Freshwater Bivalve Ecotoxoicology - Chapter 5 ppsx

5
Laboratory Toxicity Testing with
Freshwater Mussels
Christopher G. Ingersoll, Nicola J. Kernaghan, Timothy
S. Gross, Cristi D. Bishop, Ning Wang, and AndyRoberts
INTRODUCTION
Numerous laboratory toxicity studies have been conducted with freshwater mussels in an attempt to
understand the role of contaminants in the decline of field populations of mussels(Chapter 7). In
these studies, early life stages of musselsofseveral species were highly sensitive to somemetals
and ammoniainwater exposures when comparedtomany of the most sensitive species of other
invertebrates, fish, or amphibians that are commonly used to establish U.S. Environmental Protec-
tion Agency (USEPA)Water QualityCriteria(WQC) (Augspurger et al.2003; USGS 2005a,
2005b). Importantly, results of thesestudies indicate WQC for individual chemicals established
for the protectionofaquatic organisms may not be adequately protectiveofsensitive stages of
freshwater mussels. This chapter provides asummary of methods from over 75 laboratory toxicity
studies conducted with freshwater mussels and alsoprovides an overview of astandardized method
for conducting water-only acute and chronic laboratory toxicity tests with glochidia and juvenile
freshwater mussels (ASTM 2006a). Three life stages (glochidia,juveniles, and adults) have been
used to conductlaboratory toxicity tests with mussels. Withinthis chapter,toxicity studies are
separated according to the medium of exposure (aqueous, sediment, and host fish). Each section
begins with areview of the methodsused to conduct toxicity tests (e.g., obtaining organisms,
duration of exposure, exposure chambers,and toxicity endpoints). Each section also discusses
issues that have been identified regarding the routine application of the methods (e.g., to generate
data for the derivationofWQC)and discussesresearchneeds.The final sectionofthischapter
reviews the use of the Asian clam ( Corbicula fluminea)asasurrogate for assessing effects on native
unionids. Finally, asummary of future research needs for improving methods used to conductacute
and chronic toxicity tests with freshwater mussels is provided.
AQUATIC TOXICITY TESTING WITH GLOCHIDIA, JUVENILE, AND ADULT
LIFE STAGES OF FRESHWATER MUSSELS
M ETHODS FOR C ONDUCTING A CUTE W ATER-ONLY T OXICITY T ESTS WITH G LOCHIDIA
OF F RESHWATER M USSELS

ReviewofMethods
Conditions that have been used to conductacute toxicity tests with glochidia of freshwater mussels
are summarized in Table 5.1including the testconditions recommendedinASTM (2006a). The
procedures outlined in Table 5.1 are consistent with acutetoxicity testing methods for fish, macro-
invertebrates, and amphibians (ASTM 2006c)and with acute toxicity testing methods for saltwater
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© 2007 by the Society of Environmental Toxicology and Chemistry (SETAC)
TABLE 5.1
SummaryofTest ConditionsUsed to Conduct Toxicity Tests with Glochidia of Freshwater Mussels
Conditions
Johnson et al.
(1990, 1993) Lasee (1991)
Huebner and
Pynnonen
(1992)
a
Goudreau,
Neves, and
Sheehan(1993)
Jacobson et al.
(1997)
Keller and
Ruessler (1997) McCann (1993)
Klaine, Warren,
and Summers
(1997)
USGS (Unpub-
lished Data)
Recommended Test Con-

ditions in ASTM (2006a)
1Species tested Utterbackia
imbecillis
b
Lampsilis
cardium
c
Anodonta
cygnea,
Anodonta
anatina
Villosairis Multiple species
d
Multiple species
e
V. iris U. imbecillis Multiple species
f
NA
g
2Test type Static Static Static Renewal Static Static Static Static Static, renewal,
flow-through
Static, renewal, or flow-through
(depending on chemical
tested)
3Test duration
(hours)
24 48 24, 48, 72, 144 24 24, 48 4, 24, 48 24 24, 48 6, 24, 48 6, 24 (up to 48 depending on
viability of glochidia)
4Temperature ( 8 C) 20 21 13 22 10–25 25 20 25 20 20
5Light quality Ambient lab light NR

h
NR NR NR NR NR Ambient lab light Ambient lab light Ambient lab light
6Light intensity NR NR NR NR NR NR NR NR 200 lux 100–1000 lux
7Photoperiod 16L:8D 24D Natural regime16L:8D 16L:8D 12L:12D NR 16L:8D 16L:8D 16L:8D
8Test chamber 100-mL beaker 250-mL or 300-
ml beaker
400-mL beakerBasket of mesh
netting in 4-L
chamber
12-well plate 6-well plate 12-well plate 12-well plate 200-mL dish or
300-mL
beaker
100-mL glass chamber
(minimum)
9Test solution
volume (mL)
50 200 200 NR 3.5 NR 53.5 100 75 (minimum)
10 Glochidia
collection
Shake piece of
cut gill in
water
Flush gills with
syringe
Cut gills and
press out
glochidia
using forceps
Flush gills with
syringe

Cut gills and
separate
glochidia
from
marsupia
NR Flush gills with
syringe
Flush gills with
syringe
Flush gills with
syringe
Flush gills with syringe
11 Age of test
organisms
(hours)
NR NR 3–24 NR NR NR ! 2NR ! 2to ! 24 ! 24
12 Number of
organismsper
test chamber
10 10 1000–3000 Several hundreds 50–75 50–100 40 50–100 About 1000 About 500 (1000 for repeated
sampling during atoxicity
test)
13 Number of replicate
chambers per
treatment
232, Counting 3
samples with
about 100
glochidia
2, Counting 3

samples with
about 100
glochidia
33or 43 33,Counting a
subsample
with about
100 glochidia
from each
replicate
3, Counting asubsample with
about 100 glochidia from
each replicate
14 Feeding None None None None None None None None None None
15 Aeration None None Yes None None NR NR NR None None, if dissolved oxygen is
maintained above acceptable
concentration
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16 Dilution water Reconstituted
water,
hardness
40–50 mg/L
as CaCO
3
Hardness
150 mg/L as
CaCO
3
Tap water Dechlorinated

effluent water
Dechlorinated
tap water or
Clinch River
water, VA
Reconstituted
water,
hardness 47–
76 mg/L as
CaCO
3
Sinking Creek
water, VA
Hardness
99–107 mg/L
as CaCO
3
Reconstituted
water,
hardness
170 mg/L as
CaCO
3
Depends on experimental design
17 Water quality DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,

alkalinity,
conductivity
pH, Ca, Cu,ZnDO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
ammonia,
hardness,
alkalinity,
conductivity
DO, pH, ammonia, hardness,
alkalinity, conductivity
18 Endpoint Survival (valve

closure with
culture
medium)
Survival (valve
closure with
NaCl)
Survival (valve
closurewith
KCl)
Survival (valve
closure with
NaCl)
Survival (valve
closurewith
NaCl)
Survival (valve
closure with
NaCl)
Survival (valve
closure with
salt solution)
Survival (valve
closurewith
saline
solution)
Survival (valve
closure with
NaCl)
Survival (valve closure with
NaCl)

19 Control survival
(%)
O 95 O 90 O 80 80 O 90 O 80 O 80 80 O 90 O 90 (must)
The Last Column Provides aSummary of Recommended Conditions That Can be Used to Conduct Toxicity Tests with Glochidia Based on ASTM (2006a)
a
See also Pynnonen (1995); Hansten et al. (1996).
b
Formerly Anodonta imbecillis.See also Weinstein (2001).
c
Formerly Lampsilis ventricosa.
d
V. iris, A. pectorosa, Pyganodon grandis, L. fasciola, Medionidus conradius.See also Jacobson (1990); Cherry et al. (2002).
e
Villosaosa lienosa, Villosa villosa, U. imbecillis, Megalonaias nervosa, Lampsilis teres, L. siliquoidea.See also Jacobson (1990), McCann (1993), V. iris, A. pectorosa, M. conradius.
f
Actinonaias ligamentina, Alasmidonta heterodon, Epioblasma capsaefotmis, Lampsilis siliquoidea, Lampsilis fasciola, Lampsilis abrupta, L. rafinesqueana, Potamilus ohiensis, Pleurobema plenum, Quadrula
quadrula, Quadrula pustulosa, Leptodea fragilis, Leptodea leptodon, Venustaconcha ellipsiformis, V. iris.
g
NA, not applicable; NR, not reported.
h
USGS unpublished data, Columbia, MO.
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bivalve mollusks (ASTM 2006c). Gravidfemale mussels are usually collected from the field and
held in the laboratory before isolatingglochidia to start atoxicity test (ASTM 2006a,Chapter4).
Alternatively, Zimmerman andNeves (2002) suggested glochidiaofsomespecies (including
Villosairis and Actinonaias pectorosa )couldbeextracted in the field from afemale and transported
back to the laboratory in cool water where glochidia can remainviable for several days without a
reduction in ability to successfully attachtoahost fish. This procedure may be particularly useful

whenglochidia of endangered species are extracted in the field and the female musselsare then
immediately returnedtotheir habitat. Mature glochidiaare typicallyflushed from the marsupium of
afemalemussel usingasyringe filled with water.Glochidiahave alsobeen isolated by cutting a
sectionofgill from the femalemussel and then teasingout the glochidia in water. (Thistechnique is
destructivetothe adultfemaleand maynot be appropriatefor useinisolating glochidiafor
conducting toxicity tests.)Nostudies were identified where glochidia were isolated for toxicity
testing from conglutinates released intothe water by femalemussels.
Before starting an exposure, the viability of glochidia is typicallyevaluated by aresponse
to the addition of aconcentratedsolution of NaCl or KCl.Mature and healthy glochidia will
snap shut in response to the addition of asaline solution. Immature glochidia isolated from the
marsupium of afemale will often still be enclosed in the egg membrane and will be fragile
andtend to fracture(ChrisBarnhart,Missouri StateUniversity,Springfield, MO,personal
communication). Tests are usually started if greater than 80 to greater than 90% viability of
the glochidia is observed (Huebner and Pynnonen 1992; Jacobson et al. 1997; Klaine, Warren,
and Summers 1997; ASTM 2006a). If immature glochidia are isolated from afemale mussel,
these glochidia should not be used for testing. Exposures are usually started the same daythat
glochidia are isolated from afemalebypooling glochidia from at least three females without
an extended acclimation period in the exposure water beforethe start of atoxicity test (ASTM
2006a). Theviability of glochidia isolated from each female should be evaluated before they
are pooled together. Toxicity tests can be conducted with glochidia obtained from one female
(e.g., when alimited number of endangered species are available for testing); however, the
results of tests conducted with alimited number of mussels should be interpreted with caution.
Additional research is needed to determinethe minimum number of females that should be
sampledtoobtain glochidia to start atoxicity test. This research might include an evaluation
of the variability in sensitivity of glochidia obtainedfrom individual females usingavariety
of toxicants.
ASTM (2006a)provides alist of recommended test conditions for conducting toxicity tests with
glochidia isolated from femalemussels. The list of recommended test conditions is based on the
various methods outlined in Table 5.1 and on the conditions used to conduct an inter-laboratory
toxicitytest with glochidia(ASTM2006a). ASTM (2006a)recommends that toxicity tests

with glochidiashouldbeconductedat20 8 Cwith a16L:8D photoperiod at an illuminance of
about 100–1000 lux (Table 5.1). The endpoint measured in toxicity tests with glochidia is survival
(viability)asdeterminedbythe response of organismstothe addition of asolutionofNaCl.
Glochidia that close their valves with the addition of asalt solution are classified as alive (viable)
in atoxicity test. For mostspecies, the duration of atoxicity test conducted with glochidia shouldbe
up to 24 hours with survival measured at 6and 24 hours.Control survival is typically greater than
90% at the end of 24-hourtoxicity tests conducted with glochidia. Longer duration toxicity tests with
glochidia (e.g., 48 hours) can be conducted as long as control survival greater than 90% is achieved.
However, toxicity tests conducted for greater than 24 hours with glochidia may not be as ecologi-
cally relevantgiven the short period of time betweenrelease of glochidia from afemale mussel until
encystment on ahost fish (ASTM 2006a;Chapter 4). Effect concentrations are typicallycalculated
based on the percentage of viable glochidia in the control at aparticular sampling time.
ASTM (2006a) recommends the use of glass test chambers for conducting toxicity tests with
glochidia. Test chambersshouldbeaminimum volume of 100 mL containing aminimum of 75 mL
of dilution water.Static, renewal, or flow-throughconditionscan be used depending on the
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chemical beingtested.Glochidia are not fed during the toxicity test, and aeration of dilution water is
typicallynot necessary. Dilution water shouldbeasourceofwater that has been demonstrated to
supportsurvival of glochidia for the duration of the toxicity test. For site-specific evaluations,the
characteristics of the dilution water should be as similar as possibletothe site of interest.The
number of replicates and concentrations tested dependsinpart on the significance levelselected and
the type of statisticalanalysis. ASTM (2006a)recommendsaminimum of three replicates should
be tested,each replicate containing about at least 500 glochidia (preferably 1,000glochidia/repli-
cate if survival is to be evaluated in subsamples of glochidia collected during the toxicity test).
Survival can be determined throughout the toxicity testbysubsamplingeach replicate (e.g., by
subsampling about 100 glochidia at 6and 24 hours and then placing these organisms into one well
of amulti-wellplate to determinesurvival with the addition of asaltsolution).
Toxicity tests with glochidia have been conducted for up to 144 hours,but 24 and 48-hour

exposuresare most oftenused (Table 5.1). Therelatively short durationoftoxicity tests with
glochidiaisbased on the relatively short duration betweenthe releaseofglochidia into the water
column and encystment on the host and on the relatively short survival time of glochidia after
isolationfrom the female mussel (Table 5.2). If the life history of the glochidia for aparticular
species is not known(e.g., the host required for encystment or how long glochidia released from a
female mussel can remain in the water column before encysting on ahost), it might be appropriate
to conducttoxicity tests with glochidia for longer than 24 hours as long as 90% control survival can
be achieved at the end of the test (ASTM 2006a).
IssuesRegarding the Use of Methods
Glochidia and juvenile musselsofseveral genera have been found to be highly sensitive to some
metals and to ammoniainwater exposures compared to many of the most sensitive genera of other
invertebrates,fish,oramphibiansthatare commonlytested (Chapter 7, Cherry et al.2002;
Augspurger et al. 2003; USGS 2006a, 2006b). However, concerns have been expressedregarding
the useoftoxicity data generated with glochidiainthe derivationofWQC (CharlesStephan,
USEPA,Duluth, MN;personal communication). These concerns mainly include:(1) the duration
of the toxicity tests, (2) the quality of the glochidia at the start of atest, and (3) the test acceptability
criteria. The following section provides information that attempts to address these concerns. Areas
of ongoing researchorneed for future researchare alsoidentified.
Duration of the Toxicity Test
1. How long shouldacute tests with glochidia be conducted (i.e., based on the life historyof
the species)?
There are nearly 300 species of freshwater musselsinNorth America, and the length of
time that glochidia remain viable after release from the marsupium of afemale into the
environment dependsonthe life history of the species and the temperature of the water
(Chapter 3). Longevity of glochidia after release and before attachment to ahost may
exceed one weekand may be dependent on temperature (Zimmerman and Neves 2002);
however, some reports are anecdotal (Murphy 1942;Matterson1948; Tedlaand
Fernando 1969). Glochidia of some species released in conglutinates remainviable for
days or weeks after releaseinto the environment (Chris Barnhart, personal communi-
cation). Glochidiaofseveral species,including Anodonta spp., remainviable while free

in the environment for 7–14 days (Howard and Anson 1922; Mackie 1984; Huebner and
Pynnonen 1992; Pynnonen 1995).
Table 5.2 provides asummary of laboratory studies that have evaluated survival times
of glochidia after removal from the marsupium of the female or survival time based on
results reported in toxicity tests conducted with glochidia. For example, Zimmerman and
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TABLE 5.2
SurvivalTime of Glochidia after Removal from Female Unionid Mussels
Duration of Viability
Species
Temperature
( 8 C) Day(%Survival) Reference
Actinonaias ligamentina 20 7(O 90); 8(O 75); 9(O 50) USGS (2004)
Actinonaias pectorosa 10 13 ( O 75) Zimmermanand Neves (2002)
25 5(O 75) Zimmermanand Neves (2002)
20 O 2(O 90)
a
Jacobson et al. (1997)
Alasmidonta heterodon 20 2(O 90); 2(O 75); 2(O 50) USGS (2004)
Anodonta anatine 13 O 3(O 90) Huebner and Pynnonen(1992)
Anodonta cataracta 10 O 14 ( O 90) Jacobson (1990)
Anodonta cygnea 13 O 3(O 90) Huebner and Pynnonen(1992)
Anodonta grandis 10 O 14 ( O 90) Jacobson (1990)
Elliptio complanata 57NR
b
Matterson (1948)
20 ! 1(O 90); 3(O 75) Bringolf et al. (2005)
Elliptio dilatata 20 ! 1(O 90); 1(O 75); ! 2(O 50) Bringolf et al. (2005)

Epioblasma capsaeformis 20 0.3 ( O 90) Wang et al. (2003)
Lampsilis abrupta 20 2(O 90); 5(O 75); 7(O 50) USGS (2004)
Lampsilis cardium 21 O 2(O 90)
a
Lasee (1991)
Lampsilis fasciola 20 6(O 90); 7(O 75); 8(O 50) Wang et al. (2003)
20 O 2(O 90)
a
Jacobson et al. (1997)
20 1(O 90); 2(O 75); 3(O 50) Bringolf et al. (2005)
20 2(O 90; 4(O 75); 5(O 50) Bringolf et al. (2005)
Lampsilis rafinesqueana 20 6(O 90); 6(O 75); 6(O 50) USGS (2004)
Lampsilis siliquoidea 10 9NRTedla and Fernado(1969)
20 8(O 90); 9(O 75); 10 ( O 50) Wang et al. (2003)
25 O 2(O 80)
a
Keller and Ruessler (1997)
20 1(O 90); 3(O 75); 4(O 50) Bringolf et al. (2005)
Limpsilis teres 25 0.2 ( O 80) Keller and Ruessler (1997)
Leptodea fragilis 20 1(O 90); 3(O 75); 4(O 50) Wang et al. (2003)
Leptodea leptodon 20 1(O 90); 2(O 75) Bringolf et al. (2005)
Leptodea leptodon 20 0.25 ( O 90); 1(O 75); 2(O 50) USGS (2004)
Margaritifera falcate 11 11 NR Murphy (1942)
Medionidus conradius 20 O 2(O 90)
a
Jacobson et al. (1997)
Megelonaias nervosa 25 1(O 80)
a
Keller and Ruessler (1997)
Potamilus alatus 20 6(O 90) 6(O 75); 6(O 50) Wang et al. (2003)

Potamilus ohiensis 20 5(O 90); 6(O 75); 7(O 50) Wang et al. (2003)
Pyganodon grandis 20 O 1(O 90)
0
Jacobson et al. (1997)
Quadrula quadrula 20 1(O 90); 1(O 75); 2(O 50) Wang et al. (2003)
Quadrula pustulosa 20 ! 1(O 90); 1(O 75); 1(O 50) Wang et al. (2003)
Utterbackia imbecillis 21 10 ( O 80); 14 ( O 50) Fisher and Dimock (2000)
25 O 2(O 80)
0
Keller and Ruessler (1997)
25 O 2(O 80)
a
Klaine, Warren, and Summers (1997)
20 O 1(O 90)
a
Johnson, Zam, and Keller (1990,
1993)
Venustaconcha
ellipsiformis
20 2(O 90); 3(O 75); 3(O 50) Wang et al. (2003)
Villosa iris 10 8(O 75) Zimmermanand Neves (2002)
20 5(O 90); 5(O 75); 6(O 50) Wang et al. (2003)
25 2(O 75) Zimmermanand Neves (2002)
(continued)
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Neves (2002) report that the viabilityofglochidia of V. iris was greater than 75% for
8days at 108 Cand 2days at 258 C, and viability of glochidia of A. pectorosa was greater
than 75% for 13 days at 108 Cand 5days at 258 C(Table 5.2). Similarly, glochidia of

Utterbackiaimbecillis maysurviveupto19daysbut exhibit50% mortalitywithin
13.5 days (Fisherand Dimock 2000).Survivalofisolated glochidiafrommany
species listedinTable 5.2istypically greater than 90%after twotothree days;
however, theviability of glochidiafor aparticular speciesshouldbedetermined
before the start of an exposure. For example, glochidia of L. teres and E. capsaeformis
were viable for only four to six hours and glochidiaof M. nervosa and Q. quadrula were
viable for one day after removal from the marsupium of the female (Table 5.2). There-
fore, 24 hours is areasonable time period to conducttoxicity tests with glochidia of many
species at 208 C, although shorter or longertests might be neededfor aparticular species
depending on glochidia survival time and the life history characteristics of the species
(i.e., survival of glochidia in the control mustbegreater than 90% at the toxicity test)
(ASTM 2006a).
Thetime between the release of glochidia from the marsupium of the female mussel to
attachment of these glochidia on ahost may only take afew seconds for some species,
but hours are requiredfor the gill tissue of afish to migrate to form acyst aroundthe
glochidia. Duringthat time,the glochidiamay be exposedtowater-borne toxicants.
Anodontinae speciesreleasesglochidia directly into water, whichremain viable for
days in order to effectively infest their host fish. Therefore, aprolonged glochidial test
would have ecological relevance for these species.Other species releaseglochidia in
mucus strands that coat the bottom or remain suspended on vegetation,waiting for their
hosts to swim by, and still otherspecies package glochidia in conglutinates that serve as a
lure to host fish.Hence, glochidia of these species may also be in water for extended
periods of time;however,itisnot known how exposure to water-borne contaminants
would be influenced by the mucus or conglutinatesurrounding the glochidia. Toxicity
testsconducted for24hourswithglochidia maynot be as ecologicallyrelevantas
toxicity tests conducted with juvenile mussels, but they may be useful for somepurposes
such as deriving concentrations of achemical that may be protectiveofthe species. Use
of glochidia to evaluatethe relative sensitivity of aparticular mussel species to chemicals
would be particularly useful when evaluating species where only alimited number of
adult mussels are available for methodsdevelopment or for producing juvenile mussels

for toxicity testing. Moreover,the host fish for somespecies of mussels or techniques for
transforming juvenile musselsinthe laboratory may be unknown (Chapter 4).
TABLE 5.2 (Continued)
Duration of Viability
Species
Temperature
( 8 C) Day(%Survival) Reference
22 O 1(O 80)
a
Goudreau,Neves, and Sheehan
(1993)
20 O 1(O 80)
a
Scheller (1997)
20 O 2(O 90)
a
Jacobson et al. (1997)
Villosaosa lienosa 25 O 2(O 80)
a
Keller and Ruessler (1997)
Villosa nebulosa 20 O 2(O 90)
a
Jacobson (1990)
Villosa villosa 25 O 2(O 80)
a
Keller and Ruessler (1997)
a
The value based on control survival in 24- or 48-hour toxicity tests.
b
NR, not reported.

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Therelatively short duration of toxicity tests with glochidia is based on the relatively
short duration betweenreleaseofglochidia into the water column and encystment on the
host and on the relatively short survival time of glochidia after isolation from the female
mussel. If the life historyofaparticularspecies is not known (e.g., the host requiredfor
encystment or how long glochidia released from afemale mussel can remaininthe water
column beforeencysting on ahost), it might be appropriatetoconducttoxicity tests with
glochidia for longerthan 24 hours as long as 90% control survival can be achieved at the
end of the test.
2. How long can glochidia survive and still be able to attach to ahost?
Glochidia of somespecies can still attachtoahost for several days after release from a
female depending on temperature (Chris Barnhart,personalcommunication).The
maximum time at which greater than 50% of U. imbecillis metamorphosed in atissue
culture medium was nine days after isolation from afemale(Fisher and Dimock 2000).
Zimmerman and Neves (2002) reported that glochidia can successfully attachtoahost one
to two weeks after isolation from afemale. Afuture research project could be to conducta
series of toxicity teststodetermine if thereisachangeinsensitivityovertimeafter
glochidia have been released into the environment.Sensitivity of L. siliquoidea glochidia
held for 24 hours after isolationfrom afemale was similar to newly-released glochidia in
exposures to copper (Wang et al. 2003). The sensitivity of glochidia held in an extra piece
of the marsupium in arefrigerator overnight was similar to the sensitivity of glochidia
tested immediately after isolation from afemale in toxicity tests conducted with zinc or
copper (Jerry Farris, Arkansas State University, State University, AK; personal communi-
cation). Ultimately, it is more practical to base duration of exposure on survival of control
organisms in the laboratory rather than on an estimate of the length of time glochidia can
survive and still attachtoahost (e.g., Table 5.2).
3. Are there data that indicate that effect concentrations do not change very muchduring the
last half of atoxicity test (i.e., does the EC50 at 6, 24, 48, or 96 hours differ)?

There are limited studies with glochidia that have comparedchangesintoxicity over this
timeframe.The toxicity of copper (Jacobson et al. 1997; Wang et al. 2003), ammonia
(Wang et al. 2003), and chlorine (Wang et al. 2003)decreasedover 48–96-hour exposures.
In contrast, no change in the toxicity of several pesticides was observed in 24–48-hour
exposures (Keller and Ruessler 1997; Bringolf et al. 2005). If glochidia for aparticular
species are able to survive for more than 24 hours, then a24-hourtoxicity test shouldbe
considered.Importantly,researchers areencouraged to design studies that generate
toxicity data throughout the exposure period (e.g., reporting 6, 24, and 48-hourresponses)
(ASTM 2006b). However, generating data for asix-hour exposure period is logistically
difficult in an eight-hour day.
Quality of Glochidia at the Star tofaToxicity Test
1. How shouldthe quality of glochidia be determined at the start of atoxicity test? Is the
use of asolution of NaCl (or KCl)todeterminethe percentage of glochidia exhibiting
valve closure an appropriate method to judge the acceptability of glochidia used to start
atoxicity test? Does the response of glochidia to asolution of NaCl (or KCl)relate to
the ability of glochidia to attachtoahost?Isthere an independent way of determining
if glochidia are alive or healthy at the start (or end) of atoxicity test?
Valve closure is an ecologically relevantendpoint that is criticalfor glochidia to
successfully transform on the host.Ifglochidia do not snap shut, the glochidia should
be considered ecologically dead (Huebner and Pynnonen 1992; Goudreau, Neves, and
Sheehan 1993;McCann 1993;Jacobsonetal. 1997). Theresponse of glochidiain
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toxicity tests was similar wheneither KCl or fish plasma was used to makeglochidia
close at the end of an exposure (Huebner and Pynnonen 1992). Decreased response to
KClwas considered an indication of reduced glochidiaviabilityand thus reduced
capabilitytoattachtothe fishhost(Pynnonen 1995). Asignificantcorrelation
was observed betweenthe response of glochidia to KCl and the ability of glochidia
of U. imbecillis to metamorphose to the juvenile life stage (Fisher and Dimock 2000).

Zimmermanand Neves(2002) reported acorrespondencebetween theresponseof
glochidiaof V. iris and A. pectorosa to NaCl and theabilitytoinfest ahostfish.
Jacobson et al. (1997) reportedglochidiaof V. iris that responded to the addition of
NaCl followinganexposuretocopperwere able to attach to ahostfish with no
impairmentofsubsequent metamorphosistojuvenile mussels. Results of these
studies indicatethat addition of asolution of NaCl or KCl can be used to estimate
the condition of glochidia. While either asolution of salt or fish plasma could be used
to determine thepercentageoforganisms closing,itiseasiertoworkwith NaCl
comparedtoKCl or fish plasma.
2. Should therebeaholding time for glochidia after harvesting but beforeapplication of a
saline solution to determineifglochidia that are initially closed might open?
Mature glochidia are not typically closed after being isolated from afemale mussel.
Glochidiathat are closed after isolationfrom afemale may reopenafter being held in
clean water afew hours (Goudreau, Neves, and Sheehan 1993;Dick Neves, Teresa
Newton, USGS, LaCrosse, WI; personal communications).
3. Will immature,stressed, or unhealthy glochidiaclose when exposedtoasaline
solution?Couldglochidiabealive andsuccessfully attach to ahostbut not close
when exposed to asaline solution? Are brokenglochidia frequently observed at the
start of atest? Wouldthe presence of brokenglochidiabeindicative of stress during
harvesting?
Immature glochidiathatare free of theegg membrane or mature andhealthy
glochidiawill closewhenexposed to asalinity challenge. However, immature
glochidiaare generally enclosed in an eggmembraneand arefragile andtendto
fracture, thus should not be used for toxicity testing. The best approach for avoiding
the use of immatureglochidia in toxicity testing is to sample female mussels at atime
of the year whenthe organisms would be expected to be releasing matureglochidia
(JessJones, US GeologicalSurvey, Blacksburg,VA; personalcommunication).
Stressed or unhealthy glochidiacouldeither be opened or closed before the start of a
test. If stressed or unhealthy glochidia weretoclosewhenexposedtoasalinity chal-
lenge, then theseindividuals would be used in atoxicity test. Measurementofthe

viability of glochidia in the control at the end of atoxicity testwould help to identify
stressed or unhealthy glochidia. Results of reference-toxicant tests shouldalso be used
to evaluate the health of the glochidia used to conductthe test(ASTM 2006a). Broken
glochidia have not been observed at the start of atest (Chris Barnhart, Jerry Farris, Dick
Neves, TeresaNewton, Ning Wang, USGS, Columbia, MO; personal communications).
The presence of broken glochidia may indicate that the glochidia are immature and
should not be used for testing.
Test Acceptability Criteria for Toxicity Tests with Glochidia
1. What criteria should be used to judge acceptability of atoxicity test conducted with
glochidia?
ASTM (2006a)recommends that the age of glochidia should be less than 24 hours
old at the start of the toxicity test. Viability of glochidia isolated at the beginning of a
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toxicity testmust be greater than or equal to 80% (preferablygreater than or equal to
90%). Average survival of glochidia in the control at the end of atest mustbegreater
than or equal to 90%. ASTM (2006a)also recommends that subsamples of each batch
of test organisms used in toxicity tests shouldbeevaluated usingareference toxicant
(e.g., NaCl or CuSO
4
). Data from thesereference-toxicant tests can be used to assess
genetic strain or life-stage sensitivity of test organisms to select chemicals.
2. Should glochidia be rinsed before use in atoxicity test? Would rinsing glochidia before
the start of atest be stressful to the organisms?
Glochidiashould be rinsedwithculture or dilution waterafter removalfrom
marsupia to (1)eliminate tissuesorexcessmucus from theexcised glochidia that
haveahigh potential forfungal growth and subsequently could affect thesurvival
(toxicity tests) or transformation of glochidia(propagation) and(2) reduce the
number of protozoans that may be present in the excised gill that could also affect

glochidiasurvivalortransformation(ASTM2006a). Rinsed glochidiahavebeen
observed to successfully transformonfish or in artificial mediaand high control
survivalintoxicity testshas been reportedusing glochidiathathavebeen rinsed
(Huebner and Pynnonen 1992; Johnson, Keller, and Zam 1993; Myers-Kinzie 1998;
Milam et al. 2005).
3. Should glochidia be acclimated to test conditions before the start of atoxicity test?
Glochidia are not typically acclimated to the water-quality characteristics of the
dilution water beforethe start of atoxicity test(Table 5.1). Most of these exposures
are started the sameday that glochidia are isolated from marsupia of the females.
Therefore, minimaltimeisavailable to acclimateglochidia to thedilution water
before thestartofatest.Inorder to maintain organismsingood condition and
avoid unnecessary stress,ASTM (2006a)recommends that organisms shouldnot be
subjected to rapid changesintemperature or water quality beforethe start of atest.
Glochidiacan be acclimated in amixture of 50% culturewater and 50% test water
and gradually adjustedtothe test temperature within about two hours beforethe start
of an exposure(ASTM2006a). Investigators have held adultmussels undertest
conditions before isolationofglochidia (e.g., Huebner and Pynnonen 1992), which
would result in acclimating glochidia to the selected exposure temperatureinthe
toxicity test. However, brooding glochidia in the marsupium are in contact with the
hemolymph of the femalethat is physically isolated from direct contact with water
(Silverman, McNeil, and Dietz1987). In addition, glochidia are typically released
instantaneously into the surrounding water from the marsupium of the femalemussel.
Therefore, holding the female mussels in the dilution water beforeisolatingglochidia
fortoxicity testing wouldprobablyhaveaminimal influence on theability of
glochidia to acclimate to the conditions of the dilution water.
M ETHODS FOR C ONDUCTING W ATER-ONLY T OXICITY T ESTS WITH J UVENILE
F RESHWATER M USSELS
ReviewofMethods
ASTM (2006a)provides alist of recommendedtest conditions for conducting toxicity tests with
juvenile mussels. The list of recommendedtest conditions is based on the various methods outlined in

Table 5.3 and on the conditions used to conductaninter-laboratory toxicity test with juvenile mussels
(ASTM 2006a). ASTM (2006a)recommends that toxicity tests with juvenile mussels beconducted at
208 Cwith a16L:8D photoperiod at an illuminance of about 100–1,000lux (Table 5.3). Toxicity tests
are typicallystarted with newly-transformed juvenile mussels less than five days after the release
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TABLE 5.3
SummaryofTest ConditionsUsed to Conduct Toxicity Tests with Juvenile Freshwater Mussels
Conditions
Johnson et al.
(1990, 1993)
Jacobson
(1990),
Jacobson
et al. (1993) Lasee (1991)
Keller and Zam
(1991)
Klaine,
Warren, and
Summers
(1997)
Scheller
(1997)
Myers-Kinzie
(1998)
Dimockand
Wright
(1993)
Newton

et al.
(2003)
Lasee
(1991)
Wade
(1992)
a
Jacobson
(1990)
Valenti
et al.
(2005)
USGS
(2004)
USGS (2005a,
2005b, 2005c)
Recommended
Test Conditions
ASTM (2006a)
1Species
tested
Utterbackia
imbecillis
b
Villosa
nebulosa,
Villosa iris,
Anodonta
grandis
c

Lampsilis
cardium
d
Mulitple
species
e
U. imbecillis V. iris Lampsilis
siliquoidea
U. imbecillis,
Pyganodon
cataracta
L. cardium Lampsilis
ventricosa
U. imbecillis Villosa
nebulosa
V. iris Mulitple
species
f
L. siliquoidea,
Epioblasma
capsaeformis,
V. iris
g
NA
h
2Test type Renewal Static Static Static Static Static NR Static Flow
through
Renewal Renewal Artificial
stream
Renewal Renewal,

flow
through
Flow through Static, renewal or
flow-through
(depending on
duration of
exposure and
chemical tested)
3Test duration
(days)
2121–4 1–4 41,2,4 1–4 4, 10 7914 21 2, 4, 10 28 Acute:% 4
Chronic:
21–28
4Temperature
( 8 C)
20 20 21 22, 25, or 32 25 25 24 20 21 21 24 20 20 20 20 20
5Light quality Ambient lab
light
NR
h
NR NR NR NR NR NR Fluorescent NR NR NR NR Fluorescent Fluorescent Ambient lab light
6Light
intensity
NR NR NR NR NR NR NR NR NR NR NR NR NR 200 lux 200 lux 100–1000 lux
7Photo period 16L:8D 16L:8D 24 D12L:12D or
16L:8D
16L:8D NR NR NR 16L:8D 24D 24D 16L:8D 12L:12D 16L:8D 16L:8D 16L:8D
8Test
chamber
125-mL

beaker
12-well
plate
Covered
250-mL
crystallizing
dish
Petri
dish
Petri
dish
12-well
plate
Petri
dish
120-mm
diam. tub
with mesh
bottom in
4-L chamber
132 by
90 by
130 mm
chamber
Covered
250-mL
crystall-
izing
dish
50-mm

diam. glass
tub with
mesh
bottom
in 250-mL
Chamber
Dish
covered
with mesh
30-mL
beakers
submer-
ged in a
1-L glass
beaker
50- or
300-mL
beaker
300-mL
beaker
Static or
renewal: 50-mL
beakers
(minimum);
flow-through:
300-mL
beakers
(minimum)
9Test solution
volume

(mL)
100 3.5 NR 15 15 510NR1200 NR 200 150 950 30 or 200 200 Static or renewal:
30 (minimum);
flow-through: 200
(minimum)
10 Procedure
for
obtaining
juveniles
Artificial
media
Fish host Fish host Fish host
or artificial
media
Fish host or
artificial
media
Fish host Artificial
media
Artificial media Fish host Fish host Artificial
media
Fish host Fish host Fish host Fish host Fish host
11 Age of test
organisms
(days)
1–10 1–3 0, 7, 14 1–2 1–3 ! 3, 5, 9 ! 10 7–10 3–5 06–10 1–3 60 3–5, 60 60 Acute: ! 5
Chronic:
60–120
12 Number of
organisms

per test
chamber
10 10 10 10–20 15NR 10 20 50 15 15 55 10 Acute:! 5
(minimum)
Chronic:10
(minimum)
(continued)
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TABLE 5.3 (Continued)
Conditions
Johnson et al.
(1990, 1993)
Jacobson
(1990),
Jacobson
et al. (1993) Lasee (1991)
Keller and Zam
(1991)
Klaine,
Warren, and
Summers
(1997)
Scheller
(1997)
Myers-Kinzie
(1998)
Dimockand
Wright

(1993)
Newton
et al.
(2003)
Lasee
(1991)
Wade
(1992)
a
Jacobson
(1990)
Valenti
et al.
(2005)
USGS
(2004)
USGS (2005a,
2005b, 2005c)
Recommended
Test Conditions
ASTM (2006a)
13 Number of
replicate
chambers
per
treatment
22or 33 2–4 10 4NR3 62 3344 4Acute:4
(minimum)
Chronic:3
(minimum)

14 Feeding None None None None None None None None None Lab
cultured
phyto
plankton
Algae
and
silt
Algae Algae
and
sedi-
ment
None Instant
algae
mixture
i
Acute:none
Chronic
Algae
15 Aeration None None Yes NR None NR NR Yes Yes None None None Yes None None None, if dissolved
oxygen is
maintained above
acceptable
concentration
16 Dilution
Water
Reconstituted
water,
hardness
40–50
mg/L as

CaCO
3
Clinch River
water, VA
Hardness
150 mg/L as
CaCO
3
Reconstituted
water, hardness
47–76 mg/L as
CaCO
3
Recon
stituted
water,
hardness
99–107
mg/L as
CaCO
3
Sinking
Creek
water, VA
Hardness 100
or 200 mg/L
as CaCO
3
NR Hardness
133

mg/L as
CaC0
3
Hardness
150
mg/L
as CaC0
3
Tennessee
River
water
Clinch
River
water,
VA
Reconstituted
water, hardness
100 mg/L as
CaCO
3
Reconstituted
water, hardness
170 mg/L as
CaCO
3
Reconstituted
water-
hardness
170 mg/L as
CaCO

3
Depends on experimental
design
17 Water
Quality
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conductivity
DO, pH,
hardness,
alkalinity,
conduc-
tivity

pH,
hardness
NR DO, pH,
hardness,
alkalinity,
condu-
ctivity
DO, pH,
hardness,
alkalinity,
conducti-
vity
DO, pH,
hardness,
alkalinity,
conducti-
vity
NR NR DO, pH,
ammonia,
hardness,
alkalinity,
condu-
ctivity
DO, pH,
ammonia,
hardness,
alkalinity,
conductivity
DO, pH, ammonia,
hardness,

alkalinity,
conductivity
18 Endpoints Survival
(move-
ment)
Survival
(gaped
valves, foot
activity or
stained with
neutral red)
Survival
(foot or
ciliary
movement)
Survival
(activity
and
heartbeat)
Survival
(gaped
valves
with foot
and ciliary
activity)
Survival
(heart
beat and
ciliary
action)

Survival
(foot or
valve
move
ment)
Survival (foot,
valve or
ciliary
activity,
heartbeat)
Survival,
growth,
ratio of
stressed to
alive
Survival
(foot or
ciliary
move-
ment),
growth
(length and
height)
Survival
(Ciliary
action)
Survival
(extruded
foot
and

gaping
valves)
Survival,
growth
Survival
(foot or
shell
move-
ment) and
growth
(shell
length)
Survival (fool or
shell movement)
and growth
(shell length)
Survival (foot
movement),
growth (shell
length)
19 Control
survival
(%)
O 95 100 96 NR O 90 O 80 99 O 90 O 95 97 O 90 100 90 O 90 O 88 Acute:O 90 (must)
Chronic:O 80
(should)
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The last column provides asummary of recommended conditions that can be used to conduct toxicity tests with juvenile mussels as outlined in ASTM (2006a). In the last Column, Acute Tests are Tests Conducted for up to 96 hours and Chronic Tests are Tests Conducted for at

least 21 days.
a
See also Masnado, Geis, and Sonzogni (1995), McKinney and Wade (1996), Keller, Ruessler, and Kernaghan (1999).
b
Formerly A. imbedllis.
c
See also McCann (1993) for two- to four-day exposures with Villosa iris, A. pectomsa, M. conradius.
d
Formerly L. ventricosa.
e
A. imbecillis, V. lienosa, V. villosa, U. imbecillis, Lampsilis straminea daibomensis, L. subangulata, Elliptic icterina.See also Keller (1993), Keller and Ruessler (1997).
f
V. iris, E. capsaeformis, L. fasciola, L. siliquoidea, L. abrupta,L.rafinesqueana, L. leptodon.
g
Bringolf et al. (2005) has adapted this method to conduct 21-day toxicity tests with four-month old juvenile A. ligamentina.
h
NA, not applicable; NR, not reported.
i
See USGS (2005a) for adescription of the procedure used to prepare this instant algae mixture.
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from the host;however,sometoxicity tests have been started with two- to four-month-old juvenile
mussels. Acute toxicity tests with juvenile mussels are typically conducted for 96 hours with survival
measured at 48 and 96 hours. Chronic toxicity tests started with two- to four-month-old juvenile
mussels have been conducted for 21–28 days with measures of survival (based on movement of the
foot) and growth (based on shell length).Control survival is typically greater than 90% at the end of
96-hourtoxicity tests conducted with juvenile mussels and is typicallygreater than 80% at the end of
toxicity tests conducted for 10–28 days with juvenile mussels(Table 5.3; ASTM 2006a).
In acute static tests, glass test chambersshouldbeaminimum volume of 50 mL containing a

minimum of 30 mL of dilution water (ASTM 2006a). In chronic tests or in flow-through tests, glass
chambersshould be aminimum volume of 300 mL containing aminimum volume of 200 mL of
dilution water. Static,renewal, or flow through conditions can be used depending on the chemical
beingtested.Juvenilemussels are not typicallyfed during acute toxicity tests. Algae have been used
as afood source in toxicity tests conducted for 10–28 days (Table 5.3; ASTM 2006a).
Thenumber of replicates and concentrations tested dependsinpart on the significance level
selected and the type of statistical analysis. In 96-hourtoxicity tests, ASTM (2006a)recommendsa
minimum of 20 organisms shouldbeexposed to each concentration (e.g., four replicateseach
containing aminimum of fivejuvenile mussels). It may be desirable to test only five juvenile
mussels in each replicate whenalimited number of testorganisms are available or whentest
organismsare relativelysmall (e.g., when juvenile musselsare small, it maybedifficultto
observe more than about five testorganisms simultaneouslyinareplicate test chamber under the
microscope). However, some investigators have tested 10–20 juvenile mussels in each replicate. In
chronic toxicity tests, aminimum of three replicates shouldbetested,each replicate containing a
minimum of 10 juvenile mussels.
Toxicity tests with juvenile mussels are typicallystarted with organisms that have been trans-
formed with afish host (ASTM 2006a); however, artificial media has alsobeen used to transform
juvenile mussels for use in toxicity testing (Johnson, Keller, and Zam 1993; Clem 1998; Hudson et al.
2003). ASTM (2006a) recommendstesting of juvenile mussels that have been transformed on afish
host due to uncertaintiesregardingthe sensitivity of juvenile mussels transformed using artificial
media. Numerous investigators have observed high mortality of juvenile musselsabout four to six
weeks after transformation (e.g., Anne Keller, USEPA, Athens, GA; Don Cherry, Jerry Farris, Teresa
Newton; personal communications). As aresult of this problem,the duration of toxicity tests started
with newly-transformed juvenilemussels is lessthan 14 days with survival or growth measured at the
end of the exposures (Table 5.3). Food (mixturesofdifferent species of algae) and sediment have
been addedtoexposurechambers, but some investigatorshavefound that newly-transformed
juvenile mussels will survive for at least 14 days without the addition of food (Table 5.3;ASTM
2006a). For example, USGS (2004) determined the acute toxicity of copper in 48-hourtests with
juvenile L. siliquoidea and Lampsillis rafinesqueana that had been held for 10 days under control
conditions (e.g., with the replacement of dilution water but without the addition of food). Similar

48-hourEC50swere observed in tests conducted with juvenile mussels held for 10days before testing
compared to tests started with newly-transformed juvenile mussels. Results of these tests indicate
that the sensitivity of juvenile mussels did not change over the 10-dayexposure withoutfeeding.
Hence, toxicity tests conducted for up to 10 days without feeding may provide reliable data for
evaluating effects of chemicals on musselsinexposures longerthan 4days.
Thehigh mortalityofnewly-transformed juvenilemusselsintoxicity tests conductedfor
longerthan 14 days is likely related to alackofunderstandingofthe nutritional requirements
of mussels at this life stage. Newly-transformed juvenile musselsdepend on pedal feeding to
obtain food (cilia on the foot are used to move food intothe juvenile mussel). Juvenilemussels
gradually begin to use acombination of pedal and filter feeding to obtain food until the mussels
eventually depend on filter feeding to obtain food by about six months in laboratory cultures
supplied with asilt-clay sediment substrate. However, in the field, juvenile mussels probably
depend on acombination of filter, deposit and pedal feeding in coarser substrates (Dick Neves,
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personal communication). Research is ongoing to improveculturing methods for propagation,
holding, and feeding of newly-transformed juvenile mussels (Chapter 4; Keller and Zam 1990;
Gatenby, Neves, and Parker1996, 1997; Henley, Zimmerman, and Neves 2001; ASTM 2006a).
Once developed,theseculturing methods shouldhelp to refine methods for conducting chronic
exposures with juvenile mussels.
Investigators have reportedsuccess in conducting toxicity tests for up to 28 days starting with
two- to four-month-old juvenile mussels. Valenti et al. (2005) conducted 21-dayexposures to
mercurystarting with two-month old juvenile V. iris held in asmall amount of sediment and fed
algae ( Neochloris ). USGS(2005a,2005b, 2005c); Bringolf et al. (2005) conducted toxicity tests
starting with two- to four-month-old juvenile A. ligamentina, L. siliquoidea,or V. iris and observed
controlsurvivalgreater than 88% in 21–28-day exposures to copper, lead,zinc, cadmium,
ammonia, and several pesticides whenamixture of instant algae was used as afood source. The
instant algae mixture was prepared from commercial Instant Algae brand non-viable microalgae
concentrates (Reed Mariculture,Campbell, CA including Nannochloropsis, Isochrysis, Pavlova,

Tetraselmis,and Thalassiosira weissflogii).
IssuesRegarding the Use of Methods
Concernshavebeenexpressedregarding the usetoxicity data generated with glochidiainthe
derivationofWQC (Charles Stephan, personal communication). Charles Stephan concluded that
acute methods for testing juvenile mussels(such as thoseoutlined in Table 5.3)generally follow
standardtesting methods (e.g., ASTM 2006a,2006b),and data generatedfromthese typesof
studiesshould be useful in thederivation of WQC. However, therewere concernsidentified
regardingtoxicity tests conductedwithjuvenilemussels including:(1) thelife stagetested,
(2) the determination of deathatthe end of atest, and (3) test acceptability criteria. Thefollowing
sectionprovides information that attempts to address some of these concerns. Areas of ongoing
researchorneeds for future researchare alsoidentified.
What Life Stage Should Be Used to Start Acute or Chronic Toxicity Tests
with Juvenile Mussels?
Toxicity tests have been started with newly-transformed juvenile mussels that have either been
transformed on ahost or have been transformed with the use of an artificial medium (Table 5.3).
Glochidia, newly-transformed juvenile mussels, and two- to four-month-old juvenile mussels have
been successfully shipped via overnight carriers to otherlaboratories for use in toxicity testing
(USGS 2004; Bringolf et al. 2005; ASTM 2006a). Toxicity tests have been successfullyconducted
for 10–14 days starting with newly-transformed juvenile mussels (Table 5.3), but exposures Started
with newly-transformed juvenile mussels conducted for longer periods of time have resulted in high
mortality in controls at about four to six weeks, probably due to nutritional limitations of the diet
(e.g., Newton et al. 2003). Valenti et al. (2005),USGS (2005a,2005b, 2005c) and Bringolf et al.
(2005) conducted 21–28-day toxicity tests starting with two- to four-month-old juvenile musselsof
avarietyofspecies and observed control survival greater than 88% when algae was used as a
food source.
How Should the Death of Juvenile Mussels Be Determined at the End
of aToxicity Test?
Lack offoot or shell movement, lack ofciliary activityonthe foot, lack of aheart beat, or awidegaped
valve have been used to establish death in toxicity tests with juvenile mussels (Table 5.3). ASTM
(2006a)recommendsestablishing death of juvenile mussels based on foot movement during afive-

minuteobservation period under amicroscope. If it is suspected that juvenile mussels are avoiding
exposure to achemical in atoxicity test, it may be desirable to place the suspected live test organisms
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into dilution water that does not contain any added test materialfor one to two days after the end of the
toxicity test to determine whether these test organisms are alive or dead (ASTM 2006a).
What Criteria Should Be Used to JudgeAcceptabilityofaToxicity Test Conducted
with Juvenile Mussels?
ASTM (2006a) recommends that average survival of juvenile musselsinthe control at the end of a
96-hourtestmustbegreater than or equal to 90%. An insufficient numberoftests havebeen
conductedwithjuvenilemussels for10ormoredaysfor ASTM(2006a) to providespecific
guidance on controlsurvivalinlonger-termtests.However, alimitednumberoftoxicity
tests have reported control survival greater than 80%intests conducted with juvenile mussels
for10–28 days.Therefore,ASTM (2006a)recommendsthataverage survival of juvenile
mussels in the control at the end of atest conducted for 10–28 days should be greater than or
equal to 80%. ASTM (2006a) also recommends that subsamples of each batch of test organisms
used in toxicity tests should be evaluated usingareference toxicant(e.g., NaCl or CuSO
4
). Data
from these reference-toxicanttests can be used to assess genetic strainorlife-stage sensitivity of
test organisms to select chemicals.
M ETHODS FOR C ONDUCTING W ATER-ONLY T OXICITY T ESTS WITH A DULT F RESHWATER M USSELS
ReviewofMethods
Conditions that have been used to conducttoxicity tests with adult freshwater musselsare sum-
marizedinTable 5.4.Specific standardized methods have not been developedfor conducting
toxicity tests with adult mussels, but the proceduresoutlined in Table 5.4 are generally consistent
with guidance for conducting laboratory toxicity tests with early life stages of freshwater mussels
(ASTM 2006a). Exposures have been conducted under static (Keller, Ruessler,and Kernaghan
1999), renewal(Mane 1979; Holwerda and Herwig 1986), and flow-through (Naimo, Waller, and

Holland-Bartels 1992a, 1992b; Imlay 1973; Kernaghan et al. 2003)conditions. Alimited number of
species have been used to conducttoxicity studies with adults, and these musselsare typically
collected from the field.
Adults have been held underlaboratory conditions from one daytoseveralmonths
before thestart of toxicity testing.Toxicity testsare conductedunderawidevariety of
conditions, with exposurechambersranging from 10 to 1,500 L. Duetothe relatively low
abundanceofadultmussels,the numberofreplicates pertestconcentrationand thenumber
of musselstested is generally low. Replication ranges from one to sixchambers,each
containingbetween 9and 125 organisms. The testshavebeenconducted from 48 hours
to 8months, andavarietyofendpointshavebeenusedtoevaluatetoxicity.Survival, as
measured by cessation of siphoning activityand inabilitytoreact to stimulation,isa
common endpoint assessedinmostadult toxicity studies.Inaddition,sublethalendpoints,
such as respiration rate, condition indices, glycogen content, andother biochemical par-
ameters, are frequently measured in toxicity testsconducted with adults.Water quality
analysis,including temperature, dissolvedoxygen, pH,conductivity, hardness,and alkalinity,
areroutinelymeasured duringadult toxicity tests. In addition, bioaccumulationhas alsobeen
determined (e.g., Holwerda andHerwig1986;Naimo, Waller,and Holland-Bartels1992a,
1992b).Controlsurvivalfor adulttoxicity studiesistypicallygreater than 80 to greaterthan
90% at theend of theexposures.
IssuesRegarding the Use of the Methods
Issues regarding the use of adultmussels in toxicity tests are similar to thosefor toxicity tests with
glochidia andjuvenilemussels. Some issues that have been raised in relation to glochidia or
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TABLE 5.4
SummaryofTest Conditionsfor Conducting Toxicity Tests with Adult Freshwater Mussels
Conditions
Keller and
Ruessler (1997)

Mane,Kachole,
and Pawar(1979)
Naimo, Atchison,
and Holland-Bar-
tels (1992b)Imlay(1973)
Raj and Hameed
(1991),Jacobson
(1990)
Holwerda and
Herwig (1986) Farris et al. (1991)
Kernaghan et al.
(2003)
Nicola Kerna-
ghan, University
of Florida, FL
(Unpublished
Data)
Chris Ingersoll,
USGS, Columbia,
MO (Unpublished
Data)
1Species tested Villosaosa lienosa,
Elliptic
icterina,
Utterbackia
imbecillis
a
Indonaia caeruleus Lampsilis
ventricosa
Amblycorypha

carinata,
Lampsilis
radiata
silquoidea,
Fusconaia
flava, Amblema
plicata
Villosa iris,
Actinonaias
pectorosa,
Pyganodon
grandis,
Lampsilis
fasciola,
Medionidus
conradius
Anodonta anatina Elliptio dilatata,
M.conradicus,
Pleurobema
oviforme,
Villosa iris
E. buckleyi E. buckleyiAmblema plicata
2Test type Static Static-renewalFlow-through
diluter
Flow-through Static-renewal Static-renewal Static-renewal Flow-through Static-renewal Flow-through
3Test duration 72–96 hours 48 hours 14, 28 days 36 days-8 months 96 hours, 10, 20,
30 days
7months 30 days 56 days 7, 14, 21, 30,
60 days
4, 56 days

4Temperature ( 8 C) 25, 32 30–32 21 11–21 29 NR 16–20 17 20 20
5Light quality NR
b
NR NR Fluorescent and
incandescent
NR NR NR NR Fluorescent Fluorescent
6Light intensity NR NR Subdued 16–22 foot candles NR NR NR NR NR 250 lux
7Photoperiod 12L:12D NR NR Natural conditions NR NR 10L:14D Natural conditions 16L:8D 16L:8D
8Test chamber 23-L aquaria Plexiglass aquaria 57-L glass aquaria 20-L stainless steel
chamber
Plastic containers NR 75-L fiberglass
oval stream
1500-Lplastic
containers
46-L glassaquaria 40-L glass aquaria
9Test solution
volume (L)
23 NR 57 20 10 40 60 1500 30 25 Lwith a5-cm
layer of gravel
10 Number of
organisms per
test chamber
5–10 50 10 NR 10 50 6125 30 9–10
11 Number of
replicate
chambers per
treatment
2–4 121–6 11112 4
12 Feeding None None Yes None None NR Yes None Yes Algae in pond
water

13 Aeration None None None None None NR None YesYes Yes
(continued)
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TABLE 5.4 (Continued)
Conditions
Keller and
Ruessler (1997)
Mane,Kachole,
and Pawar(1979)
Naimo, Atchison,
and Holland-Bar-
tels (1992b)Imlay(1973)
Raj and Hameed
(1991),Jacobson
(1990)
Holwerda and
Herwig (1986) Farris et al. (1991)
Kernaghan et al.
(2003)
Nicola Kerna-
ghan, University
of Florida, FL
(Unpublished
Data)
Chris Ingersoll,
USGS, Columbia,
MO (Unpublished
Data)

14 Dilution water Reconstituted
water with a
hardness of
76 mg/L as
CaCO
3
River water Reconstituted
water with a
hardness of
165 mg/L as
CaCO
3
Dechlorinated city
water and lake
water
Filtered river water Tap water River water Well water Well water
(hardness
260 mg/L as
CaCO
3
)and
reconstituted
water (hardness
80 mg/L as
CaCO
3
)
Mixture of well
water (hardness
260 mg/L as

CaCO
3
)and
pond water to a
hardness of
about 190 mg/L
as CaCO
3
)
15 Waterquality pH, hardness,
conductivity
Temperature Temperature, DO,
pH, alkalinity,
hardness,
conductivity
Temperature, DO,
pH, hardness,
alkalinity
Temperature,DO,
pH, salinity
DO, pH, Ca, Mg,
Na, Fe,
HCO
3
,Cl, SO
4
Temperature, DO,
pH,
conductivity,
hardness,

alkalinity
Temperature, DO,
pH,
conductivity
Temperature, DO,
pH,
conductivity
Temperature, DO,
pH, ammonia,
conductivity,
hardness,
alkalinity,
16 EndpointSurvival (cessation
of siphoning
activity and
inability to react
to stimulation)
Survival and
effects on
neurosecretory
cells, digestive
gland and
intestine
Respiration rate,
food clearance
rate, ammonia
excretion rate,
assimilation
efficiency,
tissue condition

index, oxygen
to nitrogen ratio
Survival Survival,
respiration rate
and body
weight
Carbohydrate and
lipid content,
lactate,
succinate,
acetate, and
propionate
Survival and
cellulolytic
activity
Survival, body
condition index,
glycogen
concentration,
sex steroid
conentration
Survival,body
condition index,
soft tissue
index, glycogen
concentration,
sex steroid
conentration
Survival, glycogen
concentration,

behavior
17 Control survival
(%)
NR 80 O 80 O 97 NR NR NR 100 NR O 80
a
Formly A. imbecilis .
b
NR, not reported.
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juvenile toxicity tests are alsoapplicable to adult toxicity tests. These concerns mainlyinclude:(1)
the length of time that adults can be held in alaboratory,(2) the conditions for maintaining adults in
the laboratory, (3) the evaluation of the health of adults, and (4) the similarities in sensitivity to
contaminants between different populations of mussels. Thefollowing section attempts to address
some of these concerns. Areas of ongoing research or needs for future researchare also identified.
How LongCan Adults Be Held in the Laboratory?
Dunn andLayzer (1997) reportedthe resultsofseveral long-term holdingexperimentsusing
fisheryponds andraceways.Survivalofadultsvariedbyspecies andaccording to holding
conditions. In theraceway experiments, survival ranged from 43 to 100% afterone year.
Survivalofmusselsheldinpondsappeared more variable,ranging from 0to100%. Other
researchers reportthatadults canbesuccessfully maintained in thelaboratoryfor aperiodof
severalmonths(e.g.,Chris Barnhart,Jerry Farris, Dick Neves, Teresa Newton; personal
communications).
What Conditions Should Be UsedtoMaintain Adults in the Laboratory?
Holding conditions for adults vary by species and season. Temperatures tested rangedbetween10
and 258 C, and musselshave been maintained in systemssupplied by pond, river, or well water.
Conditions that most closelymimic thoseinthe environmentfromwhich themusselswere
collected are recommended. Some researchfacilities have relied upon natural sources of food in
the pondorriver water to maintain an adequate diet for the captive mussels. Researchers at Virginia

Tech have successfully developed acultured algal diet to feed the mussels (Gatenby2000).
How Should the Health of Adults Held in the LaboratoryBeEvaluated?
Adultstobeused in toxicity studies are only occasionally screened for background contamination
levels (Nicola Kernaghan, University of Florida, Gainesville,FL; personal communication). The
health of adults can be evaluated by making observations of activity, behavior,and orientation of
musselsinasubstrate (Jerry Farris and TeresaNewton, personal communications).Further health
assessments may be achieved with the use of biochemical indicators, such as glycogen (Chapter 10).
Several studies have used glycogen as ameasureofthe energetic status of musselsand as an indicator
of their physiological conditionafterexposuretocontaminants (Hemelraad andHerwig1990;
Holopainen and Lamberg 1997; Kernaghan et al. 2003).
Are There Similarities in Sensitivity to Contaminantsbetween Populations of Mussels
of the Same Species?
Some species of mussels appeartobemore sensitive to certaincontaminants than others, and
several studieshavebeenconductedtocompare species sensitivities(Imlay 1973; Keller and
Ruessler 1997). SeeChapter 7for comparison of toxicity endpointsbetweenspecies.However,
no studies have been conducted to date comparingthe sensitivity of different populations of the
same species of mussel.
METHODS FOR CONDUCTING SEDIMENT TOXICITY TESTS
WITH FRESHWATER MUSSELS
R EVIEW OF M ETHODS
Conditions used to conductsediment toxicity tests with freshwater musselsare summarized in
Table 5.5.Chapter 6provides an overview of methods used to conductsediment toxicity tests with
musselsinthe field. Only afew sedimenttoxicity tests with mussels have been conducted, and
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TABLE 5.5
SummaryofTest Conditionsfor Conducting Sediment Toxicity Tests with Freshwater Mussels
Conditions
Keller,Ruessler,and

Chaffee (1998) Wade (1992) Newton et al. (2003)
USGS (2005c), Nile Kemble,
USGS, Coumbia, MO (Unpub-
lishedData)
1Species tested Lampsilis siliquoidea, Lasmigona
costata, Villosa villosa
Utterbackia imbecillis (formerly
Anodonta imbecillis)
Lampsiliscardium Lampsilis siliquoidea, Lampsilis
rafinesqueana
2Test type Renewal Renewal Flow through Renewal
3Test duration (day) 1, 29 4, 10 28
4Temperature ( 8 C) 22 24 21 20
5Light quality NR
a
NR NR Fluorescent lights
6Light intensity NR NR NR 200 lux
7Photoperiod 12L:12D24D 16L:8D 16L:8D
8Test chamber Glass cylinder (5 cm in diameter,
7.5cminheight, closed on one
end, with100 um Nitex screen)
in 5-Lglass aquaria
Glass cylinder (5 cm in diameter,
closed on one end, with 100 um
Nitex screen) in 250-mL
crystallizingdishes
Polycarbonate tube (4.5 cm in
diameter, 11 cm in height,
closed one end with 153 um
mesh Nitex screen)in12

! 8 !
13 cm chamber
300-mLbeaker
9Test solution volume (mL)NR200 800 mL of overlying water, 3cm
of sediment, and1.5 cm of sand
260 mL of overlying water and
15 mL of sieved sediment, two
volume additions/day of
overlyingwater
10 Ageorlife stage of test
organisms
Glochidia and newly-transformed
juveniles
Newly-transformed juveniles Newly-transformed juveniles Two- to four-month-oldjuvenile
mussels
11 Number of organisms per
test chamber
10, 50–100 15 20 10
12 Number of replicate
chambers per treatment
333–6 4
13 Feeding None Yes None Instantalgae mixture
14 Aeration Yes None Yes None
15 Dilution water Well water,with ahardness of
250 mg/L as CaCO
3
Sediment pore water Well water, withahardness of
123–190 mg/L as CaCO
3
Well water, with ahardness of

140 mg/L as CaCO
3
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16 Water quality Temperature, DO, pH,
conductivity
Temperature, DO, pH, hardness,
alkalinity, conductivity
Temperature, DO, pH, hardness,
alkalinity, conductivity,
ammonia
Temperature, DO,pH, hardness,
alkalinity, conductivity,
ammonia
17 Endpoints Survival Survival Survival (foot movement, ciliary
activity), growth(shell height)
Survival (foot movement), growth
(shell length)
18 Control survival (%)Glochidia O 69 Juvenile O 90 96 99–100 90–95
a
NR,not reported.
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specificstandardized methods have not been developed. However, theproceduresoutlinedin
Table 5.5 are generally consistent with the methods for conducting toxicity tests with early life
stagesoffreshwater mussels(ASTM 2006a)and with themethods developedfor conducting
sediment toxicity tests with other freshwater invertebrates (ASTM 2006d). Exposures to whole
sediment and to pore water have been conducted. Both glochidia and juvenile mussels have been

used in sediment toxicity studies. Mussel species for sediment toxicity studies have been selected
accordingtoseveral criteria including:availability of glochidia, suitabilityfor culture in the
laboratory, and similar sensitivity betweenthe surrogate and target species.Glochidia or juvenile
mussels used to start the exposures have been obtained usingproceduresoutlined above in sections
dealing with water-only toxicity tests with glochidiaand with juvenile mussels.
Whole-sedimentexposures with glochidiaornewly-transformed juveniles have been
conducted by placing screenedcylinders containing testorganisms into secondary chambers
containing sediment. Three to sixreplicates perconcentration are generally tested,with up to
100 glochidia or 10–20 newly-transformed juvenile mussels tested per replicate. The duration of
exposures started with glochidia have been 24 and 48 hours,and the duration of exposures starting
with newly-transformedjuvenilemussels have rangedfrom4to 10 days.Controlsurvival of
glochidia ranged from 69 to 79%, and control survival of newly-transformedjuvenilesranged
from 90 to 100% (Table 5.5)
USGS(2005c) and Nile Kemble (USGS, Columbia, MO;unpublished data) conducted a28-day
whole-sedimenttoxicity test starting with two-month-old juvenile L. siliquoidea and L. rafines-
queana.Sediments were sieved to alessthan 250-m mparticle size before the start of the exposure.
The sediments were sieved to obtain aparticle size that couldbeused isolate juvenile musselsatthe
end of asediment exposure. It is unlikely that this life stage of juvenile mussel wouldbeable to
consume larger sedimentparticles. Exposures were conducted in 300-mL beakers containing about
15 mL of sediment and 260 mL of overlying water with about 2volume additions/day of overlying
water.Juvenile mussels were fed an instant algae mixture twice daily (commercial Instant Algae
brand non-viablemicroalgae concentrates; Reed Mariculture, Campbell, CA) and control survival
was greater than or equal to 90%. Studies are ongoingtoevaluatethe influence of sieving sediment
on the bioavailability of contaminants to juvenile mussels and to Hyalella azteca and Chironomus
dilutus (i.e., sieved to aparticle size of 63–250 m m).
IssuesRegarding Use of Methods
Issues regardingthe useofglochidia andjuvenilemussels in sediment toxicity testsare similartothose
forwater-onlytoxicitytests with theselifestages. Theseconcernsinclude:(1) thedurationofthe
toxicity tests, (2)the qualityofthe organismsatthe startofatest,(3) thelifestage tested,(4) the
determinationofdeath at theend of atest, and(5) test acceptabilitycriteria. Many of theseissueshave

alreadybeenaddressed in earliersectionsinthischapter.However, specificissuesrelatingtosediment
toxicity testsare summarized in thefollowing section, andinformation is provided that attempts to
addressthese concerns.Areas of ongoingresearchorneedfor future research arealsoidentified.
Duration of the Toxicity Test
Typically, sediment toxicity tests with glochidia have been conducted for 24–48 hours, and tests
with newly-transformed juvenilemussels have been conducted for up to 10 days. Species-depen-
dent viability of glochidia needs to be considered, as previously discussed(Table 5.2). Sediment
toxicity tests starting with two-month-old juvenile mussels have been conducted for 28 days. In
addition, sediment tests shouldbeconducted for aduration that will enable appropriate compari-
sons to be drawn betweenwater-only and sediment toxicity tests and betweenmussels and other
standard sediment test organisms, such as theamphipod H. azteca andthe midge C. dilutus
(USEPA 2000; ASTM 2006d).
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Life Stage Tested
Glochidia, newly-transformedjuvenilemussels,and two-month-oldjuvenilemussels have been used
in sediment toxicity studies. Adultmussels,which filter-feed as opposedtopedal-feeding juvenile
mussels, have less contactwith thesediment, anditcould thereforebehypothesizedthattheywould be
less sensitivethanthe earlylifestagesofmussels.However,nostudies of sediment toxicity on adult
musselswerereviewedfor this chapter, andcomparisons betweenlifestageshavenot been conducted.
Use of Control or Reference Sediments to Establish Test Acceptability
Typically, control sediment is asediment that has previously been demonstrated to have no toxic
effects on the species being tested and is used to evaluatethe acceptability of atoxicity test(USEPA
2000; ASTM 2006d). In some instances, control sediments may have been sterilized or tested for
contaminants before the start of asedimenttoxicity test(Keller,Ruessler, and Chaffee 1998).
Sediment material collectedfromareferencesite, which is usually arelatively undisturbed
location, has been used to evaluatetest acceptability and to makecomparisons to test sediments
(USEPA 2000; ASTM 2006d). Survival of musselsincontrol or reference sediments has been
reportedinthe literature to be acceptable at greater than 90% (Table 5.5); however, no standard

methods for conducting toxicity tests with freshwater musselshave been developed to establish a
definitive testacceptability criterion for survival.
METHODS FOR CONDUCTING HOST FISH EXPOSURE TOXICITY
TESTS WITH FRESHWATER MUSSELS
R EVIEW OF M ETHODS
Two studies have evaluated exposure of glochidia encysted on fish to toxicants (Jacobson 1990;
Nicola Kernaghan, unpublished data). During the glochidial stage of development, mussels attach
to the gills of ahost fish and encyst in host tissues within 2–36 hours of attachment. While encysted,
the glochidia change form and begintoresemble adults. Although the exact processesthat occur
while the glochidia are attached to the host are not fully understood, some in vitro experiments
suggest that glochidiaabsorborganic molecules from fishtissues and requirefish plasma for
development andmetamorphosis (Isom and Hudson 1982) in atruehost–parasiterelationship.
Glochidia remainattached to the host fish from 7to10days to several months, offering asignificant
period of time during which they may be exposedtohost contaminant burdens.
NicolaKernaghan collected gravid femalemussels from areference location that has histori-
callysupported healthy populations of diverse mussel species.Hostfish were collectedfrom
captivepond populations forwhich contaminantbodyburdens were documented. These fish
were implantedwith time-release pellets of toxaphene,dichlorodiphenyldichloroethylene (DDE),
or atrazine to establish body burdens representativeofconcentrations found in fish tissue in the
environment. Three to five fish were prepared at each concentration, and an equal number of control
fish were implanted with placebo pelletsfor each test.
Following inoculationwithglochidia,fish were held separately in 40-Laquariauntil
glochidial transformation occurred.Juvenile musselswerecollected andrandomly assigned
to smallglass Petri dishes containing about20mLofwater.Fivereplicates,eachcontaining
at least10juvenile mussels, were used foreachimplanted fishand test durationwas nine
days.Aspreviously described forjuvenile water-only andsedimenttoxicitytests,cessation
of both activityand heartbeat wasused as themeasurementendpointindicating death.
Controlsurvivalofglochidia forhostexposuretoxicity testswas generally greater than
80% (NicolaKernaghan, unpublisheddata).
Jacobson(1990) investigated theeffectofcopperonthe early life stagesoffreshwater

mussels,including theencystedglochidia of V. nebulosa , A. pectorosa ,and A. grandis.
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Largemouth bass,which were used as hosts, were obtained from afish hatchery andwere
held in thelaboratoryfor five to sevendaysbeforetesting.Fishwereencysted en masse in a
rectangular 120-Lpolyethylenetank, filledwithabout 40–60L of aerated, dechlorinated tap
water. Glochidiaofaspecies were then addedtothe tank, resulting in arelativelyhom-
ogenous levelofencystment on allfish.The infested fishwereexposed to 0, 25, 100,and
200 m gCu/Lin17-Lpolycarbonate carboys at 19–218 Cand with aphotoperiod of 16:8,
light–dark.Three replicates,each containing 5or10encystedfish were prepared foreach
concentrationand test solutionswererenewed everytwo to four days.Ateachrenewal,the
tank waterwas drainedthrough a100-m msieve,and thenumberofjuvenile musselswas
recorded.Meannumberofjuvenile musselstransformed perfish wasusedasanendpointfor
this study.
IssuesRegarding the Use of Methods
Issues regarding host fish exposure studies are uniquetofreshwater mussels, as aresult of the
unique life cycleoffreshwatermussels.However, issues raised regardingstandardization of
methods for conducting toxicity tests with glochidia or juvenile life stages of freshwater mussel
described above are also applied to encystment tests. These concerns include:(1) the duration of the
toxicity tests, (2) the confirmation of exposure concentrations, (3) determination of mortality of
glochidia while attached to the host,(4) evaluating exposure of glochidia to contaminants while still
in the marsupium of the adult female mussel, and (5) testacceptability criteria. These specific issues
relating to host fish exposure studies are summarized in the following sectionand information is
provided that attemptstoaddress theseconcerns. Areas of ongoing research or need for future
researchare also identified.
Duration of the Toxicity Test
The length of host fish studies is primarily determined by the period of time that anygiven species will
naturally remain encysted on the host fish. Subsequent survival studies of juvenile musselsshouldbe
based upon the recommendations for conducting water-only toxicity studies with juvenile mussels.

Confirmation of Chemical Concentrations in Host Fish and Exposed Mussels
Due to the relatively small sizeofglochidia and juvenile mussels, it is generally not practical to
determinebioaccumulation of chemicals. Testingofhostfish to determineindirectexposure
concentrations would be useful, but further studies are neededonthe relation between the host
fish and the encysted glochidia to understand the significance of these concentrations.
Determination of Glochidia Mortality While Attached to aContaminatedHost Fish
Jacobson (1990) enumeratedall juvenile mussels transformed in each replicateand calculated a
mean number of transformed juvenile mussels per fish. Alternatively, NicolaKernaghan tested a
subsample of transformed juvenilemussels. There is no reliable method of determining glochidia
mortality while attached to acontaminated fish, requiring further research.
Would Glochidia, While Still in the Marsupium of the Adult Female Mussel,
Be Exposed to Waterborne Contaminants?
The brood chambers of adult mussels and the glochidia that they contain have been reported to be
physically isolated from the general water circulation pattern in the rest of the gills of the adult
mussels (Silverman, McNeil, and Dietz 1987). In addition, exposure of brooding adults or encysted
glochidia to copper were not very sensitive, suggesting that the glochidia are functionally isolated
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from the ambient water conditions while they are in the brood chamber (Jacobson 1990). However,
more researchisneededtofurther evaluate this question.
METHODS FOR CONDUCTING TOXICITY TESTS USING CORBICULA
FLUMINEA AS SURROGATE SPECIES
The Asian clam, C. fluminea,has been frequently used as asurrogate for otherfreshwater mussels
and target test organism since the early 1970s, more than 30 years following their introduction into
the UnitedStates. Theuse of Corbicula in toxicity testinghas supported the hypothesis that this
organism is aviable indicator of impairment in aquatic systems (Graney, Cherry, and Cairns 1983).
Consequently, thresholdresponses of Corbicula have been used to assess relative impacts on native
mussels(Moulton, Fleming, and Purnell 1996; Milam and Farris 1998). Areview by Doherty and
Cherry(1988) provides an assessment of measured effects on juvenile and adult C. fluminea in

laboratorytests includingadescription of various exposure andrecovery regimeswithsingle
chemicals in static or flow-through systems. More recent toxicity tests have includedacute and
chronic exposures with Corbicula using: (1) laboratory and in situ techniques; (2) multi-contami-
nant stressors, including metals, surfactants, pesticides, and industrial or municipal effluents; and
(3) measurement of alternative endpointstoassess damage in exposed populations. The purposeof
this sectionistoinclude additional data available since the review by Doherty and Cherry (1988)
and to evaluatevariability among responses observed in toxicity tests conducted with C. fluminea.
R EVIEW OF M ETHODS
Specificstandard methods for conducting toxicity tests with Corbicula have not been developed.
However, standardmethods have been published for conducting bioconcentration tests (ASTM
2006e)and acute toxicity tests with saltwater bivalves(ASTM 2006c)that could be adapted for use
with Corbicula.Additionally, standard methods have been developed for in situ testing specifically
using caged bivalves(primarily, estuarine and marine species) (ASTM 2005f). Techniques have
been developedtominimize variability amongexposures when measuring growth and
tissue bioaccumulation.
Asummary of toxicity test methods using C. fluminea sincethe completion of the review by
Dohertyand Cherry(1989) is presented in Table 5.6.Experimental designs for these studies include
exposures to single-chemical or complexeffluentsbymeasuring avariety of toxic endpoints. Test
methods compiled from this dataset are either derivations from USEPAguidance documents for
acute andchronic testing (USEPA 1993)orsite-specificmethods developedspecificallyfor
bivalves(Farris et al. 1989; Belanger et al. 1991).
Asubstantial portion of the studies outlined in Table 5.6 tested adult Corbicula rather than
juvenile life stages, perhaps because of the range of lethal and sublethal endpoints that can be
measured with adultorganisms (i.e., sublethal test endpoints have typicallybeen developed using
greater tissue mass than whatisavailable in juvenile mussels). Laboratorytoxicity tests that were
conducted with juvenile and adult Corbicula ranged in duration from 4(Moulton,Fleming, and
Purnell 1996)to56days (Belanger, Meiers, and Bausch. 1995; Belanger et al. 2000). In situ tests
ranged in duration from 31 (Soucek et al. 2000)to90days (Bouldin, Farris, and Milam 2003).
Although various test chambershave been used, mostofthe studies used artificial streamsina
laboratory. Artificial streams are flexible in design and have the ability to hold large volumes of

water with varying depths and currentsfor conducting tests with Corbicula.
Aqueous Toxicity Testing
Many of the tests were conducted in the laboratory usingawide range of feeding regimes and
photoperiods (Table 5.6).These tests included wholeeffluent fromindustrial andmunicipal
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© 2007 by the Society of Environmental Toxicology and Chemistry (SETAC)