failing to predict actual hulk sediment toxicity) was much greater for elutriates than for pore
water. For the more sensitive species, the percentage of 'Type I" errors (i.e., falsely
predicting bulk sediment toxicity) was generally low for both elutriates and pore water.
Based on these results, as well as the various studies cited above, pore water appears to be a
m-2
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Table
I1I-1. Summary of toxicity data for pore water versus bulk sediment, or elutriate versus bulk sediment to Pimephales promelas, Hyalella
azteca and Lumbriculus variegaius (from Ankley et al. 1991c).
Species
n
T/T1
Bulk Sediment/Pore Water
NT/NT
T/NT
NT/T
Concordance (%) False negatives (%)
P. promelas
//. azteca
L. variegaius
26
22
27
72
13
4
8
4
15
2
1
1
9
4
7
58
77
71
8
5
4
Species
Bulk Sediment/Elutriate
T/T1
NT/NT
T/NT
NT/T
Concordance (%) False negatives (%)
P. promelas
H. azteca
L. variegatus
27
21
26
5
7
1
16
6
19
4
7
5
2
I
1
78
62
77
15
33
19
Toxic (T) or Non-Toxic (NT)
Number of samples exhibiting indicated response.
1113
-------
more representative aqueous fraction than elutriate for the prediction of m situ bulk sediment
toxicity.
Overall, the appropriate aqueous fraction for use in sediment TIE is dependent on the specific
goal of the TIE. If the effect of resuspended sediments on water column chemistry or
toxicity is to be determined, then an elutriate is a more appropriate test fraction than pore
water. If, however, the purpose of the TIE is to identify compounds contributing to the
toxicity of in-place sediments, then pore water is probably the more relevant aqueous fraction
(Ankley et al. 199 Ic). Because the anticipated use of this document is primarily for the
detection of in-place toxicants, most experiments and procedures described herein were
developed using pore water as a test fraction.
in. 2 Aqueous Fraction Preparation
Regardless of the aqueous fraction used in the TIE, the extraction method will strongly
influence chemical composition and toxicity of the test sample. There are several commonly
used methods for extracting pore water from sediments. These include pressure extraction,
centrifugation at different speeds, dialysis, syringe extraction, and solvent displacement
These techniques may or may not require filtration as part of the extraction process. Each of
these, at one rime or another, has been recommended as superior for studying dissolved
nutrients, trace metals or organics in sediments: Benes and Steinnes (1974) and Carignan et
al. (1985) advocate in situ dialysis for metal analyses, Batley and Giles (1980) recommend
m-4
-------
solvent displacement for metal and organic analyses, Schults et al. (1991) suggest centriruga-
tion followed by filtration for studies of dissolved metals and nonpolar organic compounds.
Because TIE work requires large volumes of pore water (e.g., >1 L for Phase I), other
considerations being equal, methods that enable rapid preparation of relatively large sample
volumes are preferable to techniques that are labor-intensive, because of rime or sediment
volume restrictions.
Although the studies noted above and many others have compared the effects of pore water
extraction methods on nutrient or trace metal partitioning and speciarion, no work has been
reported on variations among extraction techniques with respect to toxiciry. In order to
examine this, we subjected several sediments contaminated with different types of toxicants to
pore water characterization studies. Pore water from all sediments studied was toxic to the
cladoceran Ceriodaphnia dubia. The pore water characterization experiments compared the
degree to which five commonly used extraction methods varied with respect to toxicity
recovered (as well as the types of toxicants recovered), and concentrations of dissolved and
paniculate organic carbon (DOC and POC), dissolved oxygen, and ammonia, as well as pH
and particle size. The sites studied were from the Saginaw River in Bay City, MI [polluted
with metals, oil/grease and associated poiycyclic aromatic hydrocarbons (PAHs), and
ammonia; Schubauer-Berigan et al. 1990], and the Keweenaw Peninsula in the Upper
Peninsula of Michigan (contaminated with metals, primarily copper, lead, and zinc; Ankley et
al., 1992).
m-s
-------
The methods we compared were centrifugation at 2500x$ and 10,000x$ under a normal
atmosphere in a refrigerated (4°Q centrifuge (with and without subsequent pore water
vacuum filtration through a 1.0 urn glass-fiber filter), pressure extraction (with a Teflon*-Iined
sediment press, using a 1.0 um glass fiber filter), syringe extraction (using a plastic 25 mL
syringe with an in-line 1.0 um glass fiber filter), and dialysis (using 5 mL cups sealed with a
0.45 um Nuclepore* membrane, allowed to equilibrate for 10 d at a volume of no greater than
4% of the total sediment water concentration; Hesslein 1976).
HL2.1 Saginaw River Sediment Pore Water Characterization
Dissolved oxygen (DO), pH and conductivity were similar among all extraction methods.
DOC was similar in the low-speed centrifuged sample and the pressure-extracted sample, and
was higher in the high-speed centrifuged sample (Table ni-2). Particles were virtually absent
in the samples filtered during collection (syringe-extracted and pressure extracted samples),
and had a similar size distribution for the high- and low-speed centrifuged samples. An oily
emulsion was noted in the two centrifuged samples, but not in the syringe-extracted and
pressure-extracted samples, both of which involve filtering as part of preparation. (Note: the
dialysis technique was not evaluated with the Saginaw River sample).
The toxiciry of the whole pore water samples appeared to be largely dependent on whether
the pore water was filtered prior to testing. Unfiltered samples (centrifuged at either 2500xg
or lOOOOxg) were approximately 4 times more toxic than any of the filtered samples. In the
ffl-6
-------
Table JH-2. Results of pore water characterization studies of Saginaw River (MI) sediments.
Parameter
Toxicity (TU1)
unfiltered
filtered
Centrifuged (2500 ff)
CHjClj extraction
of filters
DOC (ppm)
POC(%TOO
15
3.7
12
150
38.7
PREPARATION TECHNIQUE
Pressure
Centrifuged (10000 e>) Extracted
15
3.3
16
386
12.1
4.0
5.0
NT.2
135
0
Syringe
Extracted
3.2
3.2
N.T.
41.7
21.0
Sample emulsion
wet wt (g)
Particle size
(um; mean ± S.
[Metals] (pg/L)
Cr
Cu
Ni
Pb
Zn
D.)
unfiltered
filtered
unfiltered
filtered
unfiltered
filtered
unfiltered
filtered
unfiltered
filtered
0.90
1.70*12.9
2000
140
760
80
660
100
380
14
630
60
0.61
1.26*5.88
600
30
350
18
240
18
250
<5
350
<50
..
N.P.S
<5
<5
<3
<3
120
<120
<5
<5
<50
<50
tm
N.P.
9
5
12
5
20
17
<5
<5
<50
<50
1 TU, Toxic Units, 100%/LCjo (%)
2 N.T., not toxic.
3 NJP., particle concentration (frequency) approached zero.
ffl-7
-------
two pre-filtered pore water extraction methods (syringe and pressure extraction), toxicity was
essentially the same (4.0 and 3.2 TU1, respectively) as in the filtered centrifuged samples (3.7
and 3.3 TU for the low and high speeds, respectively; Table ni-2).
TIE work with the centrifuged samples demonstrated that much of the toxicity removed by
filtration (perhaps caused by compounds present in the oily emulsion) was extractable with
methylene chloride (for method, see Schubauer-Berigan and Ankley 1991), thus implicating
nonpolar organic compounds as toxicants. No toxicity was recovered in methylene chloride
extracts of filters of pore water that had been extracted by pressure or syringe and subse-
quently filtered (Table ffl-2). This indicates that much of the nonpolar organic toxicity was
removed from oily samples by the process of filtration during sediment extraction. This
toxicity may have comprised a substantial component of the sediment toxicity. Recent work
by others indicates that, even for non-oily samples, concentrations of nonpolar organic
compounds in pore water may be reduced by filtration. In a study of a pore water sample
spiked with dieldrin (1 mg/L, highest concentration), filtration through a 1 um glass fiber
filter reduced dieldrin concentrations to less than 1% of those in unfiltered pore water (P.
Kosian and A. Cotter, AScI Corp., ERL-Duluth, personal communication). Researchers in
other laboratories recently have noted excessive losses of nonpolar organic compounds due to
filtration of water samples (R. Ozretich, U.S. EPA, ERL-N Pacific Division, personal
communication), and in this laboratory, pore water from sediments highly contaminated with
DDT and metabolites (DDD, DDE) exhibited a loss of toxicity when filtered.
1 Toxic units (TU) are defined as 100%/LC50(%) for pore water or elutriate tests.
m-8
-------
Both of the centrifugarion methods showed a reduction of metal concentrations upon filtration
(chromium, copper, nickel, lead and zinc) by 85% to 98% (Table ffl-2). Nickel, lead, and
zinc were present at similar concentrations in the filtrate of the centrifuged samples as in the
syringe-extracted and pressure-extracted pore waters. Further filtration had no effect on metal
concentrations in the pressed and syringe-extracted samples. Considering the numerous
toxicants identified in these samples, although filtration removed a large portion of the "total"
metals from the centrifuged samples and very little from the other (pie-filtered) extraction
methods, it is difficult to discern whether any of the metals removed by filtration in the
centrifuged samples were actually bioavailable. This consideration was further addressed
with the metal-contaminated Keweenaw Waterway sediments.
QL2.2 Keweenaw Waterway Sediment Pore Water Characterization
DO, pH, conductivity, and DOC were similar for all the extraction methods (although the
DOC was lower for the dialyzed sample). POC concentration was highest for the low-speed
centrifuged sample, was about half this value in the high-speed centrifuged sample, and was
more than two orders of magnitude less in the pressure-extracted and dialyzed samples (Table
ffl-3). Particle size distributions indicated that the median particle size was generally smaller
in the pressure-extracted and dialyzed samples than in the centrifuged samples. No oily
emulsion was noted in either the sediment or the pore waters.
m-9
-------
Table III-3. Resulis of pore water characterization studies of Keweenaw Waterway (MI) sediments.
Parameter
Toxicity (TU)
unfiltered
filtered
DOC (ppm)
POC (mg/L)
Particle size
(urn; median + S.D.)
Metals
Cu,^
Pb-a—
Pbnh.™,
Zn^a,^
Z««h«i
NRJ unfiitered
filtered
Centrifuged
(2500 K)
18
11
27.1
123
2.18+32.8
luK/1.1 TU2
11000 111
6000 60
4800 4.8
2600 2.6
1300 21
500 7.9
7.6
6.4
Centrifuged
(10000 it)
5.6
1.4
27.1
61.3
52.4+25.9
lug/Li TU
1400 14
680 7
350 0.35
220 0.22
80 1.3
<20 <0.32
2.8
5.4
Pressure Extracted
<2
1.3
20.7
0.3
0.63+13.1
fug/Li HI
410 4.1
340 3.4
91 0.09
92 0.09
260 4.1
80 1.3
5.9
3.8
Syringe Extracted
13
1.9
NM1
NM
NM
fug/LI TU
3500 35
700 7
570 0.57
87 0.09
460 7.3
50 0.83
3.3
4.3
Dialyzed
5.6
1.2
(NC)
(NC)
6.3
3.7
lue/U
320
110
22
8
60
30
0
(11)
TU
3.2
1.1
0.02
0.01
1.0
0.5
0.75
1.3
NM, not measured due to inadequate sample volume
TU, potential toxic units of metal in sample, based on laboratory-determined metal LC^s at the test pH (99ug/L for
1000 Mg/L for Pb, and 63 ug/L for Zn).
NR, non-bioavailable ratio, calculated as the sum of potential metal toxic units divided by the actual toxicity in the
sample. A higher ratio indicates a greater proportion of unavailable metals in the sample.
IH 10
-------
The low-speed centrifuged sample and the syringe-extracted sample displayed the greatest
toxicity (18 and 13 TU, respectively; Table E-3). The dialyzed and high-speed centrifuged
samples displayed moderate toxicity (5.6 TU each), while the pressed sample contained less
than 2 TU. All samples (with the possible exception of the pressed sample) lost toxicity upon
filtration. The low-speed centrifuged, high-speed centrifuged, dialyzed, and syringe-extracted
pore waters lost 40%, 75%, 80%, and 85% of their respective toxicities after filtration. The
fact that the latter two samples lost toxicity due to filtration is surprising, considering that
these samples were filtered during the extraction procedure. This may indicate that oxidation
occurring in the pore water during and after extraction from the sediments, is causing
insoluble metal precipitation to occur in the sample. This phenomenon has been noted by
others in pore water extractions of trace metals in sediments (Carignan et al. 1985).
Copper, zinc, and lead were detected in whole and filtered pore water (Table ni-3). Metal
concentrations were highest in the low-speed centrifuged sample, followed by the syringe-
extracted, high-speed centrifuged, pressure-extracted, and dialyzed samples. These samples,
respectively, lost an average of 47%, 81%, 52%, 35%, and 63% of the whole pore water
metal concentration (the molar sum of copper, zinc and lead). In order to determine the
relative bioavailability of the metals in each of the pore water samples, potential LC^ values
due to metals were calculated based on single-metal C. dubia toxicity tests in dilution water
at the pore water pH (approximately 8.5). For each of the samples, the potential TUs were
calculated for each of the metals assuming total availability. Because the toxicity of several
canonic metals has been shown to be additive to C, dubia (Spehar and Fiandt 1986), the
m-11
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individual metal TUs were summed for each extraction method Since TEE had previously
shown that these metals were the sole acute toxicants at this site, a "non-availability ratio"
(NR) could be calculated for each extraction technique by dividing the total potential metal
TUs by the actual sample toxicity. A ratio close to 1 would indicate that the metals in the
sample, independent of their sample concentration, were actually all bioavailable, based on
dilution water LC^s for the metal The higher the ratio, the less likely it is that the metals
were totally available. Of all the extraction methods, dialysis gave the NR closest to unity
(0.75), followed by high-speed centrifugation (2.8) and syringe extraction (3.3). The sediment
press and low speed centrifugation procedures resulted in higher NR values (6.2 and 7.3,
respectively), suggesting that these methods extracted relatively high concentrations of
unavailable metals. In a study comparing various pore water extraction methods for metals,
Carignan et al. (1985) also found that centrifugation at low speeds (5000 rpm) recovered
higher concentrations of copper, zinc, and organic carbon than either centrifugation at higher
speeds (10000 rpm) or in situ dialysis.
HI.2.3 Recommended Pore Water Preparation Method
Based on sample volume considerations for TIE work, as well as results of the studies above
and reported by others (e.g., Capel 1986; Schults et al. 1991), we recommend that pore water
be isolated via centrifugation without subsequent filtration. Although the specific mechanism
is not known through which filtration removes toxicants from pore water samples (e.g.,
removal of contaminants associated with particles, filtration of oxidized metal-ligand
ffl-12
-------
complexes, sorption to the filter, etc.), data from our laboratory clearly indicate that any pore
water isolation technique that requires or incorporates filtration as part of the extraction
process is likely to remove bioavailable metals and nonpolar organics. Our data also suggest
that speeds ranging from 2,500xg to lO.OOOxg are suitable for pore water preparation. The
lower speeds, however, may result in the presence of unavailable metals in pore water. The
speed of centrifugation has been shown in other research not to affect the partitioning of
nonpolar organics, such as PCBs, into pore water (Capel 1986). Finally, to reduce artifacts
induced by temperature fluctuations (Bischoff et al. 1970), we recommend that pore water
samples be prepared under cool (ca., 4°C) conditions. This can be achieved either through
the use of a refrigerated centrifuge, or through sample preparation in a controlled temperature
room (e.g., walk-in cooler).
In our pore water characterization studies, there are several factors which we did not address
(for example, the effects of oxidation on speciation of pore water nutrients and contaminants).
Further research is required to extend existing knowledge of pore water's suitability for
evaluating sediment toxiciry.
in.3 Use of Benthic Species for Aqueous Testing
Another facet differentiating sediment TIE from effluent TIE involves the selection of species
for testing. Sediments contain epibenthic and benthic species and communities quite different
from the pelagic species used in effluent toxicity and TIE studies. Several common benthic
ffl-13
-------
species [e.g., Rhepoxynius dbronius (Swartz et aL 1982), H. azteca (Nebeker and Miller 1988,
Nelson et aL 1991), Chironomus riparius and Chironomus unions (Adams et al. 1985, Nelson
et aL 1991), among others] have been determined to be sensitive to a wide variety of
sediment contaminants, based on bulk phase toxicity tests (see Giesy and Hoke 1989 for a
review of sediment toxicity test species). Therefore, in addition to using water column
species for sediment TIE, for some studies it may be desirable to subject benthic organisms to
the TIE pore water manipulations to identify the compounds responsible for the observed
sediment toxicity. This is especially critical if the absence of one or more specific benthic
species of concern is a primary factor triggering the TIE studies. Although most TIE studies
have been performed with pelagic species, such as C, dubia or fathead minnow, the TIE
methods may be appropriately used (with few modifications) for any species amenable to
aqueous testing in small volumes. Thus, methods were developed (Schubauer-Berigan and
Ankley 1991; Ankley et al. 1991c) and are described here and in Section IV for testing
benthic species, such as H. azteca, C. tentans and L. variegatus, in small-volume aqueous
samples. We also address the relative sensitivity of several different benthic and pelagic
species to pore water from various sediments contaminated with several types of toxicants, as
well as to aqueous solutions containing single chemicals such as metals, nonpolar organic
compounds and ammonia, all common sediment contaminants.
Because TIEs involve comparing the effects of various physical and chemical manipulations
on sample toxicity to a baseline toxicity value, it is especially important to have good
performance control and manipulation blank sample survivorship in the toxicity tests (U.S.
m-14
-------
EPA 1991a;b). We have found that it is relatively easy to obtain satisfactory (80-90%)
control survival in 96-h aqueous exposures for the epibenthic/benthic species H. azteca and C.
tentans by placing an artificial substrate, consisting of a small (2.25 cm2) Nitex* screen, into
the test chamber. This prevents floating organisms, and appears to reduce cannibalism among
H. azteca. We have had difficulties in testing several C. tentans individuals in the same test
cup; we frequently observe cannibalism in both pore water and control water in water-only
tests with S organisms in 30 mJL L. variegatus, an aquatic oligochaete that we also have
used for TIE work, survives very well in small-volume aqueous tests without the use of an
artificial substrate. If other epibenthic or benthic species are to be used in die TIE, consider-
ation and preliminary study should be given to using the organism in aqueous phase tests
before commencing with the TIE. These considerations should include, but are not limited to,
the routine availability of the organisms, acute sensitivity of the organisms to toxicants,
requisite test volumes, optimal or feasible test temperature, light conditions, number of
organisms allowable per test chamber, and a feeding regime (if needed).
Some epibenthic/benthic species with which we have worked appear to be comparable in
sensitivity to many contaminants as the more commonly used pelagic species, C. dubia and
fathead minnow. In a study comparing the relative sensitivity of two epibenihic or benthic
(H. azteca, L. variegatus) and two water-column (C. dubia, fathead minnow) species to
acutely toxic pore water and elutriate from seven contaminated sites, we found H. azteca to
be most sensitive to both pore water and elutriate, followed by C. dubia and the fathead
ffl-15
-------
minnow. L variegatus was the least sensitive to both pore water and elutriate samples (Fig.
m-1; Ankley et aL 1991c).
These observations with field samples also have been corroborated by comparison of the
sensitivity of these species to the metals lead, zinc, copper, nickel and cadmium (Table ffl-4).
Although variations in metal toxicity to the species were observed at different pHs, H. azteca
were generally as sensitive or more sensitive than C. dubia, which were in turn usually more
sensitive than fathead minnows.
One interesting phenomenon that we have noted is that the sensitivity of H. azteca to
ammonia does not appear to be influenced by pH (at least, within the pH ranges of our tests;
pH 6 to 8.7; Table 1H-4). Thus, while for fathead minnows, C. dubia and L. variegatus, un-
ionized ammonia (more prevalent at higher pHs) is the more toxic form of ammonia (U.S.
EPA 1988; 199la), it appears for H. azteca that ammonium ion may be at least as toxic as
the un-ionized form. [Note that at pH 8.0, only 5.38% of the total ammonia is present in the
un-ionized form; at pH 8.5 (which approximates pH / for many pore waters), 15.2% of the
total ammonia is present as un-ionized ammonia.]
ffl.3.1 Selection of TIE Species
In bulk sediment toxicity tests, the route of exposure for upper water column species to
toxicants is probably through the water overlying the sediment. In contrast, epibenthic and
m-16
-------
Figure HM. Relative sensitivities of Pimiphales promt las, Ceriodophnia dubia, Hyalella
azteca, and Lumbriculm variegatus to sediment pore water and sediment
elutriate. Error ban indicate the standard error of the mean for the ranks.
Letters in parentheses above the bars indicate differences in ranks among the
species; means with different letters differed significantly (p < 0.05) from one
another (from Ankley et aL 1991c).
i '
C/)
-------
Ttbie m-4. Trends in metal and ammonia toxicity with respect to test pH. LC^s (expressed as
ug/L of metal or mg/L of ammonia) were determined at 48 b for C. dubia, and 96 h
for fathead minnows, H. azteca, and L variegaau. Tests were perfonned in very hard
reconstituted water (see text).
Metal Species
C. dubia
Zn H. azteca
P. promelas
C. dubia
Ni H. azteca
P. promelas
C. dubia
Pb H. azteca
Fathead minnow
C. dubia
Cu Fathead minnow
H. azteca
C. dubia
Cd Fathead minnow
H. azteca
N:NH3 H. azteca
Lumbriculus varieganu
pH6-6J
LC,
>530
1200
830
>200
1960
>4000
280
<90
1410
10
15
17
563
54
228
20* (9.0)
>1000
pH 7-7 .5
LC,
360
1500
333
137
1940
3360
>2700
>5400
>5400
28
44
__i
350
74
«
232 (14)
62
pH 8-8.5
LC*
95
289
502
13
890
3080
>2700
>5400
>5400
201
>200
87
121
<5
4-15
211 (12)
13
1 Test not performed.
2 Value represents the mean of 6 LC50 values determined at that pH. with the standard deviation in
parentheses.
ID-18
-------
benthic test species are additionally exposed to in-place sediment toxicants through both pore
water and direct sediment contact Therefore, when using sediments in TIE studies we
proceed via a two-phase approach. First, bulk sediment tests designed to evaluate the toxicity
of in-place sediments are performed using benthic organisms, in order to mimic exposure to
in-place sediment toxicants (Ankley et al. 1991c). Then, pore water toxicity tests with
benthic organisms (and, if necessary, water column species; see below) are conducted to
demonstrate that pore water samples also are toxic (bulk sediments will rarely be toxic in the
absence of pore water toxicity; Table ffl-1, Ankley et al. 1991c). Although benthic species
should be used to evaluate the acute toxicity of bulk sediments and corresponding pore water,
in some instances it may be desirable or necessary to use pelagic species for some portion of
the TIE because of limitations in the availability of benthic organisms. If this is the case, the
same benthic species that were used to establish initial bulk sediment and pore water toxicity
should be employed in Phase HI (confirmation phase) of the TIE to ensure that the com-
pounds responsible for pore water toxicity to the primary TIE species also are causing toxicity
to the benthic organism of concern.
In selecting species for the bulk sediment/pore water tests and/or the TEE manipulations,
toxicity should be evaluated initially for several species with differing sensitivities to various
compounds. Because of its relatively great sensitivity to a wide variety of compounds, we
have found H. azteca to be a good choice for the sediment and pore water testing and TIE
manipulations. This species is additionally functional, depending upon the strain available,
because it may be tested in both freshwater and estuarine environments. Also, H. azteca is
m-i9
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relatively simple to culture; current (Nelson et aL 1991) or pending (U.S. EPA, in prepara-
tion) documents are available describing culturing and testing procedures. We also have used
a variety of pelagic species (cladocerans, fishes) as well as other types of benthic organisms
(oligochaetes, chironomids) for TEE studies. Regardless of the species used in the bulk of the
TIE studies, however, sediment toxicity should be confirmed with the most sensitive benthic
species tested, or the species of concern (if applicable).
IIL4 Test Volume Considerations
Sediments typically contain 30-50% water (on a weight basis). Of this, approximately 50% is
extractable as pore water (depending on method of extraction). Therefore, pore water
recovered by centrifugation from sediments averages approximately 20% of the total sediment
weight. Obviously, sampling constraints will limit the volume of sediment that can be
collected from individual sites. Consequently, sediment studies inherently differ from effluent
studies in the volume of aqueous sample available for the TEE process. As a result, we have
used a variety of techniques to conserve sample volume through all phases of the TIE.
The first volume consideration to be addressed is the toxicity test volume. Most of the
species with which we have worked (Le., fathead minnow, C. dubia, D. magna, D. pulex, H,
azteca, and L. variegatus) are amenable to testing in small sample volumes (e.g., 5 organisms
per 10 mL replicate). Conserving pore water by testing in small volumes is essential if
sediments are available in limited quantities.
in-20
-------
Another useful tool for reducing the volume (and effort) required for TIE testing with pore
water is the simultaneous testing of two species in the same test chamber. This method
provides the additional advantage of minimizing differences in chemical test parameters (e.g.,
pH) during the testing of two species, and can provide valuable information regarding the
relative sensitivity of the species to pH-dependent toxicants, such as ammonia or metals, in
the sample. We have successfully tested C. dubia and fathead minnow in the same 10 ml
volume throughout an entire Phase I evaluation. We do not recommend, however, such
simultaneous testing of species until it has been determined that the species are compatible,
and that test conditions (e.g., adequate DO) can be maintained throughout the test
The effluent Phase I lit iiwuial recommends using all the manipulations described for each
sample, which require a total volume of 2-3 L (U.S. EPA 1988; 1991a). However, because of
the limited volumes of pore water often available, we frequently omit steps such as the pH-
adjusted (pH 3 and 9) C,, manipulations, at least in the initial Phase I manipulations (this
approach also has been recommended in the chronic TIE document; U.S. EPA 1991b). These
require large volumes of sample (i.e., 200 mL each), and in our experience usually do not add
significantly to the insights gained through the other TIE manipulations (one exception would
be for a sample containing ionizable organic compounds). Thus, we recommend using the
Phase I manipulations in a tiered manner to conserve sample volume and time. A tiered
approach, in which all pH 3- and pH 9-adjusted manipulations are deleted to conserve sample
and effort, has been recommended for chronic Phase I TIE. For acute sediment TIEs,
however, we recommend performing the pH 3- and pH 9-standing, aeration and filtration tests
in-21
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because of their relatively low volume requirements (Le., 35 mL each), and the potential for
useful information to be gleaned from these tests (Schubauer-Berigan et al., 1992).
Pore water storage considerations are of additional importance when volumes available for
TIE are limited. We recommend storing the extracted pore water for no more than 48 h prior
to use because we have observed marked temporal fluctuations in toxicity, metal availability
(e.g., through precipitation during storage), and concentrations of volatile compounds. In fact,
storage effects on toxicity actually can serve as a useful TIE "manipulation" for implicating
hydrogen sulfide as a sample toxicant, as this compound is highly volatile and easily
oxidized, and usually disappears after a few days' storage. Because storage time should be as
brief as possible, the volume required for immediate testing must be determined prior to
extracting the pore water, and only the amount needed for those tests should be prepared.
The possibility exists that the pore water matrix may vary among different extractions;
therefore, it is important to carefully track the toxicity (and concentrations of any suspect
toxicants) each time a new batch of pore water is prepared. In our experience, this has not
been a particularly troublesome aspect of sediment TIE.
ffl.5 Common Sediment Contaminants: Ammonia, Metals and Hydrogen Sulfide
Many sediments contain toxic concentrations of ammonia and hydrogen sulfide (Ankley et al.
1990a, Schubauer-Berigan et al. 1990, Schubauer-Berigan and Ankley 1991). Although these
contaminants are to a large extent derived from natural microbial processes, in many cases the
IH-22
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accumulation of these compounds may be exacerbated by the presence of other, anthropogen-
ically-derived materials (e.g., high levels of organic matter). In addition, ammonia concentra-
tions in sediments may be increased by direct loading through effluent discharges. In either
case, the sediment researcher is often confronted with these two contaminants in a sample,
sometimes simultaneously, and must ascertain whether any other toxicants are present
Further, metals also are common sediment toxicants; unfortunately the simultaneous presence
of metals and ammonia or hydrogen sulfide can result in confusing TIE results because the
pH-dependent behavior of metals can mimic that of ammnnia or hydrogen sulfide. We
encounter this situation routinely during sediment TIE, and in concert with others in the
effluent research group, have developed several techniques for circumventing the difficulties
associated with separating the effects of various toxicants in samples. Examples of these
methods are: (a) performing TIE tests at altered pH to avoid the effects of one toxicant, (b)
testing alternative species with differing sensitivities to certain of the compounds, (c)
performing TIE manipulations at concentrations below the effects concentration for the
ammonia and/or hydrogen sulfide, and (d) performing specialized techniques for recovering
volatile or filterable toxicants (e.g., hydrogen sulfide and metals or organic compounds).
Each of these approaches, and its applicability for assisting in the identification of common
sediment contaminants, is described below.
UL5.1 The Graduated pH Test
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Adjusting the test pH of samples subjected to various manipulations can be a very powerful
tool in sediment TEE. The bioavailability and/or toxicity of ammonia, hydrogen sulfide and
metals are highly pH dependent, even within the relatively narrow range of physiological
tolerance for most pelagic, epibenthic and benthic organisms (e.g., pH 6-9). Results of such
toxicity tests are often a reliable starting point for determining, for instance, whether toxicants
other than ammonia are present in the pore water. Often, the results of the graduated pH test
will be the only substantive clue to the nature of the toxicants in the sample, and we
recommend that careful attention be given to maintenance of test pH and water quality
parameters during this test
The un-ionized form of ammonia (NH3) is generally thought to be more toxic than the ionized
form to aquatic organisms (H. azteca appears to be an exception to this observation; cf.,
Section in.3). While the un-ionized form of the ammonia is much more prevalent at pH 8
than at pH 6 (5% vs. 0.0568% un-ionized), the un-ionized ammonia itself is more toxic at pH
6 than at pH 8 (U.S. EPA 1985). The net result of these pH effects on bioavailability and
toxicity is that the same amount of total ammonia is approximately 3 times more toxic at pH
8 than at pH 6 (see U.S. EPA 1988). Because we have found that the initial pH of pore
water samples drifts to 8.5 or greater, we often perform the graduated pH test at pHs of 6.5,
7.5 and 8.5. If ammonia is the sole sample toxicant, the sample may be non-toxic at the
lowered pHs in the graduated pH test
IH-24
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The toxicity of hydrogen sulfide is also highly pH dependent: total sulfides are approximate-
ly 12 times more toxic at pH 6.5 than at pH 8.7, despite the fact that molecular H,S is more
toxic at pH 8.7 than at lower pH (Broderius et al. 1977). Often in the graduated pH test, we
have observed enhanced toxicity at pH 6.5, which tends to disappear after prolonged storage
of the pore water (Le., longer than 1 day). Because of the high volatility and potential for
oxidation of hydrogen sulfide, this compound tends to be relatively unstable in aqueous
samples.
Metals are another class of compounds whose toxicity and/or bioavailability are dependent on
pH within the range of the graduated pH test Recent work at this laboratory with pelagic
(C. dubia and fathead minnow) and epibenthic/benthk (H. azteca and L. variegatus) species
indicates that zinc and nickel show increased toxicity at pH 8.5 relative to that at lower pHs
(Table ffl-4). Lead and copper show the opposite trend, and are more toxic at pH 6.5 than at
pH 7.5 or pH 8.5 (Table HI-4). Cadmium appears for fathead minnows to be more toxic at
pH 6.5 and pH 8.5 than at neutral pH. Thus, the graduated pH test may serve additionally to
distinguish between toxic and nontoxic metals when several are present simultaneously in a
pore water sample.
One difficulty with the graduated pH test is that interpretations may be confounded when
several pH-dependent toxicants (e.g., one or more metals, sulfides, and/or ammonia)
simultaneously occur at toxic concentrations in a sample. This is because different combina-
tions of these toxicants (e.g., copper, lead and HjS, or zinc and ammonia) may tend to mask
ID-25
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other compounds possessing the same trends in pH-dependent toxicity. However, by
simultaneous use of the graduated pH test and other TIE manipulations (i.e., EDTA test,
sodium thiosulfate additions, and ion-exchange chromatography, or compound spiking), the
identification of the individual toxicants can be achieved.
HL5.1.1 Methods of pH Control
Pore waters are usually well buffered; adjustment of the sample pH alone generally does not
sufficiently maintain pH because CO, in the ambient test environment tends to lead to
reestablishment of the equilibrium (initial) pH. In initial guidance for TIE (U.S. EPA 1988),
the addition of acids/bases was recommended for pH control in the graduated pH test Since
then, two additional successful methods of pH control have been used in both sediment and
effluent TTEs to negate this effect The first incorporates methods to control CO2 exchange
between air and water, thus steering the bicarbonate buffering system of the test water to an
altered pR This generalized approach may or may not require initial acid/base adjustments,
and consists of CO2 injections into the controlled airspace above the sample. The second
method utilizes hydrogen-ion buffers designed to be inherently nonreactive and non-toxic to
biological tissues and organisms (Ferguson et al. 1980; Neilson et al. 1990) for controlling pH
at approximately the pK, of the buffer.
When small organisms are used (e.g., C, dubia), a very simple method for controlling the pH
is the so-named "closed cup" technique (U.S. EPA 1988). Using this method, sample
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dilutions are acid/base adjusted and added in sufficient quantity to fill a small chamber, which
is then sealed with a cover to remove headspace. The pH usually holds well for 48 h using
this method; however, we have experienced difficulties in maintaining adequate DO concen-
trations for both H. azteca and fathead minnows in 96 h tests. Another disadvantage with the
closed cup pH control method is that the pHs can be read only at test termination (or when
there is total mortality in the test chamber), as it is very difficult to eliminate headspace and
maintain pH once the closed chamber has been opened.
A second method for controlling pH involves manipulating the headspace concentrations of
CO;, and is more effective when larger organisms such as fathead minnow, L. variegatus, H.
azteca, and C. tentans are used. This technique employs rectangular glass chambers with a
small hole bored into the end (U.S. EPA 199la). We have found most aqueous extracts of
sediments to be sufficiently buffered to require acid/base adjustment before headspace gas
adjustments are marie. Chambers are flushed with CO2 and then stoppered. This method is
useful because the pH may be taken several times during the exposure period, provided that
chambers are re-flushed with CO, after each exposure to ambient air. In addition, smaller
volumes of pore water or elutriate are required for CO^ chambers than for the closed cup
method, and DO concentrations can be more easily maintained.
A third pH control method involves the use of zwitterionic hydrogen ion buffers designed to
be nonreactive with biological tissues (Ferguson et al. 1980). Three of these buffers have
been shown to exhibit low toxiciry to several aquatic organisms (Neilson et al. 1990). Mes
in-27
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(2-(N-morpholine)ethane-sulfonic acid) buffers the pH from 6.0-6.2. Mops (2-(Af-morpho-
line)propanesulfonic acid) and Popso (pipenmne-M^-bis(2-hydroxypropanesulfonic acid)
buffer solution pHs at 7.0-7.2 and 7.8-8.2, respectively. We also have found the buffers to be
relatively nontoxic to the benthic, epibenthic and pelagic organisms we use routinely for TIE
work (i.e., fathead minnow, C, dubia, H. azteca, C. tentans, and L, variegatus; Table ffI-5,
U.S. EPA 1991a). We have tested these buffers extensively with different types of com-
pounds to determine their efficacy in TIE. The Mes buffer appears to interfere slightly with
the toxicity of some metals (e.g., lead and copper LCjoS increased by 2x for C. dubia when
Mes was used) but does not impede the ability of EDTA or sodium thiosulfatc to chelatc
metals (Table ffi-6). The effect of the buffers on metal toxicity during the TIE will probably
be slight if there are more than 2 TU due to the metals in the samples. We now use the
buffers Mes and Mops routinely with acutely toxic pore water samples (use of the Popso
buffer has been unnecessary for the pore water samples with which we have worked, because
the initial pH of these pore waters is close to 8.5 without adjustment). However, certain
caveats must be attached to the unexpurgated use of these buffers. First, the effective buffer
concentration (i.e., the concentration required to maintain the desired pH) is hardness and/or
alkalinity dependent The effective buffer concentration needed in a single chemical dilution
water test (generally 2.5 to 4 mM) may be an order of magnitude lower than that required to
effectively buffer pore water or effluent samples. In addition, studies with non-toxic pore
water and effluent samples have demonstrated that the buffer toxicity decreases in more
complex samples (data not shown). Second, we have encountered some pore water samples
whose toxicity is affected by the buffer. When this occurs, it may be necessary to use other
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Table DI-5. Sensitivities of C, dubia, fathead minnow, H. aztecd, and L. varieganu to the
pH-control buffers, Mes, Mops and Popso. Test duration was 48 h for C.
dubia, and 96 h for all other species.
Species
C. dubia
Fathead minnow
H. azteca
L. variegana
Water type
SW1
VHW2
SW
VHW
VHW
VHW
Mes
38
62
71
>100
46
>100
Mops
62
57
77
>100
29
>100
PODSO
19
23
77
100
13
100
1 SW, Soft water
2 VHW, Very hard reconstituted water
IH-29
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Table III-6. Sensitivity of C. dubia to certain metals (tested using different pH adjustment/control techniques), and ability of EDTA
to chelate metal toxicity in the presence and absence of pH-control buffers.
Compound
Pb
Cu
Zn
pH control
Technique
CO2 adjustment
Mes buffer
closed cup
Mes buffer
closed cup
Mes buffer
closed cup
Mops buffer
closed cup
Popso buffer
24 h
480
>1000
31
41
534
820
253
339
78
136
fiH
6.3
6.3
6.2
6.3
6.7
6.2
7.2
7.3
8.2
8.2
LAo (jig/L)
48 h
430
580
12
22
328
616
205
252
70
78
EH
5.8
6.3
6.3
6.3
6.7
6.2
7.2
7.3
8.2
8.2
IEDTA) (mg/L)
required to remove toxicitv
<51.2
<51.2
<51.2
<51.2
<51.2
<51.2
<51.2
<51.2
<51.2
<51.2
111 30
-------
methods of pH control. Thus, initial tests with the pore water samples should be designed to
examine two facets of pH-affected toxicity: first, the effective buffer concentration for the
particular sample should be determined; and second, the sample toxicity using the buffers should
be compared to that with other pH-control methods, in order to detect whether the buffers
themselves interfere with sample toxicants. Once these issues have been resolved with the
sample, use of the pH-control buffers can offer several important advantages in pore water
studies: the buffers generally maintain more precise and predictable pH values than the CO2-
control methods; small volumes (10 mL) can be used with the buffers, and DO is not difficult to
maintain.
HL5.2 Alternative Species Testing
As mentioned previously, the sensitivity of different species can vary widely for different types
of compounds. Thus, relative species sensitivities can be an effective tool for differentiating
between the effects of different compounds in pore water. For example, if ammonia and some
metal more toxic at high pH (e.g., zinc) are both present in a sample at potentially toxic
concentrations, the graduated pH test would be useless in discriminating between the effects of
the two suspect compounds (i.e., both would exhibit increased toxicity at elevated pH). Thus, it
is prudent to test, in tandem, several species possessing differing responses to these types of
contaminants. Fathead minnows, for instance, are more sensitive to ammonia and hydrogen
sulfide than C. dubia, and are comparatively insensitive to some metals (Table ffl-4). L
variegatus is another species very sensitive to ammonia, but not to metals or certain nonpolar
organic compounds, while H. azteca is sensitive to both ammonia and metals. These types of
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comparisons may be useful throughout all stages of the TIE to determine whether more than one
toxicant may be present in a pore water sample, or to ascertain whether a manipulation designed
to remove one toxicant (e.g., zeolite removal of ammonia) actually removed another toxicant
(zinc).
HI.5.3 Toxicant Dilution Testing
Frequently, no Phase I manipuktion completely removes toxicity from samples containing
several toxicants, and techniques (e.g., use of cation exchange resins) that are designed to
remove one toxicant (canonic metals) also remove another toxicant (ammonia). In these cases,
another method of identifying and assessing the relative contributions of the individual toxicants
consists of performing sample manipulations below the effective concentration(s) of one or more
of the toxicants. Using a hypothetical pore water contaminated with ammonia and zinc as an
example, if a greater amount of toxicity appears to be due to zinc than to ammonia (e.g., 10 TU
vs. 2 TU), an EDTA test performed at a 100% or 50% pore water concentration would not be
expected to remove sample toxicity due to the presence of toxic amounts of ammonia. If,
however, EDTA were added to a sample dilution at which ammonia would not be expected to
cause toxicity (e.g., 25%) the toxicity contributed by zinc would be likely removed from the
sample, thus indicating the presence of more than one type of toxicant A similar approach
could be used for sulfide and another compound more toxic at low pH (e.g., lead or copper).
HL5.4 Recovering Volatile and Filterable Contaminants
ffl-32
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As was mentioned previously (cf, m.5.1), hydrogen sulfide (r^S) is volatile at lowered pHs.
This characteristic may be used as an effective Phase n tool for isolating and measuring H2S
through the volatile toxicant transfer experiment Bioavailable metals and nonpolar organics are
often removed by filtration (cf., Section UL2). Recovery of these compounds from the filter
may be achieved through the use of appropriate solvents. Techniques for isolating hydrogen
sulfides, metals and nonpolar organic compounds from contaminant mixtures are described
below.
HL5.4.1 Volatile Toxicant Transfer Experiment
This technique permits the isolation of volatile compounds, which may be contributing to the
toxicity of the pore water sample, and consists of a closed-loop sparging system that transfers
volatiles from a sample aliquot to a dilution water aliquot (U.S. EPA 1988; 1991a). The setup
can be performed at equilibrium pH to detect neutral volatiles, but a more useful application for
detecting H2S toxicity uses a "purge and trap" system. This setup relies on the higher concen-
trations of HjS (the volatile form of the compound) as compared to HS~ at pH 3 than at ambient
pH; in this system, the sample is adjusted to pH 3 and (in an airtight apparatus) sparged with
nitrogen, which is subsequently bubbled through a dilution water solution at pH 9. In theory,
all the HjS initially present in the sample should be transferred to the dilution water trap;
however, the degree of air-tightness of the system limits its efficiency due in part to the ready
oxidation of r^S to sulfate. Thus, we recommend using a smaller volume of trap water relative
to the sample volume in order to concentrate any volatiles mat might be present. The resulting
trap water may be analyzed for H^S and tested for toxicity (at either pH i or pH 6) to detect H2S
m-33
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toxicity. This procedure, when successful, serves as a powerful isolation and identification
technique for HjS, and can be used in tandem with other procedures to provide a complete
depiction of the toxic components of a sample. One confounding factor in the effort to isolate
and identify HjS toxicity is the fact that the detection limit for H£ measurements via the
colorimetric (methylene blue) method (0.05 mg/L; APHA 1980) is significandy greater than its
LCj0 for some species, such as fathead minnows (0.015 mg/L; Broderius and Smith 1977;
Broderius et aL 1977). Thus, while the volatile toxicant trap method may successfully isolate
HjS from the pore water sample, the chemical analyses may not be sufficiently sensitive to
detect its presence at toxic concentrations in the sample.
HI.5.4.2 Recovering Filterable Toxicity
Bioavailable nonpolar organic compounds and metals can be removed from pore water samples
via filtration with glass-fiber or nylon filters. In some cases these classes of compounds may be
recovered individually from the filter by sequential extraction with appropriate solvents
(Schubauer-Berigan and Ankley 1991). Filters can be extracted first (with or without sonica-
tion) with a solvent such as methylene chloride to remove nonpolar organics. Filters then are
removed from the solvent, and set aside for further extraction. A solvent transfer (from
methylene chloride to methanol to water) allows the eventual testing of the solvent extract in
either dilution water or non-toxic sample matrices. If the latter test solution can be used, this
may give an effective estimation of the sample matrix effects on compound availability.
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Subsequent extraction, including sonication, of the filters with an acidic (pH 3) dilution water or
non-toxic sample can provide information on toxic metals that may have been removed by
filtration. Because the extraction is likely to be somewhat inefficient, it is helpful to concentrate
the extraction water relative to the sample volume passed over the filters. The resulting
extraction water can be tested for toxicity (perhaps using EDTA to confirm metal toxicity), and
metals measured in the sample.
One concern with such extraction procedures is mat the metals or nonpolar organic compounds
recovered in the extracts may not be representative of those actually available in the sample.
Testing the extracts in nontoxic sample matrices addresses this issue to some extent; however,
note that these extraction procedures merely represent initial steps for the characterization and
identification of the toxicants, and must be followed by other identification and confirmation
procedures.
EQ.6 C1S Fractionation Considerations
Many of the types of nonpolar organic compounds that accumulate in sediments are less polar
than those typically found to be toxic in effluents. For example, we have identified toxic Cu
SPE fractions containing benzenes, PCBs, PAHs and long-chain aliphatic hydrocarbons from
sediment pore water samples from die Illinois and Saginaw Rivers (Schubauer-Berigan et al.
1990; Schubauer-Berigan and Ankley 1991). In these studies, we were unsuccessful in
recovering nonpolar organics from the Clt SPE column using the methanol/water scheme
recommended in the Phase 0 TIE manual (U.S. EPA 1989a); we used instead an increasingly
ffl-35
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nonpolar methylene chloride/methanol elution series to extract the more nonpolar compounds
that appeared to be causing the sample toxicity.
More recent work in our laboratory has characterized effective fractionation techniques for
isolating the highly nonpolar organics present in sediment pore water. Work with standards
containing a series of compounds with log K^ values ranging from 3 to 8 confirmed that the
C,, SPE fractionation techniques do not provide predictable recoveries or separations for the
more nonpolar compounds (unpublished data). We have observed mat for compounds with log
K,, values of greater than 5, one "peak" of chemicals is recovered in the 100% methanol
fractions, and another peak is recovered in the less-polar methylene chloride/methanol fractions.
However, the same standard fractionated by high-performance liquid chromatography (HPLQ
shows that increasingly nonpolar compounds are sequentially recovered, and more predictably,
in the less polar fractions. We do not know precisely why the two methods recover high log
Kow compounds in such dissimilar manners; however, we speculate that it may result from the
differences in the column packings used in the two techniques. We are currently performing
similar analyses on standards that have been spiked with oil and grease to determine whether
these substances interfere with the fractionation of high log K^ nonpolar organics.
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IV. Sample Collection, Preparation and Initial Toxicity Tests
IV.l Shipping
After collection and homogenization, wet sediment samples should be stored in sealed giq*? or
plastic containers that have been acid-washed. Headspace in the containers should be kept to a
minimum to prevent oxidation or volatilization of compounds. Samples should be held in the
dark at 4°C until shipped. Sediments should be shipped at 4°C to the testing laboratory as soon
as possible after collection. We have had good success shipping and receiving sediments in
insulated coolers, via overnight service.
IV.2 Arrival and Storage
Upon arrival of sediments in the testing laboratory overlying water should be decanted and
discarded. The temperature and general observations concerning the appearance of the samples
should be recorded upon receipt Samples should be stored at 4°C, and must not be frozen.
Re-homogenization of the sediments should be conducted before all pore water/elutriate
preparation or bulk sediment assays. Sediments should be stored for the minimum amount of
time possible before toxicity testing and TIE analyses commence (preferably less than 14 d).
Due to the protracted nature of some TTEs, sediment samples may require holding for longer
than 14 d to complete necessary analyses. Unfortunately, the effects of storage time on
sediment toxicity and chemistry will be quite sample-specific, so it is impossible to define
standard guidance for holding times. Thus, because the potential exists for sediment samples to
IV-1
-------
change, particularly upon extended storage, baseline (whole pore water or elutriate) toxicity tests
must be performed routinely any time that the samples are used. When performing these
baseline tests, routine chemical parameters such as DO, pH, alkalinity, hardness, and when
appropriate (i.e., when toxic concentrations may be present), metal and ammonia concentrations
should also be measured. Marked changes in pore water or elutriate toxicity or chemistry over
the course of the TIE should cause data to be "flagged", and may indicate that fresh sediment
samples should be collected. Some sediment toxicants (e.g., H^S) are volatile or may be subject
to oxidation. If their presence is suspected (e.g., through the use of the volatile toxicant transfer
experiment) these also should be monitored as pore water or elutriate is isolated and tested.
IV.3 Test Fraction Preparation
Preparation of pore water or elutriate for initial toxicity tests should be conducted as soon as
possible after arrival of sediments in the laboratory. Certain chemical parameters in each
fraction should also be measured at this time. These include pH, hardness, alkalinity, conductiv-
ity, total ammonia and DO.
IV.3.1 Pore Water Preparation
Pore water is prepared by centrifuging homogenized wet sediment samples. The sediment
should be spun at 2,500 to 10,000 x g for 30 minutes at 4°C. The higher speeds may be
preferable, particularly if metals are suspect toxicants. Between 10% and 50% of the total
volume of homogenized bulk sediment can be expected to be recoverable as pore water. The
IV-2
-------
variation can be accounted for by the physical characteristics of the bulk sediment (e.g.,
sediments with a high concentration of sand will contain less pore water than a sediment
containing large amounts of organic material). If a centrifuge is not available, the analyst can
use other pore water isolation techniques, provided they do not include any type of filtration.
The centrifuge should be allowed to reach operating temperature (i.e., 4°Q before beginning
sample centrifugation. Note that rotational speed (which generally is indicated on most
centrifuges) is not equivalent to gravitational force; the two are interconvertible, however, with
equations specific to the centrifuge and rotor used. Prior to centrifugation, the overlying water
from the wet sediment should be decanted and discarded along with any large debris before
homogenization. Homogenize the sample, using a clean teflon-coated metal spatula, until a
slurry is achieved. Transfer the homogenized sediment to centrifuge bottles. The bottles should
be acid- and distilled water-rinsed before each use, and should be made of either plastic
(polypropylene), stainless steel, glass, or teflon coated plastic. The bottles should be of a size
(preferably >230 mL) large enough to extract the necessary volume of pore water (e.g., SO to
125 mL) per bottle. After centrifuging for 30 min., gently decant (or aspirate) the resulting
supernatant into a separate container. Store the pore water at 4°C in the dark until used. Do
not filter the samples.
IV-3
-------
IV.3.2 Elutriate Preparation
Generally, elutriate samples are prepared by mixing (e.g., on a shaker table, tumbler or rolling
device) one volume of homogenized wet sediment with four volumes of a dilution water for 12
h (U.S. Army Corps of Engineers/U.S. EPA 1977). The mixture is then centrifuged for 30 min.
at 2,500 to 10,000 x g at 4°C. Between 80% and 90% of the total volume prior to centrifu-
gation can be expected to be available for toxicity testing and analysis.
To actually prepare the elutriate, homogenize sediments as described above. Place 300 mL of
dilution water (with a hardness or alkalinity matching that of the pore water) into a 2000 mL
graduated cylinder. Carefully deposit homogenized wet sediment into the graduated cylinder
until 300 mL of sediment has been added, using the amount of water displaced to measure the
volume of sediment added. Pour the contents of the graduated cylinder into a 2 L Erlenmeyer
flask (or other container suitable for the mixing apparatus used). Rinse the sides of the
graduated cylinder with a total of 900 mL of dilution water, and deposit the rinse water in the
Erlenmeyer flask. Cover the flask loosely with parafilm and place on a mixer table (or other
apparatus used for mixing). Mix the container at medium to high speed for 12 hours at 4°C
(Daniels et al. 1989). After the sediments and water have mixed for 12 h, carefully pour the
contents of the flask into centrifuge bottles, ensuring that the sediments do not settle to the
bottom of the flask. Centrifuge and decant in the same manner as for the pore water prepara-
tion (see above). Again, do not filter the sample.
rv-4
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IV.4 Toxicity Tests
The toxicity test methods described herein for pore water and elutriate can be used successfully
for the test species indicated; if other species are to be used for testing or TIE work, modifica-
tions may be necessary.
Organisms we routinely test in aqueous sediment fractions are C. dttbia, H. azteca, fathead
minnow, L. variegatus, and, on occasion, C. tentans. Methods for culturing these organisms are
available (Nelson et al. 1990; Phipps and Ankley 1990; U.S. EPA 1989c; U.S. EPA 1987). In
addition, most of these species are available through contract laboratories that specialize in
supplying organisms for toxicity testing.
Although there are slight differences in test conditions among the different species for aqueous
phase tests, the basic procedures are similar. For example, all initial toxicity tests are performed
in duplicate with five organisms per chamber at 25°C with a 16:8 L:D photoperiod. The
exposure volume for all species (with the exception of C. tentans) is 10 mL, and test results are
recorded at least every 24 h, along with appropriate water quality characteristics. In setting up
the aqueous phase tests, 30 mL polystyrene cups are used as the test chambers, and a stepwise
0.5 dilution series is utilized. Starting with the 50% concentration, place 10 mL of the
appropriate dilution water into each of the 50% concentration cups and each cup in the lower
dilutions. Exercise care to avoid excess sample aeration. Ten mL of pore water or elutriate
then is placed into the 100% and 50% concentration cups. Mix the dilution water with the pore
water/elutriate by drawing 10 mL of the solution into a pipette and reinjecting it back into the
IV-5
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cup, and repeating this procedure three times. Remove 10 of the 20 mL from the 50%
concentration cup and inject it into the next dilution, i.e., 25%, repeat the mixing procedure and
continue on to the next dilution. Ten mL should be discarded from the lowest pore wa-
ter/elutriate concentration to reduce volume to 10 mL. The final cup in the series will consist
solely of 10 mL of the dilution water (control). After setting up the dilution series, the
organisms are added randomly to each of two replicate cups per test concentration. Tests are
read daily, and organism survival and appropriate water quality parameters (e.g., pH, DO) are
recorded. Sample forms for recording toxicity test results are given elsewhere (U.S. EPA
1991a).
Because most pore water and elutriate samples tend to be highly colored, we suggest that older
(i.e., 24 to 48 h old) C. dubia be used for toxicity tests. Juvenile H. azteca (e.g., 7-14 day old)
and C. dubia used in the aqueous phase tests should be given an initial feeding of 67 uL of a
yeast-cerophyll-Trout Chow* (Ralston-Purina, Inc) solution (YCT) per 10 mL of test volume
(U.S. EPA 199Ib). Due to the substrate dependence of H. azteca and C. tentans, we use a
small (ca., 2.25 cm2) square of Nitex* screen (sand is more effective for C. tentans; R.A. Hoke,
AScI Corp., ERL-D, personal communication) to help prevent "floaters" and to increase control
survival in the aqueous tests. Extra care also should be given to observing or recording results
with test cups containing H. azteca or C. tentans. Do not place these cups directly upon a light
box as the extreme light intensity or heat may cause undue stress to the organisms. All aqueous
phase tests are conducted for 96 h with the exception of the cladoceran test (48 h).
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These general test conditions are consistent throughout the initial and baseline toxicity tests, as
well as the actual TIE sample manipulations, except where otherwise noted (e.g., the graduated
pH test).
rv-7
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V. Methods for Phase I Sediment TIE
Methods below describe in detail sample volumes and manipulations specific to toxicity tests
and TIE experiments with aqueous sediment test fractions (i.e., pore water, elutriate). Where
specific Phase I Phase H and Phase m TIE manipulations or procedures do not deviate from
those used for effluents, we do not describe them in detail. Therefore, in order to use this
guidance, it is essential that the analyst have and be familiar with effluent TIE manuals (U.S.
EPA 1988; 1989a; 1989b; 1991a).
V. 1 Initial Test
The purpose of these tests is to determine if the pore water or elutriate is toxic and, if so, how
toxic (i.e., to generate an LC3a), in order to identify appropriate concentrations for the TIE
manipulations. For the initial test, we generally test as many water column and benthic species
as are currently available in our cultures (up to 4). Forty mL of pore water or elutriate is
needed to test each species, and 60 or more organisms of the same age are required for each
species tested. A concentration series using 10 mL of 100%, 50%, 25%, 12.5%, and 6.25%
pore water should be prepared as described above. The concentration series also should have
duplicate control cups for each species tested. If after 24 h the LCM of the pore water is at or
below 6.25%, the test should be redone using a lower dilution series (e.g., add 3.13% and
1.56%).
V.2 Baseline Test
v-i
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If the initial toxicity test on the pore water or elutriate shows that it is acutely toxic (It., >50%
mortality at a 100% sample dilution), Phase I TIE manipulations can be initiated A baseline
toxicity test should be run each day that manipulations are performed on the sample. The
baseline test is needed as a reference point to determine whether a manipulation affected the
toxicity of the sample. The baseline also is needed to track the stability of sample toxicity
throughout the TIE.
The test concentrations of the baseline test are determined by the initial toxicity test Exposure
concentrations should be at 4x, 2x, Ix, and 0.5x the 24 h LQo of the initial test if the LC,0
was less than 25% whole sample concentration; if the LCM was greater than 25%. exposure
concentrations should be 100%, 50%, 25%, and 12.5%. The baseline test concentrations
(including the control) should be run in duplicate.
V.3 TIE Toxicity Tests and Volume Considerations
Toxicity tests conducted on samples that have undergone the TIE manipulations described below
differ in two important respects from either the initial or baseline toxicity tests. First, although
test conditions (e.g., volumes) are identical to those in the initial baseline tests, only a single
replicate (rather than duplicates) is used per concentration. Second, in conjunction with the TIE
manipulations, it is essential that there is strict adherence to the use of the appropriate
blanks/controb described elsewhere (U.S. EPA 1988; 1991a).
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The actual sample volume needed to conduct the TIE tests will depend upon the 24 h LC 0 from
the initial pore water or elutriate test All volumes listed are assuming that the initial LCJO was
greater than 25%. If the initial LC*, was less than 25%, a smaller sample volume would be
needed because of the necessity to dilute to 4x the LC^ for subsequent testing and TIE work.
Due to logistical concerns, e.g., the often limited supply of pore water, pore water should be
conserved throughout routine analyses and testing.
V.4 pH Adjustments
The pH adjustment tests remain identical to those for effluent TIE (U.S. EPA 1988; 1991a).
However, the volume of sample needed will be reduced if the pH adjustment/Cj, SPE manipula-
tion is not conducted. For example, due to sample volume limitations, we recommend that the
sediment TIE manipulations occur through a tiered approach (cf.. Section IDL4). If the Q, SPE
column removes sample toxicity at pH i, additional effort should be directed toward recovering
the pH i toxicity before performing pH adjustmem/Q, fracnonation. Besides the volumes
required for the EDTA and sodium thiosulfate tests, the first-tier tests (i.e., all manipulations
except the pH 3- and pH 9-Q, fractionanon) require 300 mL of sample at initial pH (pH i). and
100 mL sample volumes at pH 3 and pH 11, for a total volume of 500 mL. This volume
includes the amount needed to conduct the standing, aeration, filtration, and Clt SPE manipula-
tions and tests at pH i, and the three former manipulations and tests at the two altered pHs. A
total volume of 330 mL of dilution water is required for blanks and controls for the pH adjust-
ment tests, 130 mL at pH i and 100 mL at pH 3 and pH 9.
V-3
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The pH 3 and pH 11 adjustments are made by adding either 1.0, 0.1, at 0.01 N concentrations
of HO and NaOH to the pore water or elutriate sample. Follow the procedures and precautions
noted elsewhere (U.S. EPA 1988; 1991a). Table V-l gives the sodium chloride tolerances of H.
azteca and L. variegcuus, which may be used to determine whether the amount of NaCl resulting
from the acid/base adjustments is sufficient to cause toxicity. Comparable tolerance values for
C. dubia and fathead minnows are given elsewhere (U.S. EPA 1988; 199la).
Once the pH adjustments to the sample aliquots have been made, the analyst may proceed with
the aeration, filtration, and C, SPE manipulations. The 300 mL aliquots of pH adjusted
solutions should be divided, as described above, into the necessary volumes for the subsequent
manipulations.
The 30-mL aliquots of sample at pH 3, pH i, and pH 11 and corresponding pH-adjusted dilution
water blanks/controls are set aside for the pH-adjustment/standing test The pH 3 and pH 11
solutions are readjusted to pH i after the manipulations described below have been completed.
This is necessary to ensure that any effect on toxicity due to pH adjustment alone can be
evaluated relative to pH adjustment and subsequent manipulations.
V-4
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Table V-l. Species sensitivity to Phase I additives.
Additive
Species
LC»(g/L)
SRW1 MHRW2 VHRW3
EDTA Hyalella azteca
Lumbriculus variegatus
0.08
7.0
0.16
0.23
7.4
NajSA
MeOH
Nad
1 SRW,
2 X/TLTDX
Hyalella azteca 0.35
Lumbriculus variegatus 14
Hyalella azteca
Lumbriculus variegatus
Hyalella azteca
Lumbriculus variegatus 7.0
Soft reconstituted water
\I mA*4AW**«Kl«f I*a«v4 MfeAAMfl***'! 1*^*4
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V.5 Filtration
Conditioning of the filter apparatus and the 1.0 urn glass fiber filters remains identical to the
procedure used for effluent TIE work (U.S. EPA 1988; 1991a). Typically, we use a vacuum
apparatus for filtration; however, if vacuum filtration reduces sample toxiciry, then a pump
(positive pressure) filtration apparatus should be used until it has been determined that the
toxiciry removed was not caused by volatile compounds (e.g.,
The number of filters needed to filter a given aliquot of pore water or elutriate may be
considerably greater than the number needed to filter effluents. For example, due to clogging
caused by high particle content, it may be necessary to use as many as 10 filters to effectively
filter the 240 mL of pore water needed at pH i. Of course, if the tiered Phase I approach is
used (cf, Section V.4), sample volumes filtered at pH 3 and pH 11 (Le., 40 mL) will reduce
the number of filters required at these pH values. All filters may be prepared simultaneously
using a 100 mL distilled water rinse followed by a dilution water blank rinse at the appropri-
ate pH (U.S. EPA 1988; 1991a). Because of the common presence of "filterable" toxiciry in
sediment TIE work (e.g., Schubauer-Berigan and Ankley, 1991), we routinely save filters used
during Phase I for future toxicant recovery experiments. These are stored in appropriately-
labeled, sealed glass containers (i.e., one each for pH 3, pH i, and pH 11) and stored at 4°C
Once 240 mL of sample at pH i has been filtered, the filtrate is separated into one 40 mL
aliquot and one 200 mL aliquot The 40 mL will be used for the pH adjustment/filtration
V-6
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toxicity test, and the remaining 200 tnL will be used for the C,, SPE manipulation. Ail 40
mL of sample filtered at pH 3 or pH 11 is used for the pH-adjustment/filtration toxicity test.
V.6 Aeration
The pH adjustment/aeration tests and procedures remain identical to those described for
effluent Tffis (U.S. EPA 1988; 199la). Briefly, 30 mL of pH 3, pH i, and pH 11 sample and
their corresponding blanks are placed into separate 100 mL graduated cylinders and aerated
for 1 h. The rate of aeration should be maintained at 500 mL per minute. The pH of each
cylinder should be checked and readjusted with 0.01-1 N HQ or NaOH to the desired pH
midway through the aeration procedure. After 1 h of aeration, the sample should be removed
from the aeration vessel and transferred to a clean beaker using a siphon or pipette to prevent
any re-solution of sublated compounds into the sample.
V.7 Clg Solid Phase Extraction
With the exception of volume considerations (cf., Section in.4) and the elution process (cf.,
Section in.6), the methods for the C,, SPE test remain similar to those used for effluent TIE
work (U.S. EPA 1988; 1991a). Due to the relatively common occurrence of high (>5) log
K^, compounds in sediment pore water/elutriate samples, and the inability of methanol to
elute these nonpoiar compounds from Clg, a less polar solvent often must be used with
V-7
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sediment samples. We have found that methylene chloride is a good choice due to its ability
to reliably extract high (>7) log K^, compounds from the SPE column.
Each filter blank (described in the filtration step above) and its corresponding 200 mL pH
adjusted/filtered sample is passed over a properly conditioned and fraction- blanked 3 ml. SPE
column (U.S. EPA 1988; 1991a). Due to column limitations, the pH 11-filtered sample and
its corresponding filter blank must be adjusted to a pH of 9 before passage through the SPE
column. If there are limited volumes of pore water available, only pH i-filtered pore water
should initially be passed over the C,, SPE column (Tier I; cf., Section ffl.4). If toxicity is
not removed from the sample or recovered in the column eluate (described below), two 200
mL aliquots of the sample then should be pH adjusted (i.e., one aliquot pH 3, one aliquot pH
9), filtered and passed over a properly blanked and conditioned column. Another difference
in the Cts SPE procedure used for effluents is the blank and sample elution solvents. In
effluent work, the column is blanked and (after passage of the sample over the column) eluted
with three 100% methanol fractions in an attempt to recover toxicity from the column. For
sediment samples, the column is eluted with three 100% methylene chloride fractions after the
methanol elutions. Methylene chloride is miscible with methanol and has the additional
quality of being more volatik than methanol. These two properties allow a solvent transfer
of the methylene chloride fractions and blanks into methanol prior to toxicity testing. This is
essential because methylene chloride is more toxic to aquatic organisms than methanol (e.g.,
the former has a 48 h C. dubia LC50 of 0.46%, while that for methanol is 2.1%; NETAC
unpublished data). The transfer is accomplished by partially evaporating the fractions and
V-8
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blanks containing mcthylenc chloride with nitrogen (until the methylene chloride is removed)
and subsequently restoring the initial volume with methanol. (The solvent transfer process is
described in detail below.)
The 100% methanol fractions and their corresponding blanks then are tested as described
elsewhere (U.S. EPA 1988; 1991a). A 300 uL aliquot of the methanol test fraction is injected
into 20 mL of dilution water to give 3x the concentration of compounds in the original
samples. Dilutions of the 20 mL (3x) cup then are msufc The 3x concentration (containing
1.5% methanol) was selected to be below the methanol tolerance level of die organisms tested
(Table V-l; U.S. EPA 1988; 199la).
Before toxicity testing, the three methylene chloride fractions and corresponding blanks are
exchanged into methanoL With a Pasteur pipette, the methylene chloride sample fractions
and blanks are placed in appropriately-labeled 15 mL glass graduated centrifuge tubes. The
centrifuge tubes are used to accurately measure the amount of solvent in each fraction. To
ensure complete transfer of any contaminants, the scintillation vials used to capture the eluted
fractions from the Clg column should be rinsed with another 1 mL of methylene chloride and
this also should be transferred to the graduated centrifuge tube. Using a water bath to
maintain temperature at 20-25°C, the methylene chloride fractions then are evaporated to
approximately 500 uL using a gentle nitrogen stream (approx. 500 mL/min). This should be
done in a well ventilated hood Next, place a teflon coated, magnetic micro-stir bar into each
V-9
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test fraction and blank and add approximately S mL of high grade methanol to each centri-
fuge tube while stirring over a magnetic stir plate. Place the centrifuge tubes in the water
bath and sparge with nitrogen (500 mL/min) until a final volume of 500 uL is reached.
Repeat this methanol addition/rinsing process 3 times to ensure volatilization of all methylene
chloride from the fractions and blanks. After the third methanol rinse and subsequent
aeration process, the fraction and blanks are restored to their original 1 mL volumes with
methanol: using a 500 uL Hamilton syringe, measure and transfer each fraction into a clean
scintillation vial. Subtract the measured fraction volume (approximately 500 uL) from 1 mL
to obtain the quantity of methanol needed to bring the final volume to 1 mL. Rinse the
corresponding centrifuge tube with this volume of methanol and transfer to the scintillation
vial. Repeat this process for each fraction and its corresponding blank, making sure to
thoroughly clean the syringe with methanol between each transfer, by filling and emptying it
3 times with clean methanol. The fractions and blanks then are tested for toxicity using the
same procedures as for 100% methanol fractions (U.S. EPA 1988; 1991a).
To summarize the Phase I Cu procedure, 2 dilution series from each post C1S sample, (Le.,
one pH i sample for Tier 1), are tested, one dilution series corresponds to the first 100 mL of
sample passed through a C1§ column and one dilution series corresponds to the second 100
mL of sample passed through the C1S column (U.S. EPA 1988; 1991a). A dilution series for
each elution fraction also is tested (6 elution fractions from each pH). Therefore, if samples
are run at all three pH values, a total of 24 dilution series will be tested; 6 post column plus
V-10
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18 elurion fractions. In addition, the blanks are tested (only at the 3x concentration) for each
elun'on fraction.
V.8 Readjustment of Samples to pH i/Toxicity Testing
Upon completion of the pH adjustment/filtration, aeration and SPE manipulations, all
solutions (including the standing pH test) are returned to pH i of the sample before toxiciry
testing. This includes all pH-adjusted blanks which have undergone the same manipulations
as the pore water or elutriate samples. Again, care should be taken not to overshoot the
desired pH, and cause excessive changes in sample volume or ionic strength. After all
samples are returned to pH i, five organisms are added to each container, and toxiciry tests
conducted as described above. Note that, particularly in the samples subjected to pH
manipulations, the pH should be monitored and recorded carefully over the course of the
toxiciry test (U.S. EPA 199la).
V.9 EDTA Chelation Test
Decreasing concentrations of pore water/elutriate are used in conjunction with varying
additions of EDTA to help determine the degree of toxiciry associated with canonic metals in
the pore water or elutriate sample. Because the hardness of a water may affect the toxicity of
EDTA as well as its ability to chelate toxic canonic metals, the analyst must consider sample
hardness when setting test concentrations of EDTA. To aid in identification of appropriate
V-ll
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test concentrations, the toxicity of EDTA to the TIE test species must be ascertained for a
water with a hardness typical of that in the pore water or elutriate. Table V-l indicates these
values for H. azuca and L. varieganu; comparable data for C. dubia and fathead minnow and
options for the selection of appropriate test concentrations of EDTA are described elsewhere
(U.S. EPA 1988; 1991a). We generally recommend use of the "dilution" option described in
the latter document This particular option consists of setting up four dilution series of 100%,
50%, 25% and 12.5% whole pore water or elutriate (or 4x, 2x, Ix and 0.5x die LCy,). To
each of these dilution sets is added one of three decreasing quantities of die appropriate
EDTA concentration, thus forming a 3 x 4 matrix of EDTA level vs. pore water concentra-
tion. The three quantities of EDTA added should range from an amount approximating die
LCjQ of EDTA for the organism to a quantity that should not be toxic. Typical EDTA
concentrations used are Ix EDTA LCj,,, 0.5x EDTA LQ* and 0.25x EDTA LC». Because
many pore water or elutriate samples will have a hardness appreciably larger dian attainable
with standard reconstituted waters, Ix the EDTA LCM was chosen as the high addition level
to ensure adequate binding capacity of die EDTA (although recent studies at ERL-Duluth
indicate that EDTA chelates heavy metals on a 1:1 molar basis regardless of the sample
hardness; U.S. EPA 1991a; Schubauer-Berigan et al. 1992). A sample data sheet for the
dilution version of the EDTA test can be found in U.S. EPA (1991a). The baseline toxicity
test serves as the control for die EDTA test
V. 10 Sodium Thiosulfate Test
V-12
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We have successfully used sodium thiosuifate (NajSjO,) as both a reducing agent for toxic
oxidants and as a metal chelator in TIE work with pore water/elutriate samples. The Na^O,
toxicity test is procedurally similar to the EDTA addition test (U.S. EPA 1988; 1991a). For
general methods refer to the EDTA test and follow an identical format. As with the EDTA
test, we generally recommend the 3 x 4 matrix method, using Na^O, vs. pore water/elutriate
concentrations. The matrix method allows for better quantitation of sample toxicity relative
to EDTA/Na^Oj effectiveness, especially when mixtures of toxicants are present In our
experience, 0.2, 0.1, and 0.05 mL additions of a 20.5 g/L NajSjC^ stock solution to the 10
mL test volumes results in an acceptable range of concentrations needed to chelate or oxidize
most sample toxicants. These concentrations range from lethal or near lethal concentrations
of NajSjO, for a number of test species (at the 0.2 mL addition), to a concentration well
below toxic levels (Table V.I; U.S. EPA 1988; 1991a).
We have found that when using NajS2O3 for metal chelation, in some instances an additional
reductant may be needed. For example, if a sample contains both oxidants (they need not be
in toxic concentrations), and cationic metals, the Na^O, will preferentially reduce the
oxidants, and itself become oxidized. Therefore, because the reduced form of NajS2O3 is
needed to complex metals, it may not be an effective chelator in the presence of oxidants.
We have successfully dealt with this issue by using small quantities of SO2-saturated, distilled
water, to reduce excess oxidants before the addition of Na^O,. Therefore, the NajSjOs test
should be performed in duplicate, one with the SOj addition and one without the SO2
addition. (Note that an essential control in this test is the use of SO2 alone). To perform
V-13
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these tests, two aliquots of sample will be needed The volumes required for these tests will
be determined by the initial LCM of the sample, i.e., 100% vs 4x the LC^ Assuming the
initial LC^ of the sample is 25% or greater, two 80 tnL aliquots of sample will be needed to
complete the test. We have found that 10 uL of SO2-saturated, distilled water injected into
one of the 80 mL aliquots will generally reduce oxidants within a sample to the point that
NajSjOs will effectively chelatc metals. The SOj-saturated, distilled water is prepared by
gently bubbling SO2 into a beaker containing approximately 50 mL of distilled water for 15
minutes (U.S. EPA 199la). Extreme caution must be *a.ken when working with SO.: work
under a well ventilated hood while saturating the distilled water.
As an alternative to the use of SO2, we are currently exploring the use of sodium bisulfate as
a less-hazardous reductant that (theoretically) will not simultaneously chelate metals.
V. 11 Graduated pH Test
In our experience, the ambient pH of pore water/elutriate will drift up over the course of a
test to between 8.0 and 8.8. Therefore, only adjustments below pH 8.0 generally need to be
mad* We have used a number of methods for adjusting pH in aqueous samples. The
methods currently used to adjust sample pH are: (a) additions of HC1 and/or NaOH (closed
cup method), (b) additions of CQj to the test chamber headspace to control the bicarbonate
buffering system, or (c) the use of hydrogen ion buffers. Consult the latest version of the
Phase I manual for details on the use of these three methods (U.S. EPA 1991a).
V-14
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The particular pH control procedure ultimately used will be quite specific to sample and/or
test objectives; regardless of the choice "v^, the analyst should be aware of the strengths
and limitations of each approach.
V.I 1.1 Graduated pH test: Closed-cup method
In this approach, additions of HC1 are used in combination with the previously mentioned 30
mL polystyrene cups and lids (Coming* 35 mm/Tissue Culture Dish 35mm x 10 mm style).
In contrast to the various Phase I tests described above which use 10 mL sample volumes,
this particular version of the graduated pH test utilizes approximately 30 mL per test cup.
The use of the lids and larger sample volumes limits the degree of gas exchange between the
sample and the ambient air, thereby better maintaining pR Problems with this particular
approach can be inadequate maintenance of DO (particularly in tests with large organisms
such as fathead minnow), and/or the inability to monitor the pH of the sample throughout the
test The DO limitations arise due to the very nature of the pH control, (i.e., gas exchange
limitation); therefore it is imperative to measure the DO of the sample at test termination.
The inability to reseal the test chambers once opened precludes pH monitoring. Once the lids
are removed from the cups, the minimal headspace needed for pH control is extremely
difficult to maintain. This problem can be addressed by refrigerating extra sample dilutions at
the start of the test and adding pH-adjusted aliquots of these solutions as needed during the
test to reseal the test chambers.
V-15
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A 180 mL aliquot of sample is needed to perform the uoreplicated closed-cup graduated pH
test This volume requirement will be reduced if the initial LC» of the sample is less than
25%, in which case 180 mL of 4x the LCM concentration will be needed. The dilution series
procedure for the graduated pH test differs from previously mentioned tests due to the extra
volume of sample needed. The dilution series should be mfl4c, in triplicate (one for each of
the 3 pHs tested), using appropriately labeled glass beakers, as for the baseline pore water
test Each concentration in each replicate series should have a final volume of 30 mL. When
all dilutions have been made, label a dilution series for each of the three desired test pHs
(e.g., 6, 7, 8). Using 1.0, 0.1 and 0.01 N HC1, adjust each solution in the two lower dilution
series to the desired pH. Generally the samples at pH i will be at approximately 8; therefore,
this sample need not be pH adjusted. However, if the pH of an elutriate/pore water sample is
lower than 8, it can be adjusted up with stepwise additions of 1.0, 0.1, and 0.01 N NaOH
until the desired pH is achieved. Due to the high alkalinity of most pore water/elutriate
samples, the pH has a tendency to drift away from its adjustment point (e.g., pH 6, pH 7)
rather quickly. Therefore, pH control procedures (i.e., adding solution and sealing the cup)
should be initiated within 1 h of pH adjustment. After all of the pHs have been adjusted, the
solutions should be dispensed into 30 mL polystyrene cups. Randomly add 5 test organisms
into each cup and feed as necessary. Ensuring that all test organisms are well below the
surface of the solutions, place the lid on the solution surface and gently press down until a
seal is formed between the cup, solution and lid. Care must be taken not only to eliminate
any air trapped between the lid and pH adjusted solutions, but also to not crack or rupture the
test cup. The cups it the high pH value (i.e., > 8) do not require a lid to maintain their pH,
V-16
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in fact, we have found thai placing a lid on the higher pH solution tends to reduce pH over
the course of the test When reading and recording the closed-cup graduated pH test certain
precautions must be observed. Due to the inability to reseal the test cups once opened,
chemistry measurements (pH, DO) are taken only from test cups with complete mortality,
unless, as mentioned previously, test solution renewals are performed. Chemical analyses
from all test cups should be recorded at test termination to ensure that pH and DO have been
maintained at appropriate levels throughout the test
V.I 1.2 Graduated pH Test: COj Method
To control sample pH with CO,, flush the headspace of a sample in a gas tight container with
a measured amount of COy'air. The major limitation of this approach is the potential toxicity
caused by the amount of CO2 needed to adjust the samples, given the high alkalinity values
commonly observed in sediment pore waters and/or elutriates.
To perform the graduated pH test using CO2, gas tight containers large enough to hold at least
two 30 mL polystyrene cups must be obtained. We have successfully used 1 L Nalgene*
screw-top jars (available from most supply houses), 1 L latch-top gasket-sealed canning jars,
or have constructed our own containers out of glass (cf., Section HL5.1.1; U.S. EPA 1991a).
The amount of COj needed to adjust the pH of the samples, and dilutions of the samples, will
depend upon alkalinity. Therefore, unless sample and dilution water alkalinities are the same,
V-17
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each dilution of the sample may require a different amount of CO2 to maintain the desired
test pH.
Perform six 10-mL dilution series with the test sample as described earlier. Place duplicate
cups at each concentration into each of 3 separate COj test containers for a total of 3 repEcate
dilution series. (Note that testing of several concentrations within the same controlled-
headspace chamber should be avoided, because of observed contamination of test cups with
volatile compounds present in the higher concentration test cups.) For the CO2 method, we
use duplicate rather than single replicates at each concentration, primarily because duplication
increases the likelihood of obtaining at least one test cup at each concentration with the
appropriate pH. As opposed to the closed-cup method, relatively little work or sample
volume is involved in adding the duplicate at each test concentration. After appropriately
labeling chambers for the concentration and desired pH, use a 1 L gas syringe to flush 8-10%
CCyair into the pH 6 dilution series containers, stopper and allow to equilibrate (a test
adjustment run initially without test organisms should establish the effective CO2 spiking
concentration, which will be sample-specific). This procedure is repeated with the pH 7
dilution series using a 2-5% CCyair concentration. Because in most samples ambient pH will
drift to above 8, no CO2 should be injected into the pH 8 containers, nor should the chambers
for this series (pH 8) be sealed. After 2 h of equilibration, the pH of the samples should be
taken. The pH invariably will be slightly different from the target pH, but should be within
0.5 pH units of the desired pH, If the desired pH (6-6.5, 7-7.5, 8 and above) is not achieved,
different volumes of CO2 will need to be used. This can be ascertained only through trial and
V-18
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error. If after equilibration the desired pHs are maintained, the test organisms can be
introduced to the individual cups. If possible, the test should be read (and pH recorded) twice
daily. The additional test readings are needed to better monitor and maintain pH. At each
reading, the CO2 chambers must be reflushed with the predetermined appropriate volume of
CO2. If the CO2 adjustments alone are unsuccessful in maintaining the desired pH, it may be
necessary to first adjust the pH of the sample using HC1 additions.
V.I 1.3 Graduated pH Test: Buffer Method
We also have successfully used minimally toxic hydrogen ion buffers (Mes-pH 6, Mops-pH 7,
Popso-pH 8) to maintain sample pH in the graduated pH test (cf.. Section HL5.1.1). Major
advantages of this approach include excellent pH control, low maintenance (e.g., no need for
continual CO2 flushing), adequate DO, rapid test set up time, and small sample volume
requirements. The limitation of this approach is the possibility that the buffers might interfere
with sample toxicants. Another problem encountered with the buffers is the inability of the
Mops buffer to maintain consistent pH across all sample dilutions. Therefore, before using
the buffers for pH control in extended TEE work with a sample, a comparison toxicity test
using both the buffers and the closed-cup or CO2 flushing method should be conducted to
ensure that sample toxicity is not altered by the buffers, and that the buffers will maintain
constant pH at all sample dilutions.
V-19
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Because of possible buffer toxicity artifacts or interactions with sample contaminants and/or
toxicity, the lowest molar concentration of buffer that will maintain the desired pH must be
determined. Generally 10-25 mM concentrations of buffer will maintain a pore water or
elutriate pH while being non-toxic to the test organism (Table EH-5; U.S. EPA 1991a).
Initially a 100% sample with a corresponding dilution water blank should be tested at several
buffer concentrations, e.g., 10, 15, 20 and 25 mM. We have found that the lowest buffer
concentration that maintains pH in the 100% sample also will generally maintain pH in the
sample dilutions, and therefore should be used for subsequent tests. Because there is not a
headspace limitation with use of the buffers, 10 mL sample volumes can be used. The
buffers must be added to sample and dilution water separately. The weight (in g), of buffer
needed to attain the desired molar concentration of buffer is calculated by multiplying the
volume of sample (in L) by the formula weight (FW) of the buffer (195.2 g/mol for Mes,
209.3 g/mol for Mops, 362.4 g/mol for Popso) by the molar concentration of buffer desired,
(e.g., volume x FW x M). Generally, additions of the crystalline Mes and Mops buffers to
the pore water/elutriate or dilution water will adjust the sample to the desired pH (i.e., pH 6
and pH 7, respectively). Using a magnetic stir plate and stir bar, stir the buffers into the
sample and dilution water. If the buffer addition fails to adjust the sample to the correct pH,
the desired pH can be initially achieved with the use of 1.0, 0.1, 0.01 N HQ or NaOR
Thereafter, an appropriate buffer concentration will maintain pH. In our experience, the pH 8
buffer (Popso) generally is not needed because of the inherent drift of pore water and
elutriates to pHs of 8 or above; additionally, the Popso buffer requires large quantities of
NaOH to adjust the samples and dilution waters to pH 8 after addition of the buffer. Such
V-20
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individual metal TUs were summed for each extraction method. Since TIE had previously
shown that these metals were the sole acute toxicants at this site, a "non-availability ratio"
(NR) could be calculated for each extraction technique by dividing the total potential metal
TUs by the actual sample toxicity. A ratio close to 1 would indicate that the metals in the
sample, independent of their sample concentration, were actually all bioavailable, based on
dilution water LCjoS for the metal. The higher the ratio, the less likely it is that the metals
were totally available. Of all the extraction methods, dialysis gave the NR closest to unity
(0.75), followed by high-speed centrifugation (2.8) and syringe extraction (3.3). The sediment
press and low speed centrifugation procedures resulted in higher NR values (6.2 and 7.3,
respectively), suggesting that these methods extracted relatively high concentrations of
unavailable metals. In a study comparing various pore water extraction methods for metals,
Carignan et al. (1985) also found that centrifugation at low speeds (5000 rpm) recovered
higher concentrations of copper, zinc, and organic carbon than either centrifugation at higher
speeds (10000 rpm) or in situ dialysis.
LQ.2.3 Recommended Pore Water Preparation Method
Based on sample volume considerations for TIE work, as well as results of the studies above
and reported by others (e.g., Capel 1986; Schults et al. 1991), we recommend that pore water
be isolated via centrifugation without subsequent filtration. Although the specific mechanism
is not known through which filtration removes toxicants from pore water samples (e.g.,
removal of contaminants associated with particles, filtration of oxidized metal-ligand
m-i2
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complexes, sorption to the filter, etc.), data from our laboratory clearly indicate that any pore
water isolation technique that requires or incorporates filtration as part of the extraction
process is likely to remove bioavailable metals and nonpolar organics. Our data also suggest
that speeds ranging from 2,500xg to lO.OOOxg are suitable for pore water preparation. The
lower speeds, however, may result in the presence of unavailable metals in pore water. The
speed of centrifugation has been shown in other research not to affect the partitioning of
nonpolar organics, such as PCBs, into pore water (Capel 1986). Finally, to reduce artifacts
induced by temperature fluctuations (Bischoff et al. 1970), we recommend that pore water
samples be prepared under cool (ca., 4°Q conditions. This can be achieved either through
the use of a refrigerated centrifuge, or through sample preparation in a controlled temperature
room (e.g., walk-in cooler).
In our pore water characterization studies, there are several factors which we did not address
(for example, the effects of oxidation on speciation of pore water nutrients and contaminants).
Further research is required to extend existing knowledge of pore water's suitability for
evaluating sediment toxicity.
III.3 Use of Benthic Species for Aqueous Testing
Another facet differentiating sediment TIE from effluent TIE involves the selection of species
for testing. Sediments contain epibenthic and benthic species and communities quite different
from the pelagic species used in effluent toxicity and TIE studies. Several common benthic
m-13
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large amounts of NaOH may consequently increase the conductivity of the sample and/or
dilution water and thereby cause arnfactual toxicity. After making pH adjustments with the
buffers, organisms can be added to the samples and tests performed using normal protocols.
As with any version of the graduated pH test, pH should be closely monitored both during
and at the termination of the test
V-21
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VI. Methods for Phase n Sediment TIE
Phase n TIE procedures were developed to identify specific toxicants within the different
classes of compounds characterized in Phase L The Phase n procedures for sediments are
quite similar in approach, yet differ in several important instances, from those developed for
complex effluents (U.S. EPA 1989a). The methods typically involve steps to separate and/or
concentrate the toxicants from the nontoxic sample components, and at present are largely
limited to metals, nonpolar organic compounds and ammonia. Phase n procedures as they
have been applied specifically to sediment pore waters are described here. Because we have
encountered toxicants in sediments not commonly seen in effluents (e.g., H,S, filter-associated
toxiciry), we have provided additional sections where appropriate. For sediments, we have
found that certain Phase I TIE manipulations (e.g., filtration) may greatly reduce toxiciry
without providing specific evidence for any particular class of compounds as the suspect
toxicants. Therefore, we address possible approaches for assessing the components of toxiciry
removed by filtration, a relatively nonspecific sample manipulation. In addition, advice is
provided concerning the use of volatile toxicant transfer techniques useful for working with
hydrogen sulfide. Finally, some guidance is presented concerning the use of multiple
manipulations in Phase n. Not all of the procedures recommended below will be applicable
for every sample; they are merely suggestions of techniques that have been successful for the
sediments with which we have worked. As with effluent TIE, the interpretations by the
researcher will be of paramount importance to the success of the TIE.
VI-1
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VI. 1 Filter-Removable Toxicants: Metals and Nonpolar Organic Compounds
The toxicity of sediment pore waters often is reduced by as much as 40%-90% upon
filtration. We have observed the filtration effect with pore water from sediments of all types,
ranging from oily, highly organic sediments to very sandy sediments. These losses of toxicity
have been attributed to both metals and nonpolar organic compounds. We address first the
possible approaches for identifying the nonpolar toxicants retained by filters, and second,
techniques for recovering metals from filters.
VI. 1.1 Nonpolar Organic Compounds: General Overview
The Phase n procedure for effluents suggests using C,, SPE to separate nonpolar toxicants
from a filtered sample, and recommends a subsequent compound recovery scheme in which
the column is eluted with methanol/water fractions. This procedure has not been particularly
successful in pore water TIEs, due to a combination of several factors. First, as mentioned
previously, filtration frequently removes a large component of the pore water toxicity.
Additionally, the elution schemes developed for recovering nonpolar effluent toxicants from
the SPE column have not been successful for the types of organic toxicants often encountered
in sediments (Le., those with a high log K,,). Finally, sediments may be contaminated with a
large number of similar types of compounds (e.g., PAHs in oily samples), making separation,
concentration and identification steps more difficult and protracted than for effluents.
VI-2
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One method of identifying nonpolar organic compounds removed by filtration is to recover
toxicants from the filters by solvent extraction. Methylene chloride has proven a useful
extraction solvent because it is less polar and more volatile than (yet nriscible with) methanol.
Methylene chloride/methanol extracts then can be solvent-exchanged into water and passed
over a Ct, column. A subsequent solvent elution series (e.g., methylene chloride/methanol)
then may be used to recover toxicants from the column and to isolate the compounds causing
toxiciry.
The second approach to identifying "filterable" nonpolar organics is to circumvent the
filtration step prior to C,, fracdonation. Filtration is required for samples containing high
suspended solid concentrations (e.g., most elutriates and pore waters) to prevent column
plugging, as well as to ensure that nonpolar compounds on particles or in solution actually
will sorb to the resin, rather than being removed by physical filtration by the resin matrix.
We have avoided filtration of pore water samples altogether by centrifuging at a speed
somewhat higher than that used for initial isolation of the pore water (e.g., 20,000-
30,000 x g). This approach may remove sufficient paniculate concentrations from the sample
to avoid plugging the Cu SPE column; however, the column should be observed during
passage of the sample through the SPE packing to determine whether physical filtration rather
than column loading appears to be taking place. Post-Q, samples should be collected and
tested at several points in the fractionation process to assure that column overloading and
subsequent toxicity breakthrough has not occurred.
Vl-3
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In the general Phase n procedure, separation of non-polar compounds into fractions is
achieved by isolating the nonpolar compounds from aqueous sample with Qi SPE columns
and then eluting the columns with a graded, increasingly non-polar sequence of eight metha-
nol/water solutions. The last, most non-polar solution recommended for these elutions is
100% mcthanol (U.S. EPA 1989a). By this treatment, non-polar compounds are separated
into eight fractions according to their polarity, the most polar eluting in the earliest fractions
and the least polar eluting in the last fractions. Sediment pore waters, however, frequently
contain many potentially toxic nonpolar compounds, some of which are so non-polar that they
are not efficiently eluted by this water/methanol scheme. The non-polar character of these
compounds is reflected by their high (i.e., >5) log K,, values. The 100% methanol fraction is
too polar to elute such compounds from the SPE column. Consequently, a modified fraction-
ation procedure has been developed in which methylene chloride has been incorporated into
the water/methanol elution scheme.
In the modified procedure, the 100% methanol fraction is replaced by 50% methanol/50%
methylene chloride, and three additional fractions have been added, all three consisting of
100% methylene chloride. This modified method generates a total of eleven fractions.
Because methylene chloride is very toxic to aquatic animals, fractions containing methylene
chloride must be exchanged into methanol prior to testing. In addition, compounds eluting in
the methylene chloride fractions do not elute sharply in any one fraction, but rather tend to
spread out over several fractions. Therefore, to ensure that the toxicity of such compounds
will not be diluted out prior to toxicity testing, it is prudent to combine the methylene
VI-4
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chloride fractions, eliminate and/or exchange the methylene chloride into methanol, reduce the
volume down to that of an individual fraction and treat the combined fractions as one single
fraction. If further separation of the high log K^, compounds contained in this combined
fraction is required, the best course of action would be to further fractionate the toxic
fractions using HPLC.
VI. 1.1.1 Nonpolar Organic Compounds: Filter Extraction
For Phase n evaluations, a large aliquot of pore water should be filtered through a number of
1 um glass fiber filters, prepared and blanked as described in the Phase I TIE document (U.S.
EPA 199la). The volume of pore water required for the Phase n Clt fractionation depends
on the amount of toxicity in the samples and the number of nonpolar organic compounds
contributing to toxicity, among other variables (U.S. EPA 1989a). One to two L of pore
water/elutriate will usually be sufficient for aqueous sediment fractions containing 1-4 TU.
For very oily samples, filters may require changing after the passage of as little as 30 mL:
changing them more frequently than absolutely necessary will reduce required filtration time
considerably. The total volume filtered and the number of filters required should be recorded.
Filtered (and unfiltered) sample aliquots should be tested for toxicity.
If filtration removed toxicity in Phase I, then attempts should be made to extract the toxi-
cant(s) from the filters, first with a nonpolar solvent (e.g. methylene chloride/methanol) and
subsequently with a dilution water adjusted to pH 3. For the methylene chloride filter
Vl-5
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extractions, filters (1-10 per beaker) should be soaked in a small beaker containing 20 mL
methylene chloride for I h (if available, the beakers should be placed in cold-water sonication
baths). The filters should be removed from the beakers using forceps and set aside to dry in
a hood (see Section VI. 1.2). If more than 10 filters were extracted, then the solutions from
two or more beakers should be combined at this point. The methylene chloride solution
should then be solvent-exchanged, first to methanol and then to dilution water, by evaporating
half the methylene chloride solution volume (e.g., 10 mL) in a hood and replacing it with an
equal volume of 100% methanol. The sides of the beaker should be carefully rinsed with the
resulting mixture. Next, the methylene chloride/methanol mixture should be evaporated to
half its original volume (e.g., 10 mL), and replaced with 100% methanol, carefully rinsing the
sides of the beaker with the resulting solution. The resulting methanol solution should be
evaporated to half the original volume, and dilution water added to restore the volume. At
this point it may be necessary to vigorously mix the solution using a vortex mixer or tissue
homogenizing probe in order to dissolve the extracted compounds in the methanol/water
solution. The solution should once again be evaporated to half-volume, restored to volume
with dilution water, and mixed thoroughly with a vortex mixer or tissue homogenizer. The
solution should then be diluted to a volume (e.g., 250 mL if 1 L was the original sample
volume) that is 1/4 the original filtered sample volume, and tested for toxicity at test
concentrations of 4x, 2x, Ix and 0.5x the original pore water or elutriate concentration.
Blank filters should be treated identically to sample filters and tested likewise for toxicity.
The remaining solution, if toxic, (approximately 200 mL, equivalent to 800 mL of original
VT-6
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pore water) may be passed over a Ctl column and fractionated as described below (See
Section VI. 1.1.3).
VI. 1.1.2 Nonpolar Organic Compounds: High-Speed Centrifugation
As mentioned previously, an alternative to filter extraction for pre-Ci8 column treatment of
pore water is to avoid filtration entirely by centrifuging the sample at higher speed in order to
remove the bulk of the sample participates. We use an DEC* B-22 high speed centrifuge to
prepare 1200 mL of bulk sediment or a Beckman* L5-50 ultracentrifuge to prepare 200 mL
of bulk sediment sample per run at 20,000 x g 4°C for 30 min. The supernatant is then
decanted, collected and stored in glass beakers prior to testing. Unless a large capacity
centrifuge unit is employed this method will be quite time consuming if 2 L are required for
Clg fractionation. The sample should then be passed over a C,3 SPE column as described
below (Section VL1.1.3), and toxicity tests performed with the supernatant, a post-Cu sample
and a baseline pore water sample.
VI. 1.1.3 Clg SPE Fractionation
Either filtrate (if Phase I procedures determined that filtration did not affect toxicity), a
solvent-exchanged filter extract solution, or the supernatant from a sample centrifuged at
20,000-30,000 x g (see Sections VI. 1.1.2, VI. 1.1.3) may be fractionated using the Clg SPE
column.
VI-7
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The procedure for SPE fractionation of sediment pore water is very similar to the general
Phase II Ct| SPE fractionation procedure (U.S. EPA 1989a). The single major modification is
in the composition of the solvent mixtures used to elute the Cu column. Instead of the
original two-solvent elution scheme, using water and methanol, the modified procedure also
uses a third solvent, methylene chloride.
As in the original procedure, we recommend that one 6 mL high capacity Clg column be used
for every 1000 mL of sample fractionated. The column is preconditioned by pumping 25 mL
of 100% methanol through the column, followed by 25 mL of high purity distilled water. At
this point 25 mL of dilution water is passed over the column, the last 10 mL of which is
collected for a column blank toxicity test Special care must be taken not to allow the
column to go to dryness at any time during the conditioning procedure. One solvent must be
added after another in such a way that precludes air passing through the column.
After the column is conditioned, the elution blanks are collected (Table VI-1). Three mL (in
two 1.5 mL aliquots) of fractions 1-11 (i.e., 25% CHjOH/r^O to 100% CHjClj) is passed
over the column and each fraction collected in separate analytically clean labeled vials.
Each eluting solution is allowed to pass completely through the column before the next
solution is added to the column. After the elution blanks have been collected the column
VI-8
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Table VI-1. Composition of 11 recommended solvents for eluting the Ctl column in Phase
sediment TIE.
COMPOSITION OF ELUTING SOLUTIONS (% BY VOLUME)
METHYLENE
FRACTION WATER METHANOL CHLORIDE
1
2
3
4
5
6
7
8
9
10
11
75%
50%
25%
20%
15%
10%
5%
0%
0%
0%
0%
25%
50%
75%
80%
85%
90%
95%
50%
0%
0%
0%
0%
0%
0%
0%
0%
0%
0%
50%
100%
100%
100%
VI-9
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should be reconditioned with methanol and water as described above; again it is very
important not to allow the column to go to dryness during this procedure.
The column is now ready for the extraction of non-polar compounds from the sample. A
1000 mL sample aliquot is pumped through the column at a rate of 5 mLymin. Three 20 mL
samples of the post-SPE column effluent are collected after 25 mL, 500 mL and 950 ml,, of
the sample passes through the column. These aliquots can be tested to monitor for the
breakthrough of toxicity in the post C,, sample (U.S. EPA 1988; 1991a). The column is
allowed to go to dryness at this stage.
The column loaded with sample is now ready for elution. The column is eluted exactly as
described for the collection of elution blanks. If more than 1000 mL of sample is being
fractionated, and therefore more than one column is being used, then the complete fraction-
ation procedure from preconditioning, collection of the elution blanks, reconditioning and
column elution is repeated for each column. Corresponding fractions from several columns
may be combined at this stage, but dilution water blanks should be kept separate. The vials
containing the fractions are sealed with perfluorocarbon or foil-lined caps and stored under
refrigeration.
The methylene chloride must be eliminated from fractions before toxicity testing of the
fractions can take place. As discussed earlier, because concentrations of the high log
toxicants may be diluted over several fractions, it is best to combine those fractions that
VI-10
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contain methylene chloride (fractions 8 through 11), eliminate methylene chloride from this
one combined fraction and reduce the volume to 3 mL, as would be the case if a toxicant had
cluted in only a single fraction (i.e., in an ideal situation).
Eliminating methylene chloride from methanol or exchanging it into methanol is relatively
easy because methylene chloride is more volatile than methanol and can therefore be removed
from a mixture of these two solvents by evaporation under a stream of nitrogen. The solvent
exchange approach described below is essentially the same as that described above in Section
V.7. We have found that this step is readily accomplished by combining Fractions 8 through
11 (Table VI-1) for a total of 12 mL in a 50 mL glass centrifuge rube. To this is added
another 12 mL of methanol and a perfluorocarbon coated magnetic micro stir bar. The
centrifuge tube is placed in a water bath at 50°C and stirred magnetically with a stream of
nitrogen gently flowing over the surface of the solution. After the volume of the solution is
reduced to 3 mL, the sides of the rube are carefully rinsed with 3 additional mL of methanol
and the volume is again reduced by evaporation to 3 mL. These repeated evaporations and
additions of methanol ensure that the methylene chloride is eliminated from the fraction. If
only a single fraction containing methylene chloride is to be toxicity tested, then exchange
into methanol is achieved by using the above procedure. In this case, however, 3 mL of
methanol are added to the fraction, the volume is reduced to 3 mL and sides of rube rinsed
with another 3 mL methanol followed by a final volume reduction to 3 mL. Any procedure
that involves combining fractions and/or exchanging methylene chloride into methanol also
VI-11
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must be carried out with the corresponding elution blanks. In that way if any artifactual
toxicity is inadvertently introduced by these procedures it will be detected in the blank
Toxicity testing of SPE fractions is carried out as described in the general Phase n method
(U.S. EPA 1989a). When concentrating toxic fractions for GC-MS analysis or further HPLC
fractionation, SPE fractions 1 through 7 are treated in the same way described in the general
methods. However, because fractions 8 through 11 contain methylene chloride, they do not
require SPE concentration to remove water, as do the earlier fractions. Simply exchange the
methylene chloride fractions directly into methanol for GC/MS analysis.
Further fractionation of SPE fractions, including fraction(s) 8-11 (combined or individually),
by HPLC can be carried out exactly as described in the general method (U.S. EPA 1989a).
Compounds that require methylene chloride for elution from Clg SPE columns often can be
fractionated by HPLC using a water/methanol solvent gradient. We have found that high log
K,w compounds elute in the 100% methanol portion of such an HPLC gradient and therefore
would be found in the 100% methanol HPLC fractions. The 100% fractions, therefore, can
be concentrated for GC-MS analysis by evaporative volume reduction, as described above.
At this point, however, we have been unable to determine whether further useful separations
of high log K^ compounds can be achieved by reverse-phase HPLC fractionation.
GC-MS analysis of concentrated SPE and HPLC fractions is carried out in the same manner
as described elsewhere (U.S. EPA 1989a).
Vl-12
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VI. 1.2 Metals: General Overview
In addition to nonpolar organics, bioavailable metals frequently are removed by filtering
during Phase I evaluations (cf., Section m.3). If a reduction in toxicity is observed after
filtration yet little is known initially concerning which (or how many) types of compounds are
causing toxicity in the sample, attempts should be ma/fe to extract both nonpolar organics and
metals from the filters. After it has been determined that only one class of compounds is
being retained by the filters, repeated sequential solvent extractions of the filters are not
necessary. Therefore, this section is intended to reflect the initial extraction sequence that
might be attempted in Phase n, when it is presumably unknown exactly which types of
compounds are being retained by the filters.
VI. 1.2.1 Filter Extraction
Following the methylene chloride/methanol extraction described above, filters should be
extracted with a dilution water sample adjusted to pH 3 in an attempt to recover cationic
metals removed by filtration. Filters (as many as were required in the filtration step) should
be soaked in a beaker containing a volume of dilution water, adjusted to pH 3 with HC1,
sufficient to concentrate extractions by 4x their original concentration in the sample (i.e., 250
mL if the original sample volume was 1 L). Again, if possible, the filters should be sonicated
in a cold-water bath for 1 h after which filters may be discarded. The dilution water
solutions should be readjusted to pH i, and subsequently tested for toxicity at 4x, 2x, Ix and
VI-13
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0.5x whole pore water or elutriate concentrations. As noted above, blank filters should be
treated identically to the sample filters or elutriate LQo, and subsequent blank solutions tested
for toxicity.
If the pH 3 dilution water samples exhibit toxicity, then EDTA or sodium thiosulfate may be
added to determine whether toxicity was caused by canonic metals, and metals scans can be
performed on the samples for possible identifications (U.S. EPA 1989a).
VI.2 Use of Multiple Manipulations in Phase n
As mentioned previously (cf., Section IIL5), performing two (or sometimes more) manipula-
tions simultaneously is an extremely useful tool when several toxicants are present in a
sample, particularly when a mixture of metals and ammonia is present For example, we
have been able to discriminate between the effects of lead, zinc, copper, and ammonia using
the results of the graduated pH, sodium thiosulfate and EDTA tests (Schubauer-Berigan et ai.
1990). Such distinctions are possible because these compounds behave uniquely when
exposed to combinations of these tests. For example, ammonia and copper elicit different
responses in the graduated pH, EDTA and sodium thiosulfate tests. Ammonia and lead give
different responses in the graduated pH and EDTA tests, and ammonia and zinc elicit
different responses only in the EDTA test.
VI-14
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Another approach that has worked well when several classes of compounds are causing
sample toxicity is to perform various Phase I manipulations at an altered pH. For example,
one sediment with which we have worked contained toxicity due to ammonia and a nonpolar
organic. Toxicity was greatest at pH 8.5, filtration did not affect toxicity, and passage over a
Clg column at pH i appeared to reduce toxicity. By using the Mes hydrogen ion buffer to
control pH at 6.2, we were able to determine the nature of the Clf removable contaminant
without the interference of ammonia toxicity.
Simultaneous use of pH control and EDTA or sodium thiosulfate tests may successfully
discriminate between toxicity due to ammonia and that due to metals showing increased
toxicity at lowered pHs (e.g., lead, copper); however, this is not likely to be successful for
samples containing a combination of zinc (or nickel) and ammonia toxicity, because, like
ammonia, these compounds are also more toxic at higher pH (Table m.4). In such cases, we
have found that toxicity due to the metals sometimes can be removed by pH 11 filtration and
recovered from the filters using pH 3 dilution water extractions (Schubauer-Berigan et al.
1992; U.S. EPA 1991a). A recovery of toxicity and metals from the filters is evidence that
metals are causing at least some of the observed sample toxicity.
VI.3 Volatile Toxicants: Hydrogen Sulfide
Toxicity caused by hydrogen sulfide may be prevalent in some sediments. Phase I evidence
for this is loss of toxicity during aeration, and enhanced toxicity at pH 6 in the graduated pH
VI-15
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test Also, fish species (such as fathead minnows) tend to be more sensitive to hydrogen
sulfide than invertebrates, such as cladocerans (Broderius et al. 1977 and unpublished NET AC
data). If such observations are made, then the volatile toxicant transfer experiment is
warranted. The revised Phase I manual (U.S. EPA 199la) gives methodological details of
this procedure. To test for hydrogen sulfide, the sample should be purged at pH 3 and
volatilcs recovered in a dilution water adjusted to pH 9. Both purged sample and dilution
water should be subsequendy tested for toxicity at pH 6. Hydrogen sulfide should be
measured using a colorimetric method (APHA 1980) in both test solutions if toxicity is
recovered in the trap water. See Section m.5.4.1 for certain caveats regarding the analytical
detection limit for hydrogen sulfide as it relates to the toxicity of H^S to fathead minnows.
Methods are available (e.g., steam distillation) for detecting H,S at concentrations smaller
than those detected by the colorimetric technique (Broderius and Smith 1977).
VI-16
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VH. Methods for Phase m Sediment TIE
Phase m confirmation should be completed, if possible, when conducting a sediment TEE.
Given the possibility for the sample manipulations in Phases I and II to create artifacts with
respect to toxicity, failure to complete Phase m could be potentially disastrous, particularly if
decisions concerning remediation will be made based on TIE results.
As is true for effluent TIEs, multiple confirmation methods should be used in sediment TEE to
provide a "weight of the evidence" conclusion that the correct toxicant(s) have been identi-
fied. Most of the procedures used for Phase HI confirmation in sediment TIE work are quite
similar to those used for effluent TIEs (U.S. EPA 1989b). These procedures are described
below with specific considerations and examples concerning their adaptation to sediments.
VH.l Correlation
One of the most powerful Phase HI procedures for TIE work with complex effluents is the
correlation of toxicity due to the suspect toxicant(s) with observed sample toxicity. For this
approach to be successful, there must be a range of samples with sufficiently different
toxicities to develop a meaningful relationship. With effluents this is achieved by sampling
over time; this also would work with sediments in which the suspect compound varies over
time, e.g., in the case of seasonal agricultural runoff of relatively nonpersistent pesticides
(Chandler and Scott 1991). This approach also would be feasible in situations were the
vn-i
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suspect toxicant(s) are nricrobially-generated compounds, such as ammonia or hydrogen
sulfide, whose concentrations vary seasonally. However, with the majority of toxic sedi-
ments, the best strategy for maximizing variability will not be by sampling over time. Rather,
it generally would be more feasible to collect sediment samples exhibiting a gradient of
contamination with the suspect toxicant(s) from the study site of interest This gradient
theoretically may be generated using either horizontal or vertical (depth) variations in the
sediments. To date, however, we have only used the former approach. Note that if the TIE
is focused on a relatively well-defined and specific area (e.g., the outfall of a particular
effluent) the samples for the correlation approach are best derived from within that area,
rather than from an area far removed from the site. This type of approach helps to minimize
potentially confusing test results that may be related to site-specific differences in the intrinsic
physical/chemical characteristics of sediments. This sampling design is not appropriate,
however, if the question that needs to be answered is: "Is the suspect toxicant(s) from a
particular source?"
One potential complication with the correlation approach for sediments can be the identifica-
tion of a gradient of contamination by only the suspect toxicant(s). For example, if the
suspect toxicant(s) appears to emanate from a point-source discharge, and if the study location
is impacted by only one discharger, concentrations of all contaminants in the sediments, not
just the suspect toxicant, likely will decrease upon moving away from the input Another
potential problem with the correlation approach for sediment TIE is that it may be very
difficult, in relatively complex situations with multiple point and/or nonpoint source inputs
vn-2
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(e.g., in some Great Lakes harbors), to identify a gradient of sediment toxicity that corre-
sponds only to a particular suspect toxicant(s). Therefore, to use the correlation approach in
sediment TIE, the researcher should be aware of the discharge history and hydrodynamics of
the study system.
The actual application of the correlation approach to pore water or elutriate samples (once
they are collected) is essentially the same as described for effluents; several excellent
examples are described elsewhere (U.S. EPA 1989b; Amato et al. 1992). One example of the
use of the correlation method in a TIE with pore water samples was conducted using a set of
samples from the lower Fox River/Green Bay (Ankley et al. 1990a). The samples were
collected from 13 sites within the system, and the characterization and identification portions
of the TIE were completed and implicated ammonia as the suspect toxicant Toxicity of the
samples to C. dubia and fathead minnows (expressed as TUs) was then plotted against the
pore water ammonia concentrations (Fig. VII-1). In both instances there was a strong,
statistically-significant correlation (r > 0.9, p< 0.01), providing confirmatory evidence for the
role of ammonia as a sample toxicant [This particular correlation did not express the suspect
toxicant concentration on the x-axis in TUs as recommended by the U.S. EPA (1989b); this
was necessitated by the fact that we did not have an LCjo for ammonia at the pH of the pore
water samples.]
For some types of toxicants the correlation approach may be of limited value. This is
particularly true in the case of metals where, in some instances, there is considerable
vn-3
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Figure VTI-1. Correlation of concentrations of ammonia in sediment pore waters from the
lower Fox River/Green Bay watershed with toxicity of the samples to fathead
minnows and C. dubia, Toxicity is expressed as TU for (a) 96-h fathead
minnow mortality, and (b) 48-h C. dubia mortality. When no mortality was
observed, a value of zero toxic units was assigned. In the two instances in the
C. dubia test in which less than 50% mortality occurred at a pore water
concentration of 100\5, a value of 0.5 TU was assigned (from Ankley et al
1990a).
6
(a)
0.92
O • • • •
O
X
O
(b)
0.96
o . • • •
20
40
6O
AMMONIA (mg/L)
vn-4
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uncertainty as to how to define that fraction of the total concentration that is biologically
available (e.g., see Schubauer-Berigan et al. 1992).
Vn.2 Species Sensitivity
Evaluation of the relative sensitivity of two (or more) test species to the same elutriate or
pore water sample can be an extremely useful confirmation tool. The main prerequisite for
using the species sensitivity approach as a confirmation tool is, of course, the ability to
identify test species with a range of sensitivities to the suspect toxicant(s). Differences in
sensitivity among test organisms should be determined in single chemical tests performed
under similar conditions as for the elutriate or pore water. Generalizations as to the relative
sensitivity of different species cannot easily be made without actually performing the
appropriate tests. For example, although oligochaetes have traditionally been considered to be
relatively insensitive to toxic compounds, we have found L. variegarus to be among the most
sensitive of our test species to ammonia (Table V-l).
The use of relative species sensitivity as a confirmation test with the lower Fox River/Green
Bay pore water samples mentioned above has been demonstrated (Ankley et al. 1990a; Table
VII-1). Fathead minnows were the most sensitive test organisms, followed by C. dubia and
then a bacterial species (Photobacterium phosphoreum) both to the pore water samples and to
the suspect toxicant, ammonia, providing confirmatory evidence for its role in sample toxicity.
A second example of the use of relative species sensitivity is from a TIE on sediment pore
vn-5
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Table VIM. Comparison of the sensitivities of C. dubia, fathead minnow, and Photobac-
terium phosphoreum to pore water to Green Bay/Fox River sediment pore
water (from Ankley et al. 1990a). The LC*, and ECy, values are expressed in
percent pore water. The 95% confidence intervals are indicated in parentheses.
Site
1
2
3
4
5
6
7
8
9
10
11
12
13
1 Concentration
2 \ir- D.liaKU ,
Fathead minnow
96-h LC«
40.6(34.1-48.4)
30.9 (25.3-37.8)
35.4 (NC)2
35.4 (NC)
18.0 (NC)
21.8 (17.8-26.6)
35.4 (NC)
37.9 (30.4-47.2)
17.4 (NC)
21.1 (16.8-26.5)
NMJ
NM
NM
resulting in 20%
"i"infirl»n/*» limifc
C. dubia
48-h LC«
>100
63.0(51.0-77.8)
>100
84.1 (NC)
56.1 (43.0-73.3)
39.7(32.1-49.0)
84.1 (NC)
75.8 (NC)
44.5(34.1-58.2)
39.7 (32.1-49.0)
NM
NM
NM
inhibition of light production.
/vuilH nnt h* falfiilamH Hn^ frt laclc
P. phosphoreum
15-min ECJ
>100
>100
>100
MOO
MOO
MOO
MOO
MOO
MOO
MOO
MOO
MOO
MOO
r»f narrial mnrtalitv.
NM; No mortality.
vn-6
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water from Turkey Creek, MO (Table vn-2). In this case, the suspect toxicant in the
sediment sample was zinc; the sample was most toxic to C. dubia, followed by H. azteca and
finally fathead minnow. This trend closely paralleled the sensitivity of the three species to
zinc in single chemical tests (Table EQ-4), thereby lending support to the identification of zinc
as responsible for sample toxicity.
vn.3 iMass Balance
As for effluents, the mass balance approach can be useful for sediment elutriate or pore water
samples from which toxicity can be removed and subsequently recovered. The primary
method for doing this with effluents has been via the SPE column, and a specific example of
this is provided in the U.S. EPA (1989b) Phase in document With sediments, we also have
used the mass balance approach in cases where filtration removed some portion of toxicity
which subsequently could be recovered by either solvent (methanol, methylene chloride;
Schubauer-Berigan and Ankley 1991) or pH 3 dilution water extraction, respectively, of
nonpolar organics and metals (Schubauer-Berigan et al. 1990).
When using the mass balance method, with either effluents or pore water, there are a number
of factors that must be considered (U.S. EPA 1989b). One of the most important of these is
that the approach will not be particularly useful in instances where the matrix of the sample
has a great effect on bioavailability of the toxicant of concern. This is because the suspect
toxicant is removed from a relatively complex sample matrix containing a variety of natural
vn-7
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Table VII-2. Comparison of the sensitivies of C. dubia, fathead minnows and H. azteca to
sediment pore water from five sites along Turkey Creek, Joplin, MO. LCSOs
are expressed in percent pore water. Test lengths were 48 h for C. dubia and
96 h for fathead minnows and H. azteca. The 95% confidence intervals are
indicated in parentheses.
Site
1
2
32
42
5
C, dubia
35 (NC1)
<3 (NC)
>100
>100
3 (NQ
Percent LC^
Fathead ™nnow
>100
47 (34-63)
89 (NC)
71 (NO
77 (NC)
H. azteca
>100
4.5 (NC)
84 (47-100)
>100(NO
13 (10-18)
1 NC, confidence intervals not calculable due to lack of partial mortality in test concentra-
tions.
2 Toxicity at these sites was determined to be due to ammonia.
vn-8
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ligands (e.g., components of dissolved organic carbon), capable of binding and reducing the
bioavailability of contaminants, and tested in "clean" dilution water. Thus, if bioavailability
of the suspect toxicant is mediated via binding to ligands, much more toxicity may be
recovered than was removed, causing the mass balance approach to be essentially useless.
This matrix effect might possibly be avoided by spiking the SPE fractions or filter extracts
back into the sample from which the toxicants have been removed. We have reasonable
success using this approach with pore water samples from which filtration removed toxicity;
however, at least with effluents, this particular variation on the mass balance approach has not
been especially useful with post-SPE samples due to "artifactual" toxicity, corresponding to
the growth of methylotrophic bacteria, that often is observed. In the case of sediment pore
water, such artifactual toxicity seems somewhat less common, so it may be possible to
successfully spike methanol fractions into some post-SPE aqueous sediment samples as a
confirmation step. However, we have relatively little experience with this approach.
VH.4 Deletion Approach
Generally, the deletion approach will not be feasible in the confirmation phase of a sediment
TIE. The one instance where this particular technique might be useful is if the suspect
toxicant were a relatively labile, nonpersistent compound present in the sediments due to a
point or nonpoint source discharge which could be controlled in some manner. If this were
possible, it would be a powerful confirmation tool.
VH-9
-------
vn.5 Symptoms
The comparison of "symptoms" or behavior of organisms exposed to the test sample to their
behavior in single chemical tests with the suspect toxicant was proposed as a useful confirma-
tion technique for effluent TIE work (U.S. EPA 1989b), and this approach should be of equal
usefulness in sediment TEE. In particular, time to mortality is a quantifiable symptom that
can be monitored relatively easily in toxiciry tests.
Vn.6 Spiking
Spiking sediment pore water or elutriate samples with suspect toxicants can be an extremely
powerful confirmation tool if the analyst is aware of the limitations in the approach. A
number of different approaches can be employed when using spiking as a confirmation tool.
One approach is to double (or increase by some other multiple) the concentration of the
suspect toxicant in the sample and subsequently determine sample toxiciry; if the correct
toxicant was identified, there should be a corresponding doubling in toxiciry (U.S. EPA
19895). Another use of spiking in the confirmation phase is to spike the suspect toxicant into
a sample with a similar matrix as tne test sample (e.g., a non-toxic pore water from a nearby
site). If the suspect toxicant was correctly identified, spiking similar concentrations as those
observed in the test sample should result in similar toxiciry. In some instances, a useful
confirmation technique is spiking a sample from which the suspect toxicant has been removed
(e.g., via aeration, SPE, zeolite, etc.) with similar concentrations of the suspect toxicant. One
vn-io
-------
of the major problems with this last approach is that most manipulations used to remove
suspect toxicants also can significantly alter the matrix of the test sample (e.g., pH, ion
composition, organic carbon content, etc.), thereby potentially altering the toxicity of the
spiked toxicant Also, there exists the potential for manipulations to remove multiple classes
of toxicants (e.g., SPE will remove nonpolar organics and metals; zeolite will remove
ammonia and metals; etc.), so even if there is a good match between original and manipulat-
ed/spiked sample toxicity this is not, by itself, a definitive confirmation result
There are a number of factors to consider when using spiking as a confirmation tool For
example, spiking may not work well for compounds such as metals, where there is little or no
understanding of the speciation reactions that may occur in complex solutions. Because
different forms of metals have different toxicities, toxicity of the spiked metal may differ
from that of the metal in the original sample. This would make it appear that the identifica-
tion of the suspect toxicant was wrong when it was in fact correct (e.g., Schubauer-Berigan et
ai. 1992). A similar type of artifact theoretically also could occur when spiking chemicals
with a great affinity to dissolved or paniculate organic carbon (e.g., compounds with
relatively high K^J. Binding to organic carbon can reduce bioavailability (and toxicity) of
many nonpolar organic chemicals; however, this process may require a significant equilibra-
tion time. Thus, a "fresh" nonpolar organic spiked into a sample may appear more toxic than
a comparable amount of the same nonpolar organic already in the test sample (Ankley and
Burkhard 1992). This would make it appear that the incorrect toxicant had been identified,
when in fact there had been a correct identification.
vn-n
-------
A final issue that must be considered when using spiking as a confirmation tool is one that is
more or less pervasive throughout Phase m of the TIE. In our experience, when multiple
toxicants are present in a sample, they frequently behave in a somewhat independent fashion,
i.e., their toxicity is not additive. Thus when testing a sample in a dilution series, the
compound with the greatest number of TUs in the sample will "drive" sample toxicity, and
there may be no evidence that another toxicant is present. Yet, when the suspect toxicant at
the highest concentration is removed from the sample, the sample will still be toxic (albeit at
a higher sample concentration) due to the second, independently-acting toxicant This
situation is exacerbated by the fact that most (if not all) Phase I and Phase n sample
manipulations are relatively non-specific. For example, SPE can remove toxic concentrations
of both metals and nonpolar organics from the same sample; however, if the nonpolar organic
is driving sample toxicity, and it is acting independently from the metal, the nonpolar is likely
to be the only toxic compound identified. Therefore, although a correct identification may
have been made on one of the sample toxicants, not all toxicity was accounted for. This can
be a potentially serious problem if the goal of remediation is to remove all toxicity. The
scenario of independently-acting toxicants is particularly troublesome when using spiking as a
confirmation tool, because spiking lends no insight as to whether this may be occurring in a
sample.
VH.7 Matrix Changes
vn-i2
-------
Changing the sample matrix in a manner designed to alter the toxicity of specific compounds
can be a very useful confirmation technique. One factor that is routinely altered in the TIE
confirmation step with pore water and/or elutriate samples is pH. Due to the common
occurrence of pH-dependent toxicants such as ammonia, hydrogen sulfide or metals in
sediments, the graduated pH test can be an invaluable tool. In order to use alterations in pH
as a confirmation technique, it is essential that the behavior of the suspect compounds has
been well defined at the various test pHs. A positive result in the test (i.e., sample toxicity
behaves as predicted) can be a powerful piece of evidence for the confirmation. If there is
any deviation from expected behavior, over time or among samples, this can help provide
evidence that either the wrong toxicants, or not all toxicants, were identified. Some caution
should be taken, however, when extrapolating the effects of pH on toxicants tested in clean
laboratory water to the potential effects of pH on suspect toxicants in a complex matrix such
as pore water or elutriate. The pH-dependent behavior of a toxicant in one matrix may not
exactly mirror behavior observed in a very different matrix.
VH.8 Summary
The importance of a complete confirmation for TEE work cannot be overly emphasized. The
tools described above are not all-inclusive of the approaches that could be used for confirma-
tion; however, we have had success with each of the approaches described. Depending upon
the situation, other techniques may be useful for providing confirmatory evidence, and these
should be used For example, theoretically it is possible to subject bulk sediment samples to
vn-i3
-------
some types of TIE manipulations (e.g., EDTA additions, pH changes) to help confirm specific
compounds as sample toxicants; however, due to our lack of experience with these types of
manipulations, more definitive guidance cannot be presented.
One final comment concerning confirmation should be m*4t If a particular sample does not
appear to behave in an expected manner in Phase EQ, a Phase I (and Phase H) analysis should
be performed as soon as possible on the same sample. This often will result in relatively
quick explanation for the unexpected Phase En observations. In fact, in some instances a
complete or partial Phase I analysis may be desired of every sample used for either Phase Et
or Phase HI; the Phase I results ultimately could prove to be a powerful part of the confirma-
tion (e.g., see Ankley and Burkhard 1992).
VH-14
-------
vm. Literature Cited
Adams, W.J., R.A. Kimerle, and R.G. Mosher. 1985. Aquatic safety assessment of chemi-
cals sorbed to sediments, in: R.D. Cardwell, R. Purdy and R.C. Banner, eds., Aquatic
Toxicology and Hazard Assessment: Seventh Symposium. American Society for Testing and
Materials, Philadelphia, PA, pp. 429-453.
Amato, J.R., D.I. Mount, EJ. Durhan, M.T. Lukasewycz, G.T. Ankley, and ED. Robert.
1992. An example of the identification of diazinon as a primary toxicant in an effluent
Environ. Toxicol. Chem. (In Press.)
Ankley, G.T. and L.P. Burkhard. 1992. Identification of surfactants as toxicants in a primary
effluent Environ. Toxicol. Chem. (Manuscript Submitted.)
Ankley, G.T., A. Katko, and J.W. Arthur. 1990a. Identification of ammonia as an important
sediment-associated toxicant in the lower Fox River and Green Bay, Wisconsin. Environ.
Toxicol. Chem. 9:313-322.
Ankley, G.T., M.T. Lukasewycz, G.S. Peterson, and D.A. Jenson. 1990b. Behavior of
surfactants in toxicity identification evaluations. Chemosphere 21:3-12.
vm-i
-------
Ankley, G.T., GJ-. Phipps, E.N. Leonard, D.A. Benoit, V.R. Mattson, P.A. Kosian, A.M.
Cotter, J.R. Dierkes, DJ. Hansen, and JD. Mabony. 199la. Acid volatile sulfide as a factor
mediating cadmium and nickel bioavailability in contaminated sediments. Environ. Toxicol.
Chem. 10:1299-1307.
Ankley, G.T., M.K. Schubauer-Berigan, and R.A. Hoke. 1991b. Use of toxicity identifica-
tion evaluation techniques to identify dredged material disposal options: A proposed
approach. Environ. Management. (In Press.)
Ankley, G.T., M.K. Schubauer-Berigan, and J.R. Dierkes. 1991c. Predicting the toxicity of
bulk sediments to aquatic organisms with aqueous test fractions: pore water versus elutriate.
Environ. Toxicol. Chem. 10:1359-1366.
Ankley, G.T., V.R. Mattson, E.N. Leonard, Jl. Bennett, and C.W. West 1992. Predicting
the bioavailability of copper in freshwater sediments: evaluation of the role of acid volatile
sulfide. Aquat. Toxicol. (Manuscript Submitted.)
American Public Health Association (APHA). 1980. Standard Methods for the Examination
of Water and Wastewater. 15th Edition. APHA, American Water Works Association Water
Pollution Control Federation, Washington, D.C.
vra-2
-------
Batley, G.E., and MS. Giles. 1980. A solvent displacement technique for the separation of
sediment interstitial waters. In R.A. Baker, ed., C1 nntamjjnants and Sediments. Vol n. Ann
Arbor Sci., Ann Arbor, MI, pp. 101-118.
Benes, P., and E. Steinnes. 1974. In situ dialysis for the determination of the state of trace
elements in natural waters. Water Res. 8:947-953.
Bischoff, J.L., R.E. Greer, and A.O. Luistro. 1970. Composition of interstitial waters of
marine sediments: temperature of squeezing effect Science 167:1245-1246.
Broderius, S.J., and LI*. Smith, Jr. 1977. Direct determination and calculation of aqueous
hydrogen sulfide. Anal. Chem. 49:424-428.
Broderius, S.J., Li. Smith, Jr., and D.T. Lind. 1977. Relative toxicity of free cyanide and
dissolved sulfide forms to the fathead minnow (Pimephales promelas). J. Fish. Res. Board
Can. 341:2323-2332.
Burkhard, LJ>. and Ankley, G.T. 1989. Identifying toxicants: NETAC's toxicity-based
approach. Environ. Sci. Technol. 23:1438-1443.
vm-3
-------
Burkhard, LJP., EJ. Durban, and M.T. Lukascwycz. 1991. Identification of nonpolar
toxicants in effluent using toxicity-based fracnonanon with gas chromatography/mass
spectrometry. AnaL Chem. 63:277-283.
Bunon, G.A., B.L. Stemmer, K.L. Winks, P.E. Ross, and L.C. Burnett 1989. A multitrophic
level evaluation of sediment toxicity in Waukegan and Indiana Harbors. Environ. Toxicol.
Chem. 8:1057-1066.
Campbell, P.G.C. and P.M. Stokes. 1985. Acidification and toxicity of metals to aquatic
biota. Can. J. Fish. Aq. Sci 42:2034-2049.
Capel, PD. 1986. Chlorinated hydrocarbons in the porewater of lake sediments. Chapter 1
in Distributions and Diagenesis of Chlorinated Hydrocarbons in S<*U"ig"« PhD. disserta-
tion. University of Minnesota, St. Paul, MN
Cahgnan, R., F. Rapin, and A. Tessier. 1985. Sediment porewater sampling for metal
analysis: a comparison of techniques. Geochim. et Cosmochim. Acta 49:2493-2497.
Chandler, G.T. and G.L Scott 1991. Effects of sediment-bound endosulfan on survival,
reproduction and larval settlement of meiobenthic polychaetes and copepods. Environ.
Toxicol. Chem. 10:375-382.
vm-4
-------
Chapman, P.M., and R. Fink 1984. Effects of Puget Sound sediments and their elutriates on
the life cycle of Capitella capiuaa. Bull. Environ. Contain. Toxicol. 33:451-459.
Connell, D.W., M. Bowman, and D.W. Hawker. 1988. Bioconcentration of chlorinated
hydrocarbons from sediment by oligochaetes. Ecotoxicol. Environ. Saf. 16:293-302.
Daniels, S.A., M. Munawar, and C.I. Mayfield. 1989. An improved elutriation technique for
the bioassessment of sediment contaminants. Hydrobiologia. 188/189:619-631.
DiToro, D.M., J.D. Mahony, D.J. Hansen, KJ. Scott, M.B. Hicks, S.M. Mays, and M.S.
Redmond. 1990. Toxicity of cadmium in sediments: the role of acid volatile sulfide.
Environ. Toxicol. Chem. 7:483-498.
Doe, K.G., W.R. Ernst, W.R. Parker, G.R.J. Julien, and P.A. Hennigar. 1988. Influence of
pH on the acute lethality of fenitrothion, 2,4-D and aminocarb and some pH-altered sublethal
effects of aminocarb on rainbow trout (Salmo gairdnerf). Can. J. Fish. Aq. Sci. 45:287-293.
Durhan, EJ., M.T. Lukasewycz, and J.R. Amato. 1990. Extraction and concentration of
nonpolar organk toxicants from effluents using solid phase extraction. Environ. Toxicol.
Chem. 9:463-466.
vm-5
-------
Eadie, B J., P.P. Landrum, and W, Faust 1982. Polycyciic aromatic hydrocarbons in
sediments, pore water and the amphipod Pontoporeia hoyi from Lake Michigan. Chemo-
sphere 11:847-858.
Ferguson, W.J., K.I. Braunschweiger, W.R. Braunschweiger, J.R. Smith, J.J. McConnick, C.C,
Wasmann, NJ. j irvis, D.R BeU, and N£. Good. 1980. Hydrogen ion buffers for biological
research. Anal. Biochem. 104:300-310.
Giesy, J.P., RJL Graney, J.L. Newsted, C.J, Rosiu, A. Benda, E.G. Kreis, and FJ. Horvath.
1989. Comparison of three sediment bioassay methods using Detroit River sediments.
Environ. Toxicol. Chem. 7:483-498.
Giesy, J.P., and R.A. Hoke. 1989. Freshwater sediment toxicicy bioassessment: Rationale
for species selection and test design. J. Great Lakes Res. 15:539-569.
Hesslein, R.H. 1976. An in situ sampler for close interval pore water studies. Limnol.
Oceanogr. 21:912-914.
Hockett, J.R. and D.R. Mount. 1990. Use of metal chelating agents to differentiate among
sources of toxicity. Eleventh Annual Meeting of the Society of Environmental Toxicology
and Chemistry, Abstract, p. 162.
vm-6
-------
Junk, G.A. and J.J. Richard. 1988. Organics in water solid phase extraction on a small
scale. Anal. Chem. 60:451-454.
Knezovich, JJ5., F.L. Henderson, and R.G. Wilhelm. 1987. The bioavailability of sediment-
sorbed organic chemicals: a review. Water Air Soil Pollut 32:233-245.
Kuehl, D.W., G.T. Ankley, L.P. Burkhard, and D.A. Jensen. 1990. Bioassay directed
characterization of the acute toxicity of a creosote leachate. Hazardous Waste Hazardous
Mater. 7:283-291.
Long, E.R., M.F. Buchman, S.M. Bay, R.J, Breteler, R.S. Carr, P.M. Chapman, J.E. Hose,
A.L. Lissner, J. Scott, and D.A. Wolfe. 1990. Comparative evaluation of five toxicity tests
with sediments from San Francisco Bay and Tomales Bay, California. Environ. Toxicol.
Chem. 9:1193-1214.
Nebeker, A.V., and C.E. Miller. 1988. Use of the amphipod crustacean Hyalella azteca for
freshwater and estuarine sediment toxicity tests. Environ. Toxicol. Chem. 7:1027-1033.
Neilson, A.H., A.-S. Allard, S. Fischer, M. Malmberg, and T. Viktor. 1990. Incorporation of
a subacute test with zebra fish into a hierarchical system for evaluating the effect of toxicants
in the aquatic environment. Ecotoxicol. Environ. Safety 20:82-97.
vm-7
-------
Nelson, M.K., C.G. Ingersoll, and FJ. Dwyer. 1990. New standard guide for conducting
solid-phase sediment toxicity tests with freshwater invertebrates. ASTM Draft Document
El383. American Society for Testing and Materials, Philadelphia, PA.
Norberg-King, T.J., EJ. Durhan, G.T, Ankley and E. Robert 1991. Application of toxicity
identification evaluation procedures to the ambient waters of the Colusa Basin Drain.
Environ. Tox. and Chem. 10:891-901.
Phipps, G.P. and G.T. Ankley. 1990. Test methods to estimate the acute and chronic toxicity
and bioaccumulation of sediment-associated contaminants using the aquatic oligochaete,
Lumbriculus variegams. ERL-D Report No. 7896A. U.S. Environmental Protection Agency,
Duluth, MN.
Schubauer-Berigan, M.K., J.R. Dierkes, and G.T. Ankley. 1990. Toxicity identification
evaluation of contaminated sediments in Buffalo River, NY and Saginaw River, ML National
Effluent Toxicity Assessment Center Technical Report 20-90. 107 pp.
Schubauer-Berigan, M.K. and G.T. Ankley. 1991. The contribution of ammonia, metals and
nonpolar organic compounds to the toxicity of sediments interstitial water from an Illinois
River tributary. Environ. Toxicol. Chem. 10:925-939.
vm-8
-------
Schubauer-Berigan, M.K., J.R. Amato, G.T. Ankley, S.E. Baker, L.P. Burkhard, J.R. Dierkes,
J.J. Jenson, M.T. Lukasewycz, and TJ. Norberg-King. 1992. The behavior and identification
of toxic metals in complex mixtures: examples from effluent and sediment pore water
toxicity identification evaluations. (In Preparation.)
Schults, D.W., L.M. Smith, S.P. Femro, F.A. Roberts, and C.K. Poindexter. 1991. A
comparison of methods for measuring trace organic compounds and metals in interstitial
water. Water Res. (In Press.)
Spehar, R.L., and J.T. FiandL 1986. Acute and chronic effects of water quality criteria-based
metal mixtures on three aquatic species. Environ. Toxicol. Chem, 5:917-931.
Stumm, W. and J.J. Morgan. 1981. Aquatic chemistry - an introduction emphasizing
chemical equilibria in natural waters. John Wiley & Sons, New York, NY. 583 pp.
Swartz, R.C., W.A. DeBen, K.A. Sercu, and J.O. Lamberson. 1982. Sediment toxicity and
the distribution of amphipods in Commencement Bay, Washington, USA. Mar. Pollut Bull.
13:359-364.
Swartz, R.C., G.R. Ditsworth, D.W. Schults, and J.O. Lamberson. 1985. Sediment toxicity to
a marine infaunal amphipod: cadmium and its interaction with sewage sludge. Mar. Environ.
Res. 18:133-153.
vm-9
-------
Swartz, R.C., P.F. Kemp, D.W. Schults, G.R. Ditsworth, and R.J. Ozretich. 1989. Acute
toxicity of sediments from Eagle Harbor, Washington, to the infaunal amphipod Rhepoxynius
abronius. Environ. Toxicol. Chem. 8:215-222.
Swartz, R.C. D.W. Schults, T.H. DeWitt, G.R. Ditswotth, and J.O. Lambenon. 1990.
Toxicity of fluoranthene in sediment to marine amphipods: a test of the equilibrium
partitioning approach to sediment quality criteria. Environ. ToxicoL Chem. 9:1071-1080.
U.S. Army Corps of Engineers/Environmental Protection Agency Committee on criteria for
Dredged Material. 1977. Ecological evaluation of proposed discharge of dredged material
into open waters: implementation manual for Section 103 of Public Law 92-532. Environ-
mental Effects Laboratory, U.S. Army Engineer Waterways Experiment Station, Vicksburg,
MS.
U.S. Environmental Protection Agency. 1985. Ambient water quality criteria for ammonia.
EPA-440/5085-001. Environmental Protection Agency, Environmental Research Laboratory-
Duluth, Duluth, MN, and the Criteria and Standards Division, Washington, D.C.
U.S. Environmental Protection Agency. 1987. Guidelines for the culture of fathead minnows
(Pimephales promelas) for use in toxicity tests. EPA/600/3-87/001. Environmental Research
Laboratory-Duluth, MN.
vm-io
-------
U.S. Environmental Protection Agency. 1988. Methods for aquatic toxiciry identification
evaluations: Phase I toxicity characterization procedures. EPA-6XXV3-88-034. Environmental
Research Laboratory-Duluth, MN.
U.S. Environmental Protection Agency. 1989a. Methods for aquatic toxicity identification
evaluations: Phase n toxicity identification procedures. EPA-600/3-88-035. Environmental
Research Laboratory-Duluth, MN.
U.S. Environmental Protection Agency. 1989b. Methods for aquatic toxicity identification
evaluations: Phase in toxicity confirmation procedures. EPA-600/3-88-036. Environmental
Research Laboratory-Duluth, MN.
U.S. Environmental Protection Agency. 1989c. Short-term methods for estimating the
chronic toxicity of effluents and receiving waters to freshwater organisms, Second Edition.
EPA/600/4-89/001. Environmental Monitoring and Support Laboratory, Cincinnati, OH.
U.S. Environmental Protection Agency. 1991a. Methods for aquatic toxicity identification
evaluations: Phase I toxicity characterization procedures. Second Edition. EPA-600/6-
91/003. Environmental Research Laboratory-Duluth, MN.
Vffl-11
-------
U.S. Environmental Protection Agency. 1991b. Toxicity Identification Evaluation: Charac-
terization of Chronically Toxic Effluents, Phase I. EPA/600/6-91/005. Environmental
Research Laboratory-Duluth, MN.
Wells, M.J.M and J.L. Michael. 1987. Reversed-phase solid-phase extraction for aqueous
environmental sample preparation in herbicide residue analysis. J. Chromatogr. Sci. 25:345-
350.
vm-12
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