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THESIS

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LABORATORY AND FIELD ANALYSES OF THE
POTENTIAL TOXICITY OF POLYDIMETHYLSILOXANE (SILICONE)
TO BENTHIC MACROINVERTEBRATES

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Presented by

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11m chIRCIDatoOmuS-p.“

 

 

LABORATORY AND FIELD ANALYSES OF THE POTENTIAL TOXICITY OF
POLYDIMETHYLSILOXANE (SILICONE) TO
BENTHIC MACROINVERTEBRATES

By

Kevin S. Henry

A DISSERTATION

Submitted to
Michigan State University
in partial fulﬁllment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Department of Zoology and
Institute of Environmental Toxicology

1999

ABSTRACT

LABORATORY AND FIELD ANALYSES OF THE POTENTIAL TOXICITY OF
POLYDIMETHYLSILOXANE TO BENTHIC MACROINVERTEBRATES

By
Kevin S. Henry

Polydimethylsiloxane (PDMS) is widely used in a number of industrial processes
and consumer products that result in down-the-drain disposal. Negligible volatility and
water solubility and a slow degradation rate suggest that PDMS may accumulate in
sediments. Sediment bioassays with the amphipod Hyalella azteca and with larvae of the
midge Chironomus tentans were used to assess the potential for toxicity of PDMS-
amended sediments to benthic invertebrates in short-term and whole life cycle exposures.
Endpoints for short-term tests included survival and growth, while life cycle assays
considered survival, growth, reproduction, and emergence. Pctential effects of PDMS on
benthic communities were investigated in situ in plastic trays containing sediments that
had been defaunated and spiked with PDMS to concentrations of 1000 ppm (nominal
d.w.). Artiﬁcial substrate trays were placed at a depth of 10 m in Lake Orta, Italy, a lake
which has been historically contaminated with ammonia and heavy metals. Trays were
collected after 3 mo and invertebrates sieved and identiﬁed to genus or species, when
possible. Short-term and life cycle laboratory exposures indicated that environmentally
relevant concentrations of PDMS will not signiﬁcantly decrease survival, growth, or
reproduction. In situ, artiﬁcial substrate exposures exhibited a high degree of variability
among treatments, and suggested possible impacts on oligochaetes. However, this
observation was not substantiated in laboratory exposures with the oligochaete

Lumbriculus variegatus.

Copyright by
Kevin Shawn Henry
1999

Dedicated to my Mother, Jean Henry,
for encouraging my interest in every reasonable pursuit.

iv

 

ACKNOWLEDGEMENTS

Financial support for the research reported in this manuscript was provided by the
United States Department of Agriculture Water Sciences Fellowship Program, and Dow
Corning Corporation. My MSU graduate committee, which consisted of my mentor, John
Giesy, and Richard Merritt, Steven Hamilton, and Frank D'Itri, provided valuable advice and
assistance and helpful review and comment on the ﬁrst draft of this thesis. Many
contributions to the development of this research project were provided by David E. Powell
and Ronald B. Annelin of Dow Corning Corporation. I thank Michael Kaufman at Kellogg
Biological Station for his logistical and technical contributions to this research. Assistance
with invertebrate cultures and countless laboratory tasks was provided by Rebecca Graving,
Lancie Dole, Brian Nagy, Margaret Mackercher, Joyce Lau, and Christopher Chapman. I
owe special thanks to my associates at Michigan State University, especially to my
colleagues and friends Shane Snyder, Erin Snyder, Willemien Wieland, Jong-Seong Khim,
Becky Blasius, K. Kannan, Thomas Sanderson, Vince Kramer, John Besser, David
Verbrugge, and Lori Feyk, for technical and moral support. Peter Landrum at NOAA/Great
Lakes Environmental Laboratory in Ann Arbor, Michigan provided sediments and much
valuable advice on invertebrate sampling and ecology. Renato Baudo, Annamaria Nocentini,
Daria Rossi, and Monika Beltrami of the Italian Institute for Hydrobiology were responsible
for helping organize and conduct ﬁeld studies. Finally, thanks to Cliffard Dahm, Michael
Campana, John Morrice and Greg Wroblicky of the University of New Mexico for initial
encouragement, and to H. Maury Valett of Virginia Polytechnic Institute for "making my

career."

TABLE OF CONTENTS

List of Tables ........................................................................................................... vii
List of Figures ........................................................................................................ viii
Introduction ............................................................................................................... 1
Materials and Methods ............................................................................................... 3
Test Organisms and Sediments ............................................................................... 3
PDMS Fluid ........................................................................................................... 4
Sediment Dosing and Analyses ............................................................................... 5
Short-Term Laboratory Exposures .......................................................................... 5
Whole Life Cycle Assays ....................................................................................... 7
C. tentans Whole Life Cycle Assay ........................................................................ 7
H. azteca Whole Life Cycle Assay ........................................................................ 10
In Situ Bioassays .................................................................................................. 11
Statistics ............................................................................................................... 12
Results and Discussion ............................................................................................ 12
Short-Term Laboratory Exposures ........................................................................ 12
Whole Life Cycle Assays ..................................................................................... 14
In Situ Bioassays .................................................................................................. 16
Literature Cited ........................................................................................................ 40
Appendices .............................................................................................................. 43
Appendix 1: Standard Procedure for Polydimethylsiloxane (PDMS) Extraction
from Sediment ...................................................................................................... 44
Appendix 2: Standard procedure for a Short-Term Growth and Survival Assay with
the Midge Chironomus tentans ............................................................................. 48
Appendix 3: Standard Procedure for a Short-Term Growth and Survival Assay
with the Amphipod Hyalella azteca ...................................................................... 59
Appendix 4: Standard Procedure for a Whole Life Cycle Assay with the Midge
Chironomus tentans .............................................................................................. 69
Appendix 5: Standard Procedure for a Whole Life Cycle Assay with the Amphipod
Hyalella azteca ..................................................................................................... 89

vi

LIST OF TABLES

Table 1. Effects of polydimethylsiloxane on benthic invertebrates. ............................... 19

vii

 

 

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LIST OF FIGURES

Figure 1. Generic structure of PDMS (Stark et a1. 1982). .............................................. 20

Figure 2. Responses in Chironomus tentans bioassays with PDMS-spiked
sediments: Trial 1 of 2. Means, with standard error (N = 16). * indicates
statistically significant difference at or = 0.05 using Dunnett’s test for differences
between each treatment and a control. (a) growth measured as dry weight per
individual; (b) growth measured as ash-free dry weight per individual. .......................... 21

Figure 3. Responses in Chironomus tentans bioassays with PDMS-spiked
sediments: Trial 2 of 2. Means, with standard error (N = 16). * indicates
statistically significant difference at or = 0.05 using Dunnett’s test for differences
between each treatment and a control. (a) growth measured as dry weight per
individual; (b) growth measured as ash-free dry weight per individual. .......................... 23

Figure 4. Responses in Hyalella azteca bioassay with PDMS-spiked sediments.
Means, with standard error (N = 16). (a) grth measured as dry weight per
individual; (b) growth measured as ash-free dry weight per individual. .......................... 25

Figure 5. Responses of Chironomus tentans and Hyalella azteca with PDMS-
spiked sediments of low organic carbon content (0.5% OC). Means, with standard
error. (a) percent survival of Chironomus tentans (N = 16). (b) growth of
Chironomus tentans measured as dry weight per individual (N = 16). (c) percent
survival of Hyalella azteca (N = 12). ((1) growth of Hyalella azteca measured as
dry weight per individual (N = 12). ................................................................................ 26

Figure 6. Responses in Chironomus tentans whole-life cycle bioassays with
PDMS-spiked sediments. Dashed lines indicate suggested USEPA data quality
objectives. (a) percent survival at 20 d (N = 4). (b) Ash-free dry weight at 20 d
(N = 4) ........................................................................................................................... 28

Figure 7. Responses in Chironomus tentans whole-life cycle bioassays with
PDMS-spiked sediments. Dashed lines indicate suggested USEPA data quality
objectives. (a) C. tentans percent emergence (N = 8). (b) reproduction in number
of eggs/female. (c) percent of eggs hatching ................................................................. 30

Figure 8. Responses in Hyalella azteca whole-life cycle bioassays with PDMS-
spiked sediments. Dashed lines (when present) indicate suggested USEPA data
quality objectives. (a) percent survival at 28 d when animals are isolated from
sediments and placed into water-only chambers (N = 12). (b) Percent survival at
35 d (N: 8). (c) Percent survival at 42 (1 (test termination, N = 8). ............................... 32

viii

Figure 9. Responses in Hyalella azteca whole-life cycle bioassays with PDMS-
spiked sediments. Dashed lines (when present) indicate suggested USEPA data
quality objectives. (a) growth at 28 (1 when animals are isolated from sediments
and placed into water-only chambers (N = 4). (b) growth at 42 (1 (test termination,
N = 8). (c) reproduction at 42 d evaluated in number of young produced per
female ............................................................................................................................ 34

Figure 10. Results of in situ study with silicone-spiked sediments. (a) histogram of
oligochaetes and chironomids in artiﬁcial substrates. (b) percent of chironomid
predators in artiﬁcial substrates ...................................................................................... 36

Figure 11. Responses in Lumbriculus variegatus bioassays with PDMS-spiked
sediments. Means, with standard error (N = 4). (a) percent survival. (b) grth
measured as mean wet weight per individual .................................................................. 38

INTRODUCTION

Polydimethylsiloxane (PDMS) ﬂuids are synthetic polymers consisting of an
alternating silicon-oxygen backbone with methyl side groups (Figure 1). PDMS is used
in a variety of industrial and consumer applications, including heat transfer ﬂuids,
antifoams, cosmetics, waxes and polishes (Hamelink 1992). Many of these uses result in
down-the-drain disposal. The most common linear siloxanes are insoluble in water and
extremely hydrophobic, therefore, siloxanes become associated with sludge solids upon
reaching sewage conduits and waste water treatment plants (WWTPs, Watts et a1. 1995;
Fendinger et a1. 1997).

Approximately 3% of the PDMS delivered to WWTPs has been estimated to be
released to surface waters in the United States (Fendinger et al. 1997). This represents an
total annual loading of approximately 294,000 kg/y (Fendinger et a]. 1997). Upon
entering surface waters, the majority of the suspended sludge and associated PDMS will
eventually deposit into bottom sediments. The small fraction of PDMS that is not sorbed
to suspended particulates in the efﬂuent would partition to suspended solids in the river
or lake water and also eventually become deposited into the bottom sediments (Fendinger
et a1. 1997).

Degradation of PDMS is believed to occur by hydrolysis to yield monomeric
dimethylsilanediol (DMSD, Watts et a1. 1995), though this reaction is inversely related to
soil moisture content (Lehmann et a1. 1998). In one study, a year of incubation in aerated
sediments resulted in hydrolysis of 5-10% of PDMS present to DMSD (Carpenter et a1.
1996). The rate of PDMS degradation was less in sterile sediments, which suggests that

this process may be related to sediment microﬂora (Carpenter et a1. 1996). The mobility

 

 

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of PDMS in aquatic systems will be primarily limited to resuspension and deposition of
sediment particles with which it is associated. Concentrations of PDMS in sediments
decrease with depth, and are not detectable at depths which correspond to periods before
the production and use of silicone ﬂuids (Batley and Hayes 1991; Pellenberg and Tevault
1986). While PDMS tends to accumulate in sediments, concentrations can decrease
when inputs are stopped, by dilution and degradation (Carpenter et al. 1996). PDMS
concentrations in sediments downstream from WWTPs have been observed to range from
0.9 to 78 mg/kg, dw at sites receiving point source inputs. However, 90% of the
sediments tested contained less than 26 mg PDMS/kg, dw (Powell et al. 1996). The biota
exposed to PDMS would primarily be benthic invertebrates intimately associated with
these sediments.

The reported toxicity of PDMS to benthic invertebrates is low. In part, this
nominal level of toxicity is due to the fact that there is little bioaccumulation (Table 1). It
has been suggested that this low bioaccumulation potential is due to the large molecular
weight of PDMS molecules, which are believed to be too large to cross biological
membranes (Frye 1983). While PDMS has been found to be nontoxic in short-term
studies, little information was available on the probable chronic toxicity of PDMS to
sediment dwelling benthic invertebrates. PDMS is surface active so it was hypothesized
that large concentrations in sediments could change the physical characteristics of the
sediments. For instance, it has been reported that PDMS can reduce the bioavailability of
organic contaminants such as polycyclic aromatic hydrocarbons (PAHs) from sediments
(Kukkonen and Landmm 1995). Because a number of benthic invertebrates extract

organic compounds from sediments, either as an energy source or in the form of fat

soluble vitamins, it was postulated that PDMS might affect the availability of the organic
compounds by acting as a cosolvent. Finally, because PDMS may affect the texture of
sediments with respect to their viscosity or degree of consolidation, there was concern
that PDMS-contamination of sediments could change the suitability of the sediments for
benthic invertebrates. If the degree of consolidation of the sediment is altered by PDMS,
bioturbation and the production of burrows may be adversely impacted.

The objectives of this study were to determine if sediment-bound PDMS will
affect the survival, growth, or reproduction of benthic invertebrates. This was
investigated by use of ﬁeld studies of mixed populations of naturally occurring
invertebrates and controlled laboratory bioassays with several different species. Test
methodologies used included standardized short-term bioassays as well as multiple life-
stage protocols which are currently under development by the USEPA and USFWS.

MATERIALS AND METHODS
Test Organisms and Sediment

Three species of invertebrates were used in laboratory exposures: an amphipod
(Hyalella azteca, Crustacea) a midge (Chironomus tentans, Insecta) and an aquatic worm
(Lumbriculus variegatus, Oligochaeta). All of these species demonstrate characteristics
of ideal test species (Giesy and Hoke 1989). Standard test methods using these
organisms exist, and survival and growth are the primary endpoints (USEPA 1994;
ASTM 1995). Therefore, these organisms were used as surrogates for insects, crustacea
and oligochaetes in natural communities. The three organisms selected represent a range

of possible tolerances and life histories.

All three species were reared and maintained according to methods adapted from
Environment Canada (1997a, b), Kubitz et al. (1996), and USEPA (1994). Brieﬂy, H.
azteca were cultured at 23il°C in l L plastic breeding chambers kept in an incubator.
Nylon bolting cloth was used as a substratum Amphipods were fed daily 3 suspension of
yeast, cereal leaves, and trout chow (i.e. YCI‘), as well as the green alga Selanastrum
capricornutum. Partial water replacements were performed two times per week. C.
tentans and L variegatus were cultured in 19 L aerated glass aquaria to which shredded
and presoaked unbleached paper towels were added as a substratum. C. tentans were fed
a suspension of blended ﬁsh food ﬂakes (Tetramin), and L. variegatus were fed a
suspension of ground trout chow daily. Water for all cultures consisted of a mixture
containing 30% well water and 70% Barnstead E-Pure (Dubuque, IA), resulting in a ﬁnal
hardness of approximately 90 mg CaC03/L and a ﬁnal alkalinity of approximately 100
mg CaC03/L.

Test sediment for all lab studies except those conducted with “low organic
carbon” sediment was collected from a small pond in Williamston, Michigan, which
receives no regular surface water inputs. Low organic carbon sediments from Lake
Michigan were obtained from Peter Landrum at the NOAA/Great Lakes Environmental
Research Laboratory, Ann Arbor, Michigan. Sediment from Williamston contained 4%
organic carbon (OC), while that from Lake Michigan contained 0.5% DC. Sediments
were stored in high-density polyethylene buckets in the dark at 4°C until use.

PDMS Fluid
Polydimethylsiloxane ﬂuid with a viscosity of 350 cs was obtained from Dow

Corning Corporation (Midland, MI). The molecular weight of this ﬂuid was 17970 and

the log K0“, was approximately 10. Vapor pressure and water solubility are below
detection limits for PDMS of this viscosity.
Sediment Dosing and Analyses

Sediment spiking methodologies were designed to be as environmentally realistic
as possible. PDMS-amended treatments were prepared by spinning sludge, PDMS and
water in a round-bottomed ﬂask on a Rotavap in order to mix these components. This
spiked sludge was then stirred into homogenized sediments using a drill-mounted
stainless steel propeller. The sediment/sludge mixture was shaken on a platform shaker,
then permitted to equilibrate undisturbed for at least 7 d at 4°C before it was
homogenized again for use in bioassays.

Concentrations of PDMS in the sediments were determined on subsamples.
Dosed wet sediment samples were taken in duplicate or triplicate for PDMS analyses.
PDMS concentrations were determined by multiple extraction of sediment subsamples
with tetrahydrofuran (THF), followed by ICP analyses of the extracts (Fendinger et al.
1997b; Appendix 1).
Short-Term Laboratory Exposures

Short-term sediment bioassays were conducted using procedures similar to those
developed by the U.S. Environmental Protection Agency (USEPA 1994) and
Environment Canada (1997a, b). Standard operating procedures are attached (Appendix
2 and 3). Bioassays with C. tentans were conducted on cohorts reared from eggs
deposited on the same date. Larvae of uniform age (13 d post-oviposition) and uniform
size were selected for use in bioassays. Bioassays with H. azteca were performed with 7-

8 day old organisms. Tests with L. variegatus were initiated with adults.

Bioassays were conducted in a Benoit system (Benoit et al. 1982). Exposure
chambers (4-16 per replicate) were prepared from 300 mL high-form borosilicate glass
beakers with two 17 mm holes, covered with 100 um stainless steel screen, to allow
water circulation. Bioassay chambers were ﬁlled with 100 mL of test sediment and 175
mL of overlying water and placed into the exposure aquarium one day before the start of
a test. One volume of dilution water was added to each aquarium before the start of a
bioassay and the exposure chamber water was maintained at 23i1°C over the course of a
test. Test animals were added to the chambers at random to produce a total of ten
animals per chamber. Each group of replicates was placed in a 19 L glass aquarium, into
which water replacements were added by gravity from a polyethylene head tank. The
water replacement rate was two volumes per day.

C. tentans and H. azteca were fed suspensions of Tetramin or YCT, respectively,
each day. L. variegatus were not fed during a test. Feeding and water renewals
continued in this fashion until termination of the bioassay after 10 d. Dissolved oxygen
(DO) was measured in two replicates of each treatment daily before water replacement.
If persistent reductions in D0 (<40% saturation) were detected, the number of water
exchanges per day was increased. If necessary, feeding rates were also reduced
temporarily in all treatments. Composite samples of overlying water for water quality
analysis were collected on test days 0 and 10.

At the end of a bioassay, exposure chambers were removed from the aquaria and
the contents of each chamber were sieved through a 425 um (H. azteca and L. variegatus)
or 710 um (C. tentans) sieve and rinsed with culture water. The number of survivors in

each chamber was recorded and the weight of surviving C. tentans and L. variegatus

individuals determined to calculate growth. C. tentans individuals were pooled by
replicate in pre-ashed aluminum weigh boats and dried for 24 h at 60°C to determine dry
weight. Ash-free dry weight (AFDW) was also estimated to reduce the bias introduced
by gut contents (Sibley et al. 1997). To ﬁnd AFDW, the weigh pans containing the dry
larvae were ashed at 550°C for 2h. The pans with the ashed larvae were then re-weighed
and the tissue mass of the larvae determined as the difference between the weight of the
dried larvae plus pan and the weight of the ashed larvae plus pan. L. variegatus were
pooled by replicate and wet biomass determined. Oligochaetes were not dried because
they were later used in supplemental experiments which required wet biomass and which
are not described here.
Whole Life Cycle Assays

Whole life cycle assays with C. tentans and H. azteca were conducted based on
draft protocols developed by the USEPA and during a series of round-robin tests in which
our laboratory participated. Standard operating procedures are attached (Appendix 4 and
5). Minimum data quality objectives for survival, growth and reproduction have been
suggested (USEPA 1999). These values are given below where available.
C. tentans Whole Life Cycle Assay

The life cycle test for C. tentans was modified from Benoit et al. (1997) and
USEPA (1999). Bioassay chambers were prepared as described above under short—term
laboratory exposures. Specialized equipment specific to the midge whole life stage assay
is described in the standard operating procedure below (Appendix 4). Brieﬂy, adult
midges are placed into a 500 mL Erhlenmeyer ﬂask for mating. The following day, 6 - 8

of the larger C-shaped egg masses are transferred to a glass crystallizing dish containing

culture water and incubated at 23 i 1°C. C. tentans eggs incubated at this temperature
begin hatching after 48 h and the larvae start leaving the egg mass 24 h after ﬁrst hatch
(72 h total). To start a test (day 0 of test), newly hatched larvae are collected with a
Pasteur pipette from the bottom of the incubation dish with the aid of a dissecting
microscope. Twelve test organisms were pipetted directly into the overlying water of
each exposure chamber. Each midge test beaker was fed 1.0 mL of a 4 g/L Tetramin
suspension daily starting on day -1.

A total of 12 beakers per replicate were used in the test, and an additional 4
beakers were prepared and midge larvae introduced on day 10 for production of auxiliary
males to be used in reproduction. These beakers are necessary because peak female
emergence may occur up to 1 week later than that of male C. tentans (Sadler 1935). The
additional replicates ensure a sufficient supply of reproductively viable males for mating
with females from the experimental treatments during the latter stages of the emergence
period.

On day 20 of the test, the sediment from four randomly selected beakers per
treatment was sieved through a 40 mesh (425 um) screen to recover larvae for growth
and survival determinations. Growth was measured as ash-free dry weight. Data quality
objectives for mean control survival and for mean AFDW of 70% and 0.48 mg/control
individual have been suggested (USEPA 1999). After 20 d, emergence traps (screened
lids) were placed over the remaining eight reproductive replicates. Emergence traps for
the auxiliary male beakers are added at the corresponding 20 d time interval.

Male/female emergence and adult mortality in the test chambers were recorded

daily. Adults were collected daily from each individual test chamber and transferred to

the corresponding reproduction/oviposit (R/O) chamber for breeding. Each R/O unit was
checked daily for dead adults and for egg cases. Dead ﬂies were removed. For each
emerged female, at least one male derived from the corresponding reproductive replicate,
another replicate of that treatment, or from the auxiliary male beakers was transferred
into the RIO unit. A draft quality control value for percent emergence of 60% has been
suggested (USEPA 1999).

Egg masses from the RID chambers were transferred to incubation dishes and the
number of eggs determined either by estimation using a ring count method or by direct
counting after digestion in an acid solution. The ring count method involves multiplying
the mean number of eggs contained in 5 rings of the egg mass by the number of rings
present. The acid method is used when the integrity of an egg mass precludes estimation
by the ring method, such as when the mass is convoluted or distorted. Under these
circumstances, egg masses were soaked overnight in sulfuric acid. The acid dissolves the
gelatinous matrix surrounding the eggs and permits counting of individual eggs. The ring
count method is preferable as it is faster and permits determination of egg hatching
success. Each intact egg mass counted by the ring method was incubated in culture water
and incubated at 23°C until hatching was complete, which normally takes between 2 and
6 d. Hatching success was determined by subtracting the number of unhatched eggs
remaining from the number of eggs originally estimated for that egg mass. Draft quality
assurance objectives of 900 eggs per female and 90% hatch have been suggested by the
USEPA (1999).

The physicochemical properties of the sediments determine the length of this test.

In nontoxic sediments, the test typically requires 40 to 50 d for completion. However,

test duration increases in the presence of stressors which act to reduce growth and delay
emergence. The test was terminated when there was no emergence from the control
treatments for 7 d. At test termination, all beakers were sieved through a #40 mesh
screen (425 pm) to recover remaining larvae, pupae, or pupal cases. The endpoints for
the test were thus 20 (1 growth and survival, emergence, reproduction in the number of
eggs per female, and percent hatch.
H. azteca Whole Life Cycle Assay

H. azteca used in the whole life cycle assay were 7 to 8 days old. To start a test,
amphipods of known age were collected with a Pasteur pipette from the bottom of a
crystallizing dish with the aid of a dissecting microscope. A total of 12 test beakers were
set up for each treatment and test organisms pipetted directly into the overlying water of
the exposure beakers. Each beaker received a daily addition of 1.0 mL of YCI‘ starting
on day —1 , which is the day preceding test initiation.

Endpoints monitored included 28d survival and growth of H. azteca and 35 d and
42 d survival, growth, and reproduction (number of young/female). On day 28 of the
test, 4 of the replicate beakers/sediment were separated with a #40 mesh sieve (425-um
mesh) to remove surviving H. azteca. Immobile organisms isolated from the sediment
surface or from sieved material were considered dead. Dry weights were found at 20 d
for surviving amphipods in these 4 replicates. Control survival and dry weight data
quality objectives of 80% and 0.15 mg/individual have been suggested (USEPA 1999).

After 28 d, the remaining 8 of the original 12 beakers/sediment were also sieved

and the surviving H. azteca in each beaker placed into 300 mL beakers containing 250

10

mL of overlying water and a 5-cm x 5-cm piece of Nitex® screen. Each water-only
beaker received 1.0 mL of YCT stock solution and two volume additions of water daily.

Reproduction of amphipods was measured after 35 d and 42 d in the water-only
beakers by removing and counting the adults and young in each beaker. On day 35, the
adults were returned to the same water-only beakers. Adult amphipods surviving at 42 d
were preserved in an 8% sugar formalin solution. The number of adult males was
enumerated based on the size of the second gnathopod, which is enlarged in male H.
azteca. The number of surviving females was used to determine the number of
young/female/beaker produced between 28 d and 42 d. A data quality objective of 2
young/female has been suggested (USEPA 1999). Dry weight for these surviving H.
azteca were also be measured.
In Situ Bioassays

In situ bioassays were conducted in Lake Orta, Italy. This alpine lake has
received heavy metal- and ammonium-containing industrial discharges since 1926
(Mosello and Calderoni 1990). The effects of this contamination on lake biota were
almost immediate and the lake was relatively devoid of fauna by the end of 1927. Since
then, efﬂuent treatment and liming of the lake have resulted in a partial recovery
(Mosello and Calderoni 1990). Lake Orta sediments were collected and a portion of the
sediments amended with PDMS-treated sludge using the spiking method described
above. Sediments of three PDMS treatment levels (10, 100, and 1000 mg PDMS/kg, dw
nominal) were generated, in addition to a sludge blank (0 mg PDMS/kg, dw nominal),
and a control blank which contained neither sludge nor PDMS. The treated and

untreated sediments were placed into 14 L plastic trays and positioned on the lake bed in

11

10 m of water with a method adapted from Hare et al. (1995a, b). One tray of each
sediment concentration was recovered after three months, and sieved using a plankton net
to insure retention of early instar midges. The invertebrates present were then identiﬁed
to the greatest degree possible, generally genus or species.
Statistics

Mean treatment effects of PDMS were examined statistically by a combination of
parametric and nonparametric methods. Percent data were arcsin square root transformed
before testing. All data were assessed for homogeneity of variance using Bartlett's test
and for normality by Shapiro-Wilk's test. If the data met these assumptions, parametric
statistics were used. If one or more assumptions were not met, nonparametric statistics
were used. Data were evaluated by AN OVA followed by Dunnett’s test for pairwise
comparison between each treatment and the control means. Nonparametric tests were
performed using the same tests on rank transformed data. Type I (a) error for all tests
was set to 0.05.

RESULTS AND DISCUSSION

Short Term Laboratory Exposures

Survival of C. tentans and H. azteca were not affected during 10 d exposures to
PDMS-spiked sediments. Survival of C. tentans larvae exposed in two separate
experiments to concentrations of up to 1502 or 2590 mg PDMS/kg, dw was not different
than that of individuals in control sediments (Figure 2a, 3a). Survival of H. azteca was
not signiﬁcantly less in sediments containing up to 1900 mg PDMS/kg dw than that of

controls (Figure 4).

12

Growth of C. tentans was significantly less in an initial exposure to PDMS-spiked
sediments, though this result was not replicated when the exposure was repeated. The
dry weight of C. tentans exposed to treatments of 161 or 1562 mg PDMS/kg, dw was
signiﬁcantly less than that of those grown on control sediments (Figure 2b). The ash-free
dry weight of these individuals was significantly less than that of the controls only in the
1562 mg PDMS/kg, dw (Figure 2c). This experiment was repeated and the number of
replicates was doubled to 16 per treatment. In the second exposure there were no
statistically significant differences in dry weight or ash—free dry weight between any of
the silicone-spiked sediments and control sediment (Figure 3b, c). The experiment was
repeated a third time for conﬁrmation, and again there were no statistically significant
reductions in growth in PDMS-spiked sediments (data not shown).

Organic carbon (OC) content of sediment did not affect responses of C. tentans or
H. azteca to PDMS. Virtually all particles in aquatic systems carry an organic coating
that can potentially sorb chemical contaminants (Stumm 1992), and the partitioning of
hydrophobic compounds to this coating is often great (Karickhoff and Morris 1985;
Swackhammer and Skoglund 1991). Sediment OC content has been reported to inﬂuence
the bioavailability and toxicity of some compounds (e. g. Adams 1987; Swartz et al.
1990). Therefore, the potential exists that organic carbon content of sediments may
affect the degree of sorption and perhaps the toxicity of PDMS. Kukkonen and Landrum
(1995) found no reduction in growth or survival of L. variegatus in Lake Michigan
sediments of 0.5% DC content treated with 150 mg PDMS/kg dw. In the present study,
no differences in grth or survival of C. tentans or H. azteca were observed in 10 d

exposures to Lake Michigan sediments collected from the same site as those in the

13

Kukkonen and Landrum study (1995) and spiked with PDMS up to a concentration of
approximately 1200 mg PDMS/kg dw (Figure 5).
Whole Life Cycle Assays

While short-term tests may be used to study the potential for acute toxicity, they
may not be able to predict subtle effects from long-term exposures at the community
level (Benoit et a1. 1997; Ingersoll et al. 1998). In order to better evaluate the certainty
that PDMS is not causing adverse effects in aquatic environments, assessments of the
potential for chronic, sublethal impacts rather than acute toxicity were made by use of
whole life cycle studies with C. tentans and H. azteca.

Survival, growth, emergence and reproduction of C. tentans in PDMS-treated
sediments at concentrations up to 2600 mg PDMS/kg, dw were not significantly different
than from that of C. tentans on untreated sediment. Suggested data quality objectives of
70% survival and 0.48 mg/kg AFDW were achieved in the controls, and values of these
parameters at all PDMS treatments also exceeded these levels (Figure 6). Percent
emergence of adult C. tentans was not signiﬁcantly affected by PDMS-treated sediments,
although neither control nor treatment means exceeded the 60% data quality objective
suggested by the USEPA (Figure 7a). During recent round-robin testing, only 43% of the
labs participating met this emergence criterion (USEPA 1999), indicating that the 60%
level may be greater than what should typically be expected. This 60% emergence value
and the other QA parameters for whole life cycles assays were based on testing in a
single laboratory (Sibley et al. 1996; Sibley et al. 1997; Benoit et al. 1997), and may not
reﬂect likely results in all laboratories (USEPA 1999). There were no signiﬁcant effects

of PDMS on the number of eggs produced per female, and mean production for all

14

treatments was greater than the suggested egg production value of 900 eggs for controls
(Figure 7b). Percent hatch of the eggs was not signiﬁcantly different in any of the spiked
sediments from the control, and all treatments exceeded the 90% suggested data quality
level (Figure 7c). PDMS in sediments did not inﬂuence either egg production or percent
hatch, which suggests that reproduction of C. tentans will not be adversely impacted by
sediment concentrations of up to 2600 mg PDMS/kg, dw.

Survival, growth, and reproduction of H. azteca in whole life cycle assays
were not signiﬁcantly reduced in PDMS-spiked sediments relative to control sediments
up to concentrations as great as 994 mg PDMS/kg,dw. Survival of amphipods at 28, 35
and 42 d in PDMS-treated sediments was not signiﬁcantly different from that in control
sediments, although the percent survival in the control treatment decreased from 87%
after 28 d to 77% and 63% after 35 and 42 (1, respectively (Figure 8). While the data
quality objective of 80% survival after 28 days was achieved, these values were
somewhat less than the mean survivals of 90 (28 d), 89 (35 d), and 86% (42 d) in the
control sediment from West Bearskin Lake during the round-robin assay (USEPA 1999).
These contrasts may be due to differences in physicochemical parameters of the
sediments, such as grain size, which make the Williamston Pond sediment less ideal for
amphipod colonization. Growth of H. azteca was not significantly different between
PDMS-spiked and control sediments after 28 or 42 d (Figure 9a, b), although variability
was great in the least PDMS treatment after 28 (1 (CV = 43% in 9 mg PDMS/kg, dw).
This is likely due to the few replicates for this endpoint (N = 4), and could be improved
with increased replication. The control data quality objective of 0.15 mg/individual after

28 d was achieved in all treatments. Reproduction in all treatments exceeded the

15

 

suggested control data quality value of 2 young per female and there was no difference
between any individual PDMS treatment and the control (Figure 9c). The suggested
minimum data quality objective for reproduction in control treatments of 2 young per
female was exceeded in all treatments. However, the reproduction endpoint was more
variable than either growth or survival. Coefficients of variation were 64, 90, 71, and
64% in the control, 9, 104, and 994 mg PDMS/kg, dw treatments, respectively. This
great variability suggests that this may not be an appropriate measure of contaminant
effects, or increased replication may be needed to improve statistical resolution of this
endpoint.

In Situ Bioassays

While acute and sublethal endpoints can be measured in the laboratory with some
degree of accuracy and can provide unique data on the potential toxicity of sediments, the
ecological consequences of exposure to contaminants may more realistically be evaluated
at higher levels of organization (Underwood and Peterson 1988; Clements and Kiffney
1994). Field studies with artiﬁcial substrates were designed to simulate a realistic
exposure scenario; that of sewage sludge outfall into a lake ecosystem.

Measured PDMS concentrations for PDMS-spiked artiﬁcial substrate treatments
were 13.4, 75.7, and 798 mg PDMS/kg dw upon collection. The control blank, which
was treated with neither sludge nor PDMS, and the sludge blank contained 3.0 and 2.3
mg PDMS/kg, dw respectively, which indicates some level of background contamination
in these sediments. This study site lies within a bay that contains a public beach and it is
possible that the background PDMS levels are an artifact of suntan lotion deposited by

bathers.

l6

Almost all of the macroinvertebrates present in the trays after in situ incubation
were chironomids or oligochaetes. The abundance of chironomids in the three PDMS-
spiked treatments was within 30% of control blank values, while the number of
individuals (392 larvae) in the sludge blank was only about 50% of the control blank
level of 773 animals (Figure 10a). The percent of predaceous midges in the treatments
ranged from 27 to 41% (Figure 10b). Oligochaete abundances in the sludge blank and at
13.4 and 75.7 mg PDMS/kg, dw were 58, 76 and 71% the number in the control blank,
respectively (Figure 10a). The total number of oligochaetes in the greatest PDMS
treatment (798 mg PDMS/kg,dw) was only 8% that of the untreated control blank and
14% that of the sludge—only control (Figure 10a).

The most prominent difference observed among treatments was the decrease in
oligochaete abundance in the presence of 798 mg/kg PDMS/kg, dw, the greatest
concentration tested (Figure 10a). Several explanations were considered, including the
presence of a greater number of predators in this treatment, or the possibility that PDMS
may be adversely impacting oligochaetes. The proportion of predaceous midges at 798
mg PDMS/kg, dw was less than that of the control blank and the sludge blank, and
similar to that of the 13 and 76 mg PDMS/kg, dw treatments, thus suggesting that
predators are not the source of the effect.

The observation of apparent PDMS-related effects on oligochaetes under ﬁeld
conditions was the impetus for laboratory exposures with the oligochaete Lumbriculus
variegatus. Short-term laboratory bioassays were conducted which indicated no potential
toxicity from PDMS in sediments based on survival and growth (Figure 11). This lack of

observed effects is similar to the results of other studies with PDMS-spiked sediments

l7

 

and aquatic worms, including L. variegatus (Aubert et al. 1985; Guillemaut—Drai et al.
1988; Craig and Caunter 1990; Kukkonen and Landrum 1995). It is possible that the
variability among treatments in the in situ assays was due to the patchy structure of
natural invertebrate communities, and we hope to understand this better upon sorting
further samples.

In summary, the present study corroborates and expands upon previous research
(Table 1) which has indicated minimal toxicity of PDMS to sediment-dwelling
organisms. PDMS in sediments did not affect growth, survival, or reproduction of C.
tentans or H. azteca in laboratory tests at concentrations that were at least 20 fold greater
than those which occur in the environment. Reproduction of H. azteca was too variable
to observe clear trends within treatments. The suggestion that silicone may decrease
oligochaete abundance in ﬁeld studies was contradicted by the published literature and
could not be substantiated in our own lab studies. Results of this study indicate that
current concentrations of PDMS ﬂuid in the aquatic environment will not induce

signiﬁcant effects on benthic invertebrate assemblages.

18

 

Table 1. Effects of polydimethylsiloxane on benthic invertebrates.

 

 

Test Organism Maximum Endpoint and Result Reference
Concentration
Tested
Polychaete 10,000 mg/kg No reduction in growth or survival Craig and Caunter 1990
Nereis in sediment to 28 (1 though burrowing activity
diversicolor was affected, perhaps due to
pretreatment of sediments rather
than to PDMS
Polychaete 10,000 mg/L Low toxicity and Aubert et al. 1985;
Nereis in water bioaccumulation/biomagniﬁcation Guillenaut et al. 1987
diversicolor; potential in marine trophic chains
Mussels
Mytilus edulis &
Mytilus
galloprovincialis;
Crab (Crustacean)

Carcinus maenas

Oligochaete 150 mg/kg in No toxicity or bioaccumulation; Kukkonen and Landrum
Lumbriculus sediment rapid depuration of PDMS. PDMS 1995
variegatus in sediments reduced the

bioaccumulation of Benzo[a]pyrene

CH3 CH3 CH3

CH3" Si--O -- Si-- 0 -- $1-- CH3

CH3 - C'H3 - H CH3

 

 

Figure 1. Generic structure of Polydimethylsiloxane (PDMS).
n can range from 10 to >10,000 (Stark et al. 1982).

20

Figure 2. Responses in Chironomus tentans bioassays with PDMS-spiked sediments:
Trial 1 of 2. Means, with standard error (N = 16). * indicates statistically signiﬁcant
difference at CL = 0.05 using Dunnett’s test for differences between each treatment and a

control. (a) grth measured as dry weight per individual; (b) growth measured as ash-
free dry weight per individual.

21

ay 1: Survival

Ass

 

(23) Chironomus tentans

 

 

   

\ \ .

     

161
nﬁaﬁon
nhaﬁon

 

Assay 1: Ash-Free Dry Weight

Assay 1: Dry Weight

 

18
PDMS Conce
PDMS Conce

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

(2b) Chironomus tentans
(2c) Chironomus tentans

 

 

 
 

ntration

22

PDMS Conce

Figure 3. Responses in Chironomus tentans bioassays with PDMS-spiked sediments:
Trial 2 of 2. Means, with standard error (N = 16). * indicates statistically signiﬁcant
difference at on = 0.05 using Dunnett’s test for differences between each treatment and a

control. (a) grth measured as dry weight per individual; (b) growth measured as ash-
free dry weight per individual.

23

(3a) Chironomus tentans Assay 2: Survival

 

   

 

 

 

 

 

 

 

  

 

000000
00000

3E5. as

PDMS Concentration

(3b) Chironomus tentans Assay 2: Dry Weight

 

 

 

 

 

 

 

@535
a? be

2 0
PDMS Concentration

 

(3c) Chironomus tentans Assay 2: Ash—Free Dry Weight

 

 

    

\\

     

\

PDMS Concentra

 

   
 

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oooooooo
@535
a? to 003:3

tron

24

 

 

 

 

mmmmmmwmmmo

 

12).

PDMS-spiked sediments. Means, with

Figure 4. Survival in Hyalella azteca bioassay with
standard error (N

25

Figure 5. Responses of Chironomus tentans and Hyalella azteca with PDMS-spiked
sediments of low organic carbon content (0.5% DC). Means, with standard error. (a)
percent survival of Chironomus tentans (N = 16). (b) growth of Chironomus tentans
measured as dry weight per individual (N = 16). (c) percent survival of Hyalella azteca
(N = 12). ((1) grth of Hyalella azteca measured as dry weight per individual (N = 12).

26

(5b)

(58)

 

 

 

 

 

 

Km
3

 

 

3525150

news? 5..

1246

1246

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PDMS Concentration

PDMS Concentration

(5d)

(50)

 

0.20

 

I

awe can?

 

0

mo

 

PDMS Concentration

1155

PDMS Concentration

0

27

 

Figure 6. Responses in Chironomus tentans whole-life cycle bioassays with PDMS-
spiked sediments. Dashed lines indicate suggested USEPA data quality objectives. (a)
percent survival at 20 d (N = 4). (b) Ash-free dry weight at 20 d (N = 4).

28

(6a) C. tentans % survival at 20 din whole-life cycle assay.

 

 

 

 

000000
00000

ntration

PDMS Conce

(6b) C. tentans growth at 20 d in whole-life cycle assay.

 

 

E:

 

 

6.5 43 be
82:2

m

29

Figure 7. Responses in Chironomus tentans whole-life cycle bioassays with PDMS-
spiked sediments. Dashed lines indicate suggested USEPA data quality objectives. (a)

C. tentans percent emergence (N = 8). (b) reproduction in number of eggs/female. (c)
percent of eggs hatching.

30

(7a) C. tentans percent emergence in whole-life cycle assay.

 

 

ntration

\R\2

\\\\
\\
x
\\

7777777

   

PDMS Conce

 

 

\\\\\\\\\\\\\\\\

 

tentans reproduction in whole-life cycle assay.

 

 

 

 

 

(7c) C. tentans percent egg hatch in whole-life cycle assay.

 

vac—om com
mwmm mo Leena—Z

tration

31

PDMS Concen

Figure 8. Responses in Hyalella azteca whole-life cycle bioassays with PDMS-spiked
sediments. Dashed lines (when present) indicate suggested USEPA data quality
objectives. (a) percent survival at 28 d when animals are isolated from sediments and
placed into water-only chambers (N = 12). (b) Percent survival at 35 d (N: 8). (c)
Percent survival at 42 (1 (test termination, N = 8).

32

(8a) H. azteca percent survival at 28 d in whole-life cycle assay.

 

 

 

 

 

 

(8b) H. azteca percent survival at 35 din whole-life cycle assay.

 

            
          

       

 

 

 

o o 00 0 o
o a 6‘ 2
I

9
PDMS Conce

ntration

(8c) H. azteca percent survival at 42 d in whole-life cycle assay.

 

 

 

 

 

 

 

33

Figure 9. Responses in Hyalella azteca whole-life cycle bioassays with PDMS-spiked
sediments. Dashed lines (when present) indicate suggested USEPA data quality
objectives. (a) growth at 28 d when animals are isolated from sediments and placed into
water-only chambers (N = 4). (b) growth at 42 (1 (test termination, N = 8). (c)
reproduction at 42 (1 evaluated in number of young produced per female.

34

(9a) H. azteca growth at 28 din whole-life cycle assay.

   

   
    

asssssw

amass m
_____

mm\\\\\

         

  

 

 

 

 

0000.

REV Emma? DD

m

ay.

(9b) H. azteca growth at 42 d in whole-life cycle ass

 

  

        
            

 

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000000000
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000000000

 

 

 

 

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\0

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Figure 10. Results of in situ study with silicone-spiked sediments. (a) histogram of
oligochaetes and chironomids in artificial substrates. (b) percent of chironomid
predators in artiﬁcial substrates.

36

(10a) Histogram of oligochaetes and chironomids in artiﬁcial substrates.

Freqency
(# Individuals)

Percent Chironomid
Predators

1400
1200
1000
800
600
400
200
0

 

 

 

 

 

 

 

Control Sludge 13.4 75.7 798
Blank Blank
(3.0) (2.3)

PDMS Concentration

(10b) Percent of chironomid predators in artificial substrates.

 

 

 

 

 

 

50

40

30 ~

20 ~

10 -

0 _ .
Control Sludge 13.4 75.7 798
Blank Blank
(3.0) (2.3)

PDMS Concentration

37

Figure 11. Responses in Lumbriculus variegatus bioassays with PDMS-spiked
sediments. Means, with standard error (N = 4). (a) percent survival. (b) growth
measured as mean wet weight per individual.

38

 

 

 

 
 

 

 

 

(11a) Lumbriculus variegatus survival in PDMS-spiked sediments.

 

000000000

888888888

m

5 w
Q.»

m

m m
\ o z.
m .c
m
m

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m

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5233

1275

PDMS Concentration
39

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42

APPENDICES

43

APPENDIX 1: STANDARD PROCEDURE FOR POLYDIMETHYLSILOXANE
(PDMS) EXTRACTION FROM SEDIMENT
Aquatic Toxicology Laboratory Standard Operating Procedure
Department of Zoology
Michigan State University

Title:
Polydimethylsiloxane (PDMS) Solvent Extraction from Sediment for Analysis by
GPC-ICP

Description of Method:
This protocol has been adapted from SOP NE209_02.GLP provided by Northeast
Analytical (NEA) to Dow Corning Corporation. Contact at NBA is Inga C.
Hotaling (518) 346-4592. To avoid concentration and degradation of the sample,
materials should be sealed in plastic and stored at 4°C. This method involves a
multiple tetrahydrofuran (THF) extraction of test sediments to isolate the PDMS
present. Extracts are evaporated to near dryness and shipped offsite for analysis
by ICP.

Hazards and Precautions:
Care must be exercised when using solvents. THF is moderately toxic and very
volatile and ﬂammable. Extractions should be done in a well-ventilated hood.

Materials:
HPLC grade Tetrahydrofuran (THF) Available from MSU Stores.
Teﬂon (preferable) or Nalgene wash bottle for THF storage and delivery

Clean beakers

Volumetric pipette (10 mL)

Wrist-action or platform shaker.

Fume hood

Laboratory centrifuge

35 mL Kimax heavy duty centrifuge tubes (VWR#: 21023-252) For triplicate

extractions, 6 tubes per sample will be required; 4 tubes per sample for
duplicate extractions

Teﬂon-lined caps for Kimax tubes

Aluminum weigh pans for sediment dryzwet weight ratios

Drying oven

Pasteur pipets (short stem) for solvent transfer
Procedure:
Dry: Wet Weight Ratio

1) Place 5 i 0.3 g of wet, homogenized sediment onto a pre-dryed and pre-

weighed aluminum weigh pan. Record actual weight of sediment.

2) Dry sample for at least 24 h at 60°C.

3) Weigh pan + sediment and record weight.

4) Calculate dry weight as the difference between the pan + dry sediment weight

and the pan weight.

5) Calculate dryzwet weight ratio as dry sediment weight / wet sediment weight.
Sediment Sample Preparation

1) Rinse all glassware and pipets with THF before use. Dispose of THF in

hazardous waste container.

45

2)

3)

4)

5)

6)

7)

Remove sediment sample from refrigerator and allow sample to equilibrate to
ambient temperature. Homogenize sample within the sample container using
a stainless steel spoon.

Weigh approximately 10.0 g of sediment into a centrifuge tube and record
sample weight to 0.01 g. Samples should be done at least in duplicate. If
sample contains excessive water, tubes may be sealed and centrifuged for 3
minutes at high speed to isolate solids. Water may then be pipetted off and
discarded.

Add 10 mL of THF to each centrifuge tube, seal with a Teﬂon-lined cap and
shake for 10 min on laboratory shaker.

Centrifuge sample on high speed for 3 min to isolate solids. Transfer THF
into another clean and THF-rinsed centrifuge tube. Blow down extract under
a low ﬂow of nitrogen.

Repeat steps B4-B5 two more times, combining the extracts and blowing
down to dryness after the third extraction step.

Add 0.5 mL of water to prevent PDMS degradation and ship sealed and

labeled tubes to Dow Coming for analysis as soon as possible.

Extraction Blank Preparation

Repeat steps 4-7 with an empty centrifuge tube (no soil).

Reporting concentrations of PDMS results in soil, sediment and sludge.

1)

Report PDMS present in extraction blank in u g.

46

2) Report PDMS concentration in environmental samples in pg PDMS/g dry
weight of sediment. Wet weight concentration may also be reported, but

wetzdry ratios should then be given.

47

. APPENDIX 2: STANDARD PROCEDURE FOR A SHORT-TERM GROWTH
AND SURVIVAL ASSAY WITH THE MIDGE CHIRONOMUS TENTANS

Aquatic Toxicology Laboratory Standard Operating Procedure
Department of Zoology
Michigan State University
Title: Short-term growth and survival assay with the midge C. tentans.

K.S. Henry
Description of Method:

The recommended 10-d sediment toxicity test with C. tentans must be conducted
at 23 degress C with a 16L:8D photoperiod at an illuminace of about 500 to 1000 lux.
Test chambers are 300 mL high-form lipless beakers containing 100 mL of sediment and
175 mL of overlying water. Ten third instar midges are used to start a test. The numbers
of replicates/treatment depends on the objective of the test. Eight replicates are
recommended for routing testing. Midges in each test chamber are fed 1.0 mL of a 4 g/L
suspension of Tetramin daily. Each chamber receives 2 volume additions/d of overlying
water. Water renewals may be manual or automated, though we usually use manual
methods in our lab. Overlying water can be culture water, well water, surface water, site
water, or reconstituted water. We normally use 30% well water/70% reverse osmosis
puriﬁed or 70% ultrapure for our cultures and tests. For site-speciﬁc evaluations, the
characteristics of the overlying water should be as similar as possible to the site where
sediment is collected.

Referenced Documents:

48

Ankley, G.T., D.A. Benoit, R.A. Hoke, E.N. Leonard, C.W. West, G.L. Phipps, V.R.
Mattson, and LA. Anderson. 1993. Development and evaluation of test methods
for benthic invertebrates and sediments: Effects of ﬂow rate and feeding on water
quality and exposure conditions. Arch. Environ. Contam Toxicol. 25: 12-19.

Ingersoll, CG. and MK. Nelson. 1990. Testing sediment toxicity with Hyalella azteca
(Amphipoda) and Chironomus riparius (Diptera). In: W.G. Landis and W.H. van
der Schalie (eds), Aquatic toxicology and risk assessment, 13th volume. ASTM
STP 1096. Philadelphia, PA.

Kemble, N.E., W.G. Brumbaugh, E.L. Brunson, F.J. Dwyer, C.G. Ingersoll, D.P. Monda,
and DP. Woodward. 1994. Toxicity of metal-contaminated sediments from the
Upper Clark Fork River, MT, to aquatic invertebrates in laboratory exposures.
Environ. Toxicol. Chem. 13(12): 1985-1997.

Reynoldson, T.B., K.E. Day, C. Clarke, D. Milani. 1994. Effect of indigenous animals on
chronic endpoints in freshwater sediment toxicity tests. Environ. Contam. Toxicol.
13:973-977.

Sadler, W.O. 1935. Biology of the midge Chironomus tentans Fabricus, and methods for
its propagation. Mem. Cornell Univ. Agric. Exp. Stat. 173. 25 pp.

USEPA. 1994. Methods for measuring the toxicity and bioaccumulation of sediment—
associated contaminants with freshwater invertebrates. EPA-600/R-94/024.
Environmental Research Laboratory, Duluth, Minnesota. 133 p.

Procedure:

Preparation of Test Chambers

49

The day before the sediment test is started (Day —1) each sediment should be
thoroughly mixed and added to the test chambers. Sediment should be visually inspected
to judge the degree of homogeneity. Excess water on the surface of the sediment can
indicate separation of solid and liquid components, and sediment should be thoroughly
mixed. If a quantitative measure of homogeneity is required, replicate sub-samples should
be taken from the sediment batch and analyzed for TOC, chemical concentrations, and
particle size. This can be done in the soil analysis lab in the Crop and Soil Science
Department on MSU‘s campus.

Preparation of Test Chambers

Overlying water is added to the chamber in a manner that minimizes suspension of
sediment. This is best accomplished by slowly pouring the water onto a Styrofoam disk
placed on top of the sediment to dissipate the force of the water. Renewal of overlying
water is started on day -1 by performing a water exchange after all replicates for the test
have been set up. A test begins when the organisms are added to the test chambers (day
0).

Renewal of Overlying Water

Two volume additions of overlying water should be delivered to each test chamber
daily over the course of the test. Routine water quality parameters will be measured at
regular intervals during the test (see below). The number of replacements per day should
be increased if hardness, alkalinity or ammonia concentrations in the overlying water vary
by more than 50% among treatments, or if oxygen drops to less than 30% saturation.
Although contaminant concentrations are reduced in the overlying water in water-renewal

tests, organisms in direct contact with sediments will generally still receive a substantial

50

proportion of a contaminant dose directly from the whole sediment and/or from the
interstitial water. The water delivery system should be calibrated and run for a few days
before a test is started to verify that the system is functioning properly.

Acclimation of Test Organisms and Placement into Test Chambers

Test organisms should be cultured and tested at 23°C in the same water type.
However, if a sufﬁcient supply of culture water is not available for the exposure,
acclimation of test organisms to the test water is not required. Performance standards for
this test are calibrated to the response of midges at 23°C. If the test must be run at a
different temperature, then the organisms should be acclimated at a rate of 1°C every 2 h
to prevent thermal shock.

Test organisms should be disturbed as little as possible. Midges should be
introduced into the overlying water below the air-water interface. Test organisms can be
pipetted directly into overlying water (Ankley et al. 1993). In our lab we typically place 5
larvae into culture water in 30 mL plastic cups, randomize these cups, and ﬂoat them in
the exposure chambers for 15 min before dumping the contents into the chambers.
Feeding

Each beaker receives a daily addition of 1.0 mL of 4g/L Tetramin. Suspensions of
food should be thoroughly mixed before aliquots are taken. If excess food collects on the
sediment, a fungal or bacterial growth may develop on the sediment surface, in which case
feeding should be suspended for one or more days. A drop in dissolved oxygen below
40% of saturation during a test may indicate that the food added is not being consumed.
Feeding should be suspended for the amount of time necessary to increase the dissolved

oxygen concentration. If feeding is suspended in one treatment, it should be suspended in

51

all treatments. Detailed records of feeding rates and appearance of the sediment should be
maintained daily.
Overlying Water Quality

All chambers should be checked daily and observations made to assess test
organism behavior such as sediment avoidance. However, monitoring effects on
burrowing activity of test organisms may be difﬁcult because the test organisms are often
not visible during the exposure. The operation of the exposure system should be
monitored daily.

Conductivity, hardness, pH, alkalinity, and ammonia should be measured in all
treatments at the beginning and end of a test. Overlying water should be sampled just
before water renewal from about 1 to 2 cm above the sediment surface using a pipet. It
may be necessary to pool water samples from individual replicates. The pipet should be
checked to make sure no organisms are removed during sampling of overlying water.
Hardness, alkalinity, pH, conductivity, and ammonia in the overlying water within a
treatment should not vary by more than 50% during a test.

Dissolved oxygen should be measured daily and should be between 40 and 100%
saturation. If a probe is used to measure dissolved oxygen in overlying water, it should be
thoroughly inspected between samples to make sure that organisms are not attached and
should be rinsed between samples to minimize cross contamination. Aeration can be used
to maintain dissolved oxygen in the overlying water above 40% saturation, though
increasing the number of water exchanges per day is preferable. Dissolved oxygen and pH

can be measured directly in the overlying water with a probe.

52

Temperature should be measured at least daily in at least one test chamber from
each treatment. The temperature of the water bath should also be continuously monitored
using a max/min thermometer. Usually running the water bath in the ﬁberglass tanks at
25°C will result in an appropriate exposure chamber temperature (i.e. 23°C). The daily
mean test temperature must be 231°C. The instantaneous temperature must always be
23 i 3°C.

Ending a Test

Immobile organisms isolated from the sediment surface or from sieved material
should be considered dead. A #25 (710nm mesh) sieve can be used to isolate midges
from sediment. Surviving test organisms can be counted and placed into weigh pans or
preserved in 8% sugar formalin solution for growth measurements. The sugar-forrnalin
solution is made by adding 120 g of sugar to 80 mL of formalin, and bringing the solution
up to 1 L with reverse osmosis water. This solution is mixed 1:1 with sample water to
preserve organisms.

Test Data

Ash-free dry weight (AFDW) and survival are the endpoints measured at the end
of the 10 (1 test. The duration of the test starting with third instar is not long enough to
determine emergence of adults. Traditionally, dry weight (DW) has been used for the
larval growth endpoint on midge tests. However, it has been shown that the grain size of
sediments inﬂuences the amount of sediment that C. tentans larvae ingest and retain in
their gut. As a result, in ﬁner-grain sediments, a substantial portion of the measured dry
weight may be comprised of sediment rather than tissue. While this may not represent a

strong bias in tests with identical grain size distributions in all treatments, most ﬁeld

53

assessments are likely to have varying grain size among sights. This will likely create
differences in dry weight among treatments that are not reﬂective of true somatic growth.
For this reason, it has been suggested that weight of midges should be measured as ash-
free dry weight (AFDW) instead of dry weight. AFDW will more directly reﬂect actual
differences in tissue weight by reducing the inﬂuence of sediment in the gut.

To ﬁnd AFDW, all living larvae per replicate are combined in a pre-ashed
aluminum weigh boat and dried at 60°C for 24h. The samples should be cooled in a
dessicator and weighed to the nearest 0.01 mg to obtain mean weights per surviving
organism per replicate. When weighing, remove only 4 weigh boats from the dessicator at
a time and weigh them. When done, remove 4 more and so on until all are weighed. The
dry pans will absorb water out of the air and increase in weight, so they should be weighed
as soon as possible after removal from the dessicator. The dried larvae in the pan are then
ashed at 550°C for 2h. The pan with the ashed larvae is then re-weighed and the tissue
mass of the larvae is determined as the difference between the weight of the dried larvae
plus pan and the weight of the ashed larvae plus pan. In rare instances, where
preservation is required, an 8% sugar-formalin solution can be used to preserve samples.
The sugar-forrnalin solution is made by adding 120 g of sugar to 80 mL of formalin, and
bringing the solution up to 1 L with DI water. This solution is mixed 1:1 with sample
water to preserve organisms. Sugar formalin is preferable to alcohol for preservation
because it does not cause as much shrinkage as EtOH. Average AFDW of C. tentans in
control sediments must be at least 0.48 mg at the end of the test. Head capsule width can
also be measured on surviving midges at the end of a test before AFDW is determined,

though we do not do this routinely.

54

Inﬂuence of Indigenous Organisms

The inﬂuence of indigenous organisms on the response of C. tentans has not been
well studied, though survival of C. riparius was not reduced by the presence of
oligochaetes in sediments (Reynoldson et al. 1994). Therefore, it is important to
determine the number and biomass of indigenous organisms in ﬁeld-collected sediment in
order to better interpret growth data (Reynoldson et a1. 1994). Furthermore, presence of

predators may also inﬂuence the response of test organisms in sediment (Ingersoll and

Nelson 1990).

55

Table 1. Test conditions for conducting a 10 d sediment toxicity test with Chironomus

 

 

tentans.

Parameter Conditions

1. Test Type Whole-sedirnent toxicity test with renewal of
overlying water

2. Temperature 23i1°C

3. Illurninance About 500 to 1000 lux

4. Photoperiod 16L:8D

5. Test chamber 300 mL high-form lipless beakers

6. Sediment volume 100 mL

7. Overlying water volume 175 mL

8. Age of organisms Third instar larvae

9. Number of organisms/chamber 10

10. Number of replicate 8 for routine testing

chambers/treatment

11. Feeding Tetramin goldﬁsh food, fed 1.0 mL daily to each
test chamber

12. Aeration None unless D0 in overlying water drops below
2.5 mg/L

l3. Overlying water Preferably culture water. Well water, surface
water, or site water may also be used.

14. Test chamber cleaning Occasionally the outside of the screen may be
brushed gently with a toothbrush.

15. Overlying water quality Hardness, alkalinity, conductivity, pH and
ammonia at the beginning and end of a test (day
0 and day 10). Temperature and DO daily.

16. Test duration 10 d

17. Endpoints Survival and growth

18. Test acceptability Minimum mean control survival of 70% and

mean weight per surviving control organism of
0.6 mg on day 10 and suggested perforrnance-
based criteria (see below).

 

56

Table 2. General activity schedule for conducting a 10 d sediment toxicity test with
Chironomus tentans.

 

Day

Activity

 

-14
-13

-12

~11

-9to-1

-1

1to9

10

Isolate adults for production of egg masses

Place newly deposited egg masses into hatching dishes (crystallizing dishes
with water)

A larval rearing tank is prepared by ﬁlling a 5 gal ﬁsh tank 1/3 full and
adding 2 cups of ground paper towel

Examine egg masses for hatching success. If egg masses have hatched,
transfer larvae and eggs into the larval rearing tank. Feed midges with
20g/L Tetramin suspension. It is best to start several tanks if enough egg
masses are available.

Feed and observe midges.

Add sediment into each test chamber, place chambers into exposure
system, feed each chamber, and start renewing overlying water.

Measure total water quality (pH, temperature, DO, hardness, alkalinity,
conductivity, ammonia. Isolate 3rd instar larvae from the culture tanks.
Add 1.0 mL of Tetramin (4.0 g/L) into each test chamber. Transfer 10
larvae into each test chamber. Release organisms under the surface of the
water.

Add 1.0 mL of Tetramin (4.0 g/L) to each test beaker. Measure
temperature and DO daily. Observe and record behavior of test organisms.
Measure temperature, DO, pH, hardness, alkalinity, conductivity and
ammonia. End the test by collecting the amphipods with a #25 mesh sieve
(710 um mesh). Measure weight of surviving larvae.

57

Table 3. Test acceptability requirements for conducting a 10 d sediment toxicity test with
Chironomus tentans.

 

The following performance criteria for the short-term test with C. tentans have been
recommended by the EPA:

 

1. Tests must be started with 3rd instar and younger larvae.

2. Average survival of C. tentans in the control sediment must be 2 70% at the end of
the test.

3. Average size of C. tentans in the control sediment must be at least 0.6 mg at the end
of the test. _

4. Hardness, alkalinity, pH, and ammonia in the overlying water typically should not
vary by more than 50% during the sediment exposure.

 

B. Additional Requirements

 

1. All organisms in a test must be from the same source.
2. It is desirable to start tests as soon as possible after collection of sediment from the
ﬁeld.

3. Negative-control sediment must be included in a test.

4. Test organisms should be cultured and tested at 23°C.

5 The mean of the daily test temperature must be 23il°C. The instantaneous
temperature must be 23;3°C.

 

58

APPENDIX 3: STANDARD PROCEDURE FOR A SHORT-TERM GROWTH
AND SURVIVAL ASSAY WITH THE AMPHIPOD H YALELLA AZTECA

Aquatic Toxicology Laboratory Standard Operating Procedure
Department of Zoology
Michigan State University

Title: Short-term growth and survival assay with the amphipod Hyallela azteca.

K.S. Henry
Description of Method:

The recommended 10-d sediment toxicity test with H. azteca must be conducted at
23 degress C with a 16L:8D photoperiod at an illuminace of about 500 to 1000 lux. Test
chambers are 300 mL high-form lipless beakers containing 100 mL of sediment and 175
mL of overlying water. Ten 7-to 14-d old amphipods are used to start a test. The
numbers of replicates/treatment depends on the objective of the test. Eight replicates are
recommended for routing testing. Amphipods in each test chamber are fed 1.5 mL of
YCT food daily. Each chamber receives 2 volume additions/d of overlying water. Water
renewals may be manual or automated, though we usually use manual methods in our lab.
Overlying water can be culture water, well water, surface water, site water, or
reconstituted water. We normally use 30% well water for our cultures and tests. For site-
speciﬁc evaluations, the characteristics of the overlying water should be as similar as
possible to the site where sediment is collected.

Referenced Documents:

59

Environment Canada. 1997. Biological test method: Test for survival and growth in
sediment using the freshwater amphipod Hyalella azteca. EPS 1/RM/33. Method
Development and Application Section. 123 p.

Ingersoll, CG. and MK. Nelson. 1990. Testing sediment toxicity with Hyalella azteca
(Amphipoda) and Chironomus riparius (Diptera). In: W.G. Landis and W.H. van
der Schalie (eds.), Aquatic toxicology and risk assessment, 13th volume. ASTM
STP 1096. Philadelphia, PA. pp 93-109.

Kemble, N.E., W.G. Brumbaugh, E.L. Brunson, F.J. Dwyer, C.G. Ingersoll, D.P. Monda,
and DE Woodward. 1994. Toxicity of metal-contaminated sediments from the
Upper Clark Fork River, MT, to aquatic invertebrates in laboratory exposures.
Environ. Toxicol. Chem. 13: 1985-1997.

Reynoldson, T.B., K.E. Day, C. Clarke, D. Milani. 1994. Effect of indigenous animals on
chronic endpoints in freshwater sediment toxicity tests. Environ. Contam. Toxicol.
13:973-977.

USEPA. 1994. Method for measuring the toxicity and bioaccumulation of sedirnent-
associated contaminants with freshwater invertebrates. EPA-600/R-94/024.
Environmental Research Laboratory, Duluth, Minnesota. 133 p.

Procedure:

Preparation of Test Chambers
The day before the sediment test is started (Day -1) each sediment should be

thoroughly mixed and added to the test chambers. Sediment should be visually inspected

to judge the degree of homogeneity. Excess water on the surface of the sediment can

indicate separation of solid and liquid components. If a quantitative measure of

60

homogeneity is required, replicate sub-samples should be taken from the sediment batch
and analyzed for T OC, chemical concentrations, and particle size. This can be done in the
soil analysis lab in the Crop and Soil Science Department on MSU's campus.
Preparation of Test Chambers

Overlying water is added to the chamber in a manner that minimizes suspension of
sediment. This is best accomplished by slowly pouring the water onto a Styrofoam disk
placed on top of the sediment to dissipate the force of the water. Renewal of overlying
water is started on day -1 by performing a water exchange after all replicates for the test
have been set up. A test begins when the organisms are added to the test chambers (day
0).
Renewal of Overlying Water

Two volume additions of overlying water should be delivered to each test chamber
daily over the course of the test. Routine water quality parameters will be measured at
regular intervals during the test (see below). The number of replacements per day should
be increased if hardness, alkalinity or ammonia concentrations in the overlying water vary
by more than 50% among treatments, or if oxygen drops to less than 30% saturation.
Although contaminant concentrations are reduced in the overlying water in water-renewal
tests, organisms in direct contact with sediments will generally still receive a substantial
proportion of a contaminant dose directly from either the whole sediment or from the
interstitial water. The water delivery system should be calibrated and run before a test is
started to verify that the system is functioning properly.

Acclimation of Test Organisms and Placement into Test Chambers

61

Test organisms should be cultured and tested at 23°C in the same test water.
However, if a sufﬁcient supply of culture water is not available for the exposure,
acclimation of test organisms to the test water is not required. Performance standards for
this test are calibrated to the response of amphipods at 23°C. If the test must be run at a
different temperature, then the organisms should be acclimated at a rate of 1°C every 2 h
to prevent thermal shock.

Test organisms should be disturbed as little as possible. To start a test, 7 to 14 d
old amphipods are collected with a Pasteur pipette from the bottom of a crystallizing dish
with the aid of a dissecting microscope. Test organisms are pipetted very gently directly
into the overlying water of the exposure chambers. Care should be taken to release them
under the surface of the water. Alternatively, we often place 5 amphipods into culture
water in 30 mL plastic cups, randomize these cups, and ﬂoat them in the exposure
chambers for 15 min before dumping the contents into the chambers.

Feeding

Each beaker receives a daily addition of 1.5 mL of yeast, cerophyll, trout chow
suspension (i.e. YCT). Suspensions of food should be thoroughly mixed before aliquots
are taken. If excess food collects on the sediment, a fungal or bacterial growth may
develop on the sediment surface, in which case feeding should be suspended for one or
more days. A drop in dissolved oxygen below 40% of saturation during a test may
indicate that the food added is not being consumed. Feeding should be suspended for the
amount of time necessary to increase the dissolved oxygen concentration. If feeding is
suspended in one treatment, it should be suspended in all treatments. Detailed records of

feeding rates and appearance of the sediment should be maintained daily.

62

Overlying Water Quality

All chambers should be checked daily and observations made to assess test
organism behavior such as sediment avoidance. However, monitoring effects on
burrowing activity of test organisms may be difﬁcult because the test organisms are
usually not visible during the exposure. The operation of the exposure system should be
monitored daily.

Conductivity, hardness, pH, alkalinity, and ammonia should be measured in all
treatments at the beginning and end of a test. Overlying water should be sampled just
before water renewal from about 1 to 2 cm above the sediment surface using a pipet. It
may be necessary to pool water samples from individual replicates. The pipet should be
checked to make sure no organisms are removed during sampling of overlying water.
Hardness, alkalinity, pH, conductivity, and ammonia in the overlying water with a
treatment should not vary by more than 50% during a test.

Dissolved oxygen should be measured daily and should be between 40 and 100%
saturation. If a probe is used to measure dissolved oxygen in overlying water, it should be
thoroughly inspected between samples to make sure that organisms are not attached and
should be rinsed between samples to minimize cross contamination. Aeration can be used
to maintain dissolved oxygen in the overlying water above 40% saturation, though
increasing the number of water exchanges per day is preferable. Dissolved oxygen and pH
can be measured directly in the overlying water with a probe.

Temperature should be measured at least daily in at least one test chamber from
each treatment. The temperature of the water bath should also be continuously monitored

using a max/min thermometer. Usually running the water bath in the ﬁberglass tanks at

63

25°C will result in an appropriate exposure chamber temperature (i.e. 23°C). The daily
mean test temperature must be 23 i 1°C. The instantaneous temperature must always be
23 i 3°C.
Ending a Test

Any of the surviving amphipods in the water column or on the surface of the
sediment can be pipetted from the beaker before sieving the sediment. Immobile
organisms isolated from the sediment surface or from sieved material should be considered
dead. A #25 sieve (710 um mesh) can be used to isolate amphipods from sediment.
Alternatively, Kemble et al. (1994) recommended sieving sediment using the following
procedure: (1) pour about half of the overlying water through a #50 (300 um) U.S.
Standard mesh sieve, (2) swirl the remaining water to suspend the upper 1 cm of sediment,
(3) pour this slurry through the #50 mesh sieve and wash the contents of the sieve into an
examination pan. Surviving test organisms should be counted and preserved in 8% sugar
formalin solution for growth measurements. The sugar-formalin solution is made by
adding 120 g of sugar to 80 mL of formalin, and bringing the solution up to 1 L with R0
water. This solution is mixed 1:1 with sample water to preserve organisms.
Test Data

Survival is the primary endpoint recorded at the end of the 10-d sediment toxicity
test with H. azteca. Measuring growth is optional; however, growth of amphipods may be
a more sensitive toxicity endpoint than survival (e. g. Kemble et al. 1994). The duration of
the 10-d test started with 7- to 14- old amphipods is not long enough to determine sexual
maturation or reproductive effects, though the multi life stage test does consider these

endpoints.

Amphipod body lengh (i 0.1 m) can be measured from the base of the ﬁrst
antenna to the tip of the third uropod along the curve of the dorsal surface. Antenna]
segment number can also be used to estimate length or weight of amphipods. Wet or dry
weight measurements have also been used to estimate growth of H. azteca, and in our lab
we normally use dry weight. Dry weight of amphipods should be determined by pooling
all living organisms from a replicate into a homemade aluminum "weigh boat" about 1 cm
in diameter. Samples are covered and dried at 60°C for 24 h. The samples should be
cooled in a dessicator and weighed to the nearest 0.01 mg to obtain mean weights per
surviving organism per replicate. When weighing, remove only 4 weigh boats from the
dessicator at a time and weigh them. When done, remove 4 more and so on until all are
weighed. The dry pans will absorb water out of the air and increase in weight, so they
should be weighed as soon as possible after removal from the dessicator.

Inﬂuence of Indigenous Organisms

Survival of H. azteca in 28-d tests was not reduced in the presence of oligochates
in sediment samples (Reynoldson et al. 1994). However, growth of amphipods was
reduced when high numbers of oligochaetes were placed in a sample. Therefore, it is
important to determine the number and biomass of indigenous organirns in ﬁeld-collected
sediments in order to better interpret growth data (Reynoldson et al. 1994). Furthermore,
presence of predators may also inﬂuence the response of test organisms in sediment

(Ingersoll and Nelson 1990).

65

Table 1. Test conditions for conducting a 10 d sediment toxicity test with Hyalella

 

 

azteca.

Parameter Conditions

1. Test Type Whole-sedirnent toxicity test with renewal of
overlying water

2. Temperature 231°C

3. Illuminance About 500 to 1000 lux

4. Photoperiod 16L:8D

5. Test chamber 300 mL high-form lipless beakers

6. Sediment volume 100 mL

7. Overlying water volume 175 mL

8. Age of organisms 7 to 14 (1 old at the start of the test

9. Number of organisms/chamber 10

10. Number of replicate 8 for routine testing

chambers/treatment

11. Feeding YCT food, fed 1.5 mL (1800 mg/L stock) daily
to each test chamber

12. Aeration None unless D0 in overlying water drops below
2.5 mg/L

13. Overlying water Preferably culture water. Well water, surface
water, or site water may also be used.

14. Test chamber cleaning Occasionally the outside of the screen may be
brushed gently with a toothbrush.

15 . Overlying water quality Hardness, alkalinity, conductivity, pH and
ammonia at the beginning and end of a test (day
0 and day 10). Temperature and DO daily.

16. Test duration 10 d

17. Endpoints Survival. Growth optional.

18. Test acceptability Minimum mean control survival of 80% on day

10 and suggested performance-based criteria.

66

 

Table 2. General activity schedule for conducting a 10 d sediment toxicity test with
Hyalella azteca.

 

 

Day Activity

-7 Separate known-age amphipods from the cultures and place in breeding
chambers.

-6 to -1 Feed and observe isolated amphipods.

-1 Add sediment into each test chamber, place chambers into exposure
system, and start renewing overlying water.

0 Measure total water quality (pH, temperature, DO, hardness, alkalinity,

conductivity, ammonia). Transfer ten 7 to 14 (1 old amphipods into each
test chamber. Release organisms under the surface of the water. Add
1.5 mL of YCT into each test chamber.

1 to 9 Add 1.5 mL of YCT to each test beaker. Measure temperature and DO
daily. Observe and record behavior of test organisms.
10 Measure temperature, DO, pH, hardness, alkalinity, conductivity and

ammonia. End the test by collecting the amphipods with a #40 mesh
sieve (425 um mesh).

 

67

Table 3. Test acceptability requirements for conducting a 10 d sediment toxicity test with
Hyalella azteca.

 

A. The following performance criteria for the short-term test with H. azteca have been
recommended by the EPA:

 

1. Age of H. azteca at the start of the test should be 7 to 14 (1 old.
Average survival of H. azteca in the control sediment on day 28 should be
greater than or equal to 80%.

3. Hardness, alkalinity, and ammonia in the overlying water typically should
not vary by more than 50% during the sediment exposure.

B. Additional Requirements

 

1. All organisms in a test must be from the same source.

2. It is desirable to start tests as soon as possible after collection of sediment
from the ﬁeld.

3. Negative-control sediment must be included in a test.

4. Test organisms should be cultured and tested at 23°C.

5 The mean of the daily test temperature must be 23i1°C. The instantaneous

temperature must be 23_-l_-3°C.

 

68

APPENDIX 4: STANDARD PROCEDURE FOR A WHOLE LIFE CYCLE
ASSAY WITH THE MIDGE CHIRONOM US TENTANS

Aquatic Toxicology Laboratory Standard Operating Procedure
Department of Zoology
Michigan State University
Title: Whole life cycle assay with the midge Chironomus tentans.
K.S. Henry
Description of Method:

First instar larvae of the midge Chironomus tentans are placed into exposure
chambers shortly after leaving their egg masses. The test continues through emergence,
reproduction, and hatching of the F1 generation. Survival is determined at 20 d and at the
end of the test. Growth is determined at 20 d, which corresponds to the growth endpoint
in the short-term test started with 10 (1 old larvae. From day 23 to the end of the test,
emergence and reproduction are monitored daily. The number of eggs per female is
determined for each egg case, which is incubated to determine hatching success. Each
treatment is ended when no emergence occurs for 7 d in that treatment. The total length
of the test is normally 50 to 60 d.

Referenced Documents:
Benoit, D.A., Phipps, GA., and Ankley, GT. 1993. A sediment testing intermittent
renewal system for the automated renewal of overlying water in toxicity tests with

contaminated sediments. Water Res. 27:1403-1412.

69

Benoit, D.A., P.K. Sibley, J .L. Juenemann, and GT. Ankley. 1997. Chironomus tentans
life cycle test: design and evaluation for use in assessing toxicity of contaminanted
sediments. Environ. Toxicol. Chem. 16:1165-1 176.

Nebeker, A.V., M.A. Cairns, and CM. Wise. Relative sensitivity of Chironomus tentans
life stages copper. Environ. Toxicol. Chem. 3: 151-158

Reynoldson, T.B., K.E. Day, C. Clarke, and D. Milani. 1994. Effect of indigenous animals
on chronic endpoints in freshwater sediment toxicity tests. Environ. Contam.
Toxicol. 13:973-977.

Sadler, WC. 1935. Biology of the midge Chironomus tentans Fabricus, and methods for
its propagation. Mem Cornell Univ. Agric. Exp. Stat. 173. 25 pp.

Sibley, P.K., D.A. Benoit, and GT. Ankley. 1997. The signiﬁcance of growth in
Chironomus tentans sediment toxicity tests: Relationship to reproduction and
demographic endpoints. Environ. Toxicol. Chem 16:336-345.

USEPA. 1999. Methods for measuring the toxicity and bioaccumulation of sediment—
associated contaminants with freshwater invertebrates: Second edition.
Environmental Research Laboratory, Duluth, Minnesota. Draft document.

Procedure:

Test Conditions
Conditions for conducting a whole life stage sediment toxicity test with C. tentans

are summarized in Table 1. An activity schedule is give as Table 2. This test is conducted

at 23°C with a 16L:8D photoperiod. Test chambers are 300 mL high-form lipless beakers
containing 100 mL of sediment and 175 mL of overlying water (Benoit beakers). Each

test chamber receives 2 volume additions/d of overlying water. Water renewals are

70

performed manually or using an automatic dispenser system. Overlying water should be
from a source that has been shown to support reproduction of C. tentans in culture.
Reconstituted water should be avoided when possible.

For routine testing, a total of 16 replicates, each containing 12 larvae are used for
each treatment. Assignment of beakers for the 16 replicates is as follows: On test day -l
12 replicates are set up of which 4 will be used for the 20 (1 growth and survival endpoints
and 8 for determination of emergence and reproduction. Male C. tentans typically begin
emerging 4 to 7 d before females. To permit overlap of breeding individuals, an additional
4 beakers are set up on test day 10 for production of auxiliary males, which are thus
available during the prime female emergence period for each sediment treatment. Midges
in each test chamber are fed 1.5 mL of a 4 g/L Tetramin suspension daily. Endpoints
monitored include 20 d survival and growth, emergence, reproduction, and egg viability.
Specialized Equipment

Specialized equipment required for this test includes emergence traps,
reproduction/oviposit (R/O) chambers, an aspirator, and an adult collector dish.
Emergence traps are constructed from 120 mL plastic cups cups with the bottom removed
and replaced with a disc of ﬁberglass window screen. The traps ﬁt over the rim of the 300
mL beakers and trap the adult midges as they emerge. The emergence traps are also used
as R/O chambers. To serve in this application, the chambers are placed in a 100 x 20 mm
petri dish containing approximately 50 mL of culture water. At the time of the ﬁrst
emergence in each replicate, a corresponding R/O unit is initiated and placed on the shelf
within the test system for monitoring (a single R/O chamber is used to hold all adults that

emerge from a replicate).

71

 

An aspirator is constructed to aid in the collection and transfer of adults. The
aspirator consists of a 60 cc syringe with detachable aspirator unit. The barrel end of the
syringe is cut off and a retainer, used to hold the emergence traps and R/O units, is
attached. The retainer is constructed from a 7 cm diameter plastic lid, obtained from a
270 mL widemouth glass jar and a no. 10.5 rubber stopper. A hole is bored through the
center of both the lid and rubber stopper and the lid is glued onto the stopper with silicone
caulking. The open end of the syringe is then pushed through the hole of the
retainer/rubber stopper unit such that the open edge of the syringe is aligned with the
bottom of the inverted lid. A detachable tubing unit can then be ﬁxed on the open end of
the syringe for collection of midge adults. This tubing unit is made of a no. 5.5 rubber
stopper into which two 7 mm diameter holes are bored. A ﬁre-polished glass collector
tube (6 mm ID) 16 cm in length is placed into one of the two holes while a 4 cm length of
the same glass tubing is placed into the other hole. Both tubes should be ﬂush with the
bottom of the rubber stopper after insertion. A disk cut of ﬁberglass window screen is ﬁt
over the lower end of the short glass tube to prevent adults from being sucked into the
inhalant siphon of the aspirator. A 70 cm length of tygon tubing (6 mm ID, 9.5 mm OD)
is attached onto the opposite end of the short glass piece.

An adult collector dish, used in conjunction with the collector/transfer apparatus, is
constructed from a 100 x 20 mm petri dish. At the center of this dish, a 2.54 cm access
hole is drilled and covered with ﬁberglass window screen attached with silicone caulking.
Two slits within the screen, cut at 90° angles in the form of an "x" provide access (using
the collector tube) to the emergence traps or R/O units.

Collection of Egg Masses and Hatching of Eggs

72

Cultures should be maintained so that a minimum of 12 egg masses are produced
on day -3. This will require the collection of approximately 25 female and 10 male adult
midges on day -4. On day -3, 8 of the most uniform egg masses should be separated from
the total collected, transferred to a petri dish with culture water, and incubated at 23°C.
Hatching typically begins around 48 h post-oviposition and the majority of larvae leave the
egg masses 24 h after the ﬁrst hatch.

Hatching of eggs should be complete by about 72 h. After the ﬁrst 24 h period
with larvae hatched, transfer the egg masses from the incubation dish to another dish with
clean test water. Larvae that have already left the egg mass in the ﬁrst incubation dish are
discarded in order to reduce the age range of individuals used in the test. The action of
transferring the egg case stimulates the remaining larvae to leave the egg mass within a
few hours. These larvae are used to start the test.

Preparation of Test Chambers

The day before the test is started (day -1) each sediment should be thoroughly
homogenized and added to the test chambers. Sediment homogenization is best
performed by mixing the sediments in a bucket with a drill-mounted stainless steel paddle.
A stainless steel spoon should then be used to transfer 100 mL of sediment into each
beaker. Any sediment which is accidentally smeared on the sides of the beaker above the
100 mL mark should be removed with a paper towel. Otherwise this sediment may serve
as refuge for the invertebrates if they try to avoid the main mass of sediment in the bottom
of the beaker.

Overlying water is added to the chamber in a manner that minimizes suspension of

sediment. This is best accomplished by slowly pouring the water onto a Styrofoam disk

73

placed on top of the sediment to dissipate the force of the water. Renewal of overlying
water is started on day -1 by performing a water exchange after all replicates for the test
have been set up. A test begins when the organisms are added to the test chambers (day
0).
Renewal of Overlying Water

Two volume additions of overlying water should be delivered to each test chamber
daily over the course of the test. Routine water quality parameters will be measured at
regular intervals during the test (see below). The number of replacements per day should
be increased if hardness, alkalinity or ammonia concentrations in the overlying water vary
by more than 50% among treatments, or if oxygen drops to less than 30% saturation.
Although contaminant concentrations are reduced in the overlying water in water-renewal
tests, organisms in direct contact with sediments will generally still receive a substantial
proportion of a contaminant dose directly from either the whole sediment or from the
interstitial water. The water delivery system should be calibrated and run before a test is
started to verify that the system is functioning properly.
Acclimation of Test Organisms and Placement into Test Chambers

Test organisms must be cultured and tested at 23°C in the same test water.
However, if a sufﬁcient supply of culture water is not available for the exposure,
acclimation of test organisms to the test water is not required. Performance standards for
this test are calibrated to the response of midges at 23°C. If the test must be run at a
different temperature, then the organisms should be acclimated at a rate of 1°C every 2 h

to prevent therrna] shock. In addition, testing at temperatures other than 23°C need to be

74

preceded by studies to determine expected performance under other than standard
conditions.

Test organisms should be disturbed as little as possible. To start a test, larvae are
collected with a Pasteur pipette from the bottom of the incubation dish with the aid of a
dissecting microscope. Test organisms are pipetted very gently directly into the overlying
water of the exposure chambers. Care should be taken to release them under the surface
of the water. If possible, all larvae needed for the test should be placed into beakers
within 4 h of transfer of egg masses from the ﬁrst to the second incubation dish. If all
emerged larvae have been transferred from the incubation dish to the exposure chambers,
it may be necessary to wait for additional larvae to leave their egg masses. To minimize
this possibility, the test should be initiated with a sufﬁcient number of egg masses of
similar age.

Feeding

Each beaker receives a daily addition of 1.5 mL of Tetramin (4 g/L, note it has
recently been suggested that this volume be reduced to 1.0 mL/day to reduce the chance
of DO sags; USEPA 1999). Suspensions of food should be thoroughly mixed before
aliquots are taken. If excess food collects on the sediment, a fungal or bacterial growth
may develop on the sediment surface, in which case feeding should be suspended for one
or more days. A drop in dissolved oxygen below 2.5 ppm during a test may indicate that
the food added is not being consumed. Feeding should be suspended for the amount of
time necessary to increase the dissolved oxygen concentration. If feeding is suspended in
one treatment, it should be suspended in all treatments.

Overlying Water Quality

75

All chambers should be checked daily and observations made to assess test
organism behavior such as sediment avoidance. However, monitoring effects on
burrowing activity of test organisms may be difﬁcult because the test organisms are often
not visible during the exposure. The operation of the exposure system should be
monitored daily.

Conductivity, hardness, alkalinity, and ammonia should be measured in all
treatments at the beginning of the test and at least weekly until the end of the test. DO
and pH should be taken at the beginning of a test and at least three times a week until the
end of the test. Overlying water should be sampled just before water renewal from about
1 to 2 cm above the sediment surface using a pipet. It may be necessary to pool water
samples from individual replicates. The pipet should be checked to make sure no
organisms are removed during sampling of overlying water.

Routine chemistries on day 0 should be taken before organisms are placed in the
test beakers. DO and pH can be measured directly in the overlying water in an effort to
accurately measure concentrations of DO. The DO probe should be inspected between
samples to make sure that organisms are not attached and should be rinsed between
samples to minimize cross contamination.

It is likely that DO concentrations will drop signiﬁcantly during the test. Based on
tests run at the EPA and elsewhere, concentrations above 1.5 ppm do not seem to have an
effect on midge survival and development. However, if the DO drops below 2.5 ppm for
more than one day, then water replacements should be increased or aeration of all
exposure chambers should begin. Occasional brushing of the outsides of the screens on

the exposure chambers will help maintain the exchange of water during renewals.

76

Temperature should be checked at least daily in at least one exposure chamber
from each treatment. The temperature of the water bath or the exposure chamber should
be continuously monitored with a max/min thermometer. The daily mean test temperature
must be 23 1|; 1°C. The instantaneous temperature must always be 23 i 3°C.

Monitoring Survival and Growth

At 20 d, 4 of the initial 12 replicates are randomly selected for use in growth and
survival measurements. A #40 sieve (425 um mesh) should be used to isolate larvae from
the sediment. Any immobile organisms isolated from the sediment surface or from sieved
material should be considered dead. Often midge larvae tend to lose their coloration
within 15 to 20 min of death and may become rigidly elongate. Surviving larvae are kept
separated by replicate for weight measurements; if pupae are recovered, these are included
in survival data but not included in the growth data.

Traditionally, dry weight (DW) has been used for the larval growth endpoint on
midge tests. However, it has been shown that the grain size of sediments inﬂuences the
amount of sediment that C. tentans larvae ingest and retain in their gut. As a result, in
fmer-grain sediments, a substantial portion of the measured dry weight may be comprised
of sediment rather than tissue. While this may not represent a strong bias in tests with
identical grain size distributions in all treatments, most ﬁeld assessments are likely to have
varying grain size among sights. This will likely create differences in dry weight among
treatments that are not reﬂective of true somatic growth. For this reason, it has been
suggested that weight of midges should be measured as ash-free dry weight (AFDW)
instead of dry weight. AFDW will more directly reﬂect actual differences in tissue weight

by reducing the inﬂuence of sediment in the gut.

77

To ﬁnd AFDW, all living larvae per replicate are combined in a pre-ashed

aluminum weigh boat and dried at 60°C for 24h. The samples should be cooled in a
dessicator and weighed to the nearest 0.01 mg to obtain mean weights per surviving
organism per replicate. When weighing, remove only 4 weigh boats from the dessicator at
a time and weigh them When done, remove 4 more and so on until all are weighed. The
dry pans will absorb water out of the air and increase in weight, so they should be weighed
as soon as possible after removal from the dessicator. The dried larvae in the pan are then
ashed at 550°C for 2h. The pan with the ashed larvae is then re-weighed and the tissue
mass of the larvae is determined as the difference between the weight of the dried larvae
plus pan and the weight of the ashed larvae plus pan. In rare instances, where
preservation is required, an 8% sugar-formalin solution can be used to preserve samples.
The sugar-formalin solution is made by adding 120 g of sugar to 80 mL of formalin, and
bringing the solution up to 1 L with DI water. This solution is mixed 1:1 with sample
water to preserve organisms. Sugar formalin is preferable to alcohol for preservation
because it does not cause as much shrinkage as EtOH.
Monitoring Emergence

Emergence traps are placed on the reproductive replicates on day 20. Emergence
traps for the auxiliary beakers are added at the corresponding 20 (1 time interval for those
replicates. At 23°C, emergence in reference sediment typically begins on or about day 23
and continues for about 2 weeks. In contaminated sediments, the emergence period may
be extended by several weeks.

Two categories are recorded for emergence: complete emergence and partial

emergence. Complete emergence occurs when an organism has shed the pupal exuviae

78

completely and escapes the surface tension of the water. If complete emergence has
occurred but the adult has not escaped the surface tension of the water, the adult will
usually die within 24 h. Therefore, 24 h should elapse before this death is recorded.
Partial emergence occurs when an adult has only partially shed the pupal exuviae. These
adults will also die, which can be recorded after 24 h. Pupae at the sediment surface or
the air-water interface may emerge successfully during the 24 h period. However,
cannibalism of sediment-bound pupae by larvae may also occur. Data are recorded on the
attached data sheets.
Collecting Adults for Reproduction

Adults are collected daily from individual traps using the aspirator and collector
dish. With the collector dish nearby, the emergence trap is quickly moved from the beaker
onto the dish. With the syringe plunger fully drawn, the glass collector tube is inserted
through the screened access hole of the collector dish and the adults gently aspirated into
the syringe barrel. Aspirated adults can easily be seen through the translucent plastic of
the syringe. The detachable portion of the aspirator unit is then replaced with a
reproduction/oviposit (RIO) chamber. This exchange can be facilitated by placing the
thumb of the hand holding the syringe over the barrel entry port until the RIO chamber is
in place. With the RIO chamber in place, and the plunger on the table top, the barrel of
the syringe is pushed gently downward which forces the adults to move up into the RIO
unit. Adults remaining on the transfer apparatus may be prodded into the RIO chamber by
gently tapping the syringe. The transfer process is completed by quickly moving the RIO

chamber to a petri dish containing overlying water. At all times during the transfer

79

 

process, it is important to ensure that the adults are stationary to minimize the possibility
of escape.

At about day 33 until the end of the test, the auxiliary males may be needed to
support reproduction in females. Males that emerge from the auxiliary male replicates are
transferred to individual inverted RIO chambers. Each male may be used for mating with
females from corresponding treatments for up to 5 d. Males may be used for breeding
with more than one new emergent female. Males from a different replicate within the
same sediment treatment may be paired with females of replicates where no males have
emerged. Record data on data sheets.

Monitoring Reproduction

Each RIO unit is checked daily for dead adults and egg cases. Dead organisms are
removed. In situations where many adults are contained within an RIO chamber, it may be
necessary to assume that a dead adult is the oldest male or female in that replicate for the
purpose of recording time to death. To remove dead adults and egg cases from the RIO
chamber, one side of the chamber is carefully lifted just enough to permit the insertion of a
transfer pipet or tweezers.

For each emerged female, at least one male, obtained from the corresponding
reproductive replicate, from another replicate of that treatment, or from the auxiliary male
beakers, is transferred into the RIO unit using an aspirator. Females generally remain
sexually receptive up to 3 d if they have not already mated. Almost all females will
oviposit within 1 d of fertilization; however, a few will require as long as 72 h to oviposit.
A female will lay a single primary egg case, usually in the early morning. A second,

generally smaller egg case may be laid; however these second egg cases are prone to

80

 

fungus and the viability of embryos is typically poor. These second egg cases do not need
to be counted, or recorded, and the numbers of eggs are not included in the egg counts
because eggs in the secondary egg cases typically have low viability.

Counting Eggs, Egg Case Incubation, and Hatch Determination

Primary egg cases from the RIO chamber are transferred to a separate and
corresponding petri dish to monitor incubation and hatch. The number of eggs should be
estimated in each egg case by using the ring count method: i) For each egg case, the
mean number of eggs in ﬁve rings is determined; ii) these rings should be selected at
about equal distances along the length of the egg case; iii) the number of eggs/ring
multiplied by the number of rings in the egg case will provide an estimate of the total
number of eggs. This can be done in about 5 min or less for each egg case. According to
the EPA (USEPA 1999), accuracy of estimating versus a direct count method is very
close (about 95%). The ring method is best suited to the symmetrically "C" shaped egg
masses.

When the integrity of an egg case precludes estimation by the ring method (egg
case is convoluted or distorted), the eggs must be counted directly. Each egg case is
transferred to a petri dish with 2N sulfuric acid and left overnight. The acid dissolves the
gelatinous matrix surrounding the eggs but does not affect the structural integrity of the
eggs themselves. After digestion, the eggs can be counted by collecting them with a
Pasteur pipet and spread across a microscope slide for counting under a dissecting
microscope. Counting can be simpliﬁed by drawing a grid on the underside of the slide.
Alternatively, they may be counted by removing them from the petri dish with a Pasteur

pipet and counting the eggs with a push-button counter as they are collected into the

81

pipet. The direct count method requires a minimum of 10 min to complete and does not
permit determination of hatching success.

Following estimated egg counts (ring count method), each egg case is transferred
to a 60 x 15 mm plastic petri dish containing 15 mL overlying water and incubated at 23°C
until hatching is complete. Although the time required to initiate hatching at this
temperature is about 2 d, the period of time required to bring about complete hatch may
be as long as 6 d. Therefore, hatching success is determined after 6 d of incubation.
Hatching success is determined by subtracting the number of unhatched eggs remaining
after the 6 (1 period from the number of eggs originally estimated for that egg case.
Unhatched eggs either remain in the gelatinous egg case or are distributed on the bottom
of the petri dish. Note that the unhatched eggs may decompose if the incubation
temperature rises much above 23°C, so it is important that this temperature be maintained.
Test Termination

The quality of the sediments will determine the length of this test. In clean
sediments, the test typically requires 40 to 50 d for completion. However, test duration
will increase in the presence of environmental stressors, which act to reduce growth and
delay emergence. Where a strong gradient of sediment contamination exists, emergence
patterns between treatments will likely become asynchronous, in which case we have
ended the test in our lab when there is no emergence from the control treatments for 7 d.
However, it should be noted that the EPA has more recently suggested ended each
treatment separately at a point which is determined by the performance of that treatment
(USEPA 1999). If it is decided that each treatrmnt will be ended separately, a treatment

is terminated when no further emergence is recorded over a period of 7 d. In either case,

82

at test termination, all beakers of the treatment are sieved through a #40 mesh screen (425
um) to recover remaining larvae, pupae, or pupal cases. If no emergence is recorded in a
treatment at any time during the test, the treatment is ended once emergence in the control
sediment has ended. In this case, you have probably chosen the wrong test for that
sediment, and a short-term exposure with survival and growth might be more appropriate.
Interpretation of Results
Data Analyses

Endpoints measured in the C. tentans test include survival, growth, emergence,
and reproduction. This section describes some general information from the EPA, the
USFWS, etc. that will help in interpretation of results.
Age Sensitivity

Midges are perceived to be relatively insensitive organisms in toxicity assessments.
This conclusion is based in great part on using the survival and growth of third and fourth
instar animals in short-term exposures. The ﬁrst and second instars of chironomids are
more sensitive to contaminants than the third or fourth instars. For example, ﬁrst instar C.
tentans larvae were 6 to 27 times more sensitive than fourth instar larvae to acute copper
exposure (Nebeker et a1. 1984). Sediment tests must be started with uniform age and size
midges because of the dramatic differences in sensitivity of midges by age.
Physical Characteristics of Sediment
Grain Size and Organic Matter

Larvae of C. tentans appear to be tolerant of a wide range of particle size
conditions in substrates. Survival does not appear to be affected by grain size in natural

sediments, sand substrates, or formulated sediments in both 10 d and long-terrn exposures.

83

Some studies have shown that growth of C. tentans is weakly correlated with sediment
grain size composition. However, other studies by Paul Sibley at the University of Guelph
found that the correlation between grain size and larval growth disappeared after
accounting for inorganic material contained within larval guts, and he concluded that
growth of midges was not related to grain size composition in either natural sediment or
sand substrates. Avoiding confounding inﬂuences of gut contents on weight is the reason
why AFDW is now recommended, as described above. Sibley also found that emergence,
reproduction (in eggs/female), and batch success were not affected by the particle size
composition of substrates in chronic tests with C. tentans. The organic matter content of
sediments does not appear to affect survival of C. tentans larvae, but the type and quantity
of OM may be important for growth and should be considered if sediments tested are from
different sources.
Inﬂuence of Indigenous Organisms

Survival of C. riparius was not reduced in the presence of a moderate number of
oligochaetes in sediment samples, though growth was reduced when high numbers of
oligochaetes were placed in the samples (Reynoldson et al. 1994). The presence of
predators may also inﬂuence the response of test organisms in sediment (Ingersoll and
Nelson, 1990).
Relative Endpoint Variability

According to the EPA, based on coefﬁcient of variation (CV) determined from
their standard control sediment (West Bearskin Lake), the following variability has been

documented for the various endpoints in the C. tentans life cycle test: Survival (<20%),

84

growth as dry weight (<15%), emergence (<30%), reproduction as mean eggs/female
(<20%), percent hatch (<10%).
Relative Endpoint Sensitivity

Measurement of sublethal endpoints (e.g. growth) can often provide a lot of
information compared to acute endpoints (survival). Sibley et a]. (1997) found that larval
C. tentans exposed to a gradient of food stress did not experience signiﬁcant effects on
survival, yet did experience a signiﬁcant reduction in growth and reproduction. Further,
the proportion of larvae hatching in Sibley's study was high (>80%), and not
systematically related to treatment, suggesting that percent hatch may be a relatively

insensitive endpoint. We have come to much the same conclusions in our own lab studies.

85

 

Table 1. Test conditions for conducting a whole-life cycle assay with Chironomus

 

 

 

tentans.

Parameter Conditions

1. Test Type Whole-sedirnent toxicity test with renewal of
overlying water

2. Temperature 23il°C

3. Illuminance About 500 to 1000 lux

4. Photoperiod 16L:8D

5. Test chamber 300 mL high-form lipless beakers

6. Sediment volume 100 mL

7. Overlying water volume 175 mL -'

8. Age of organisms < 24 h old larvae

9. Number of organisms/chamber 12

10. Number of replicate 16 (12 at day -1 and 4 for auxiliary males on day F

chambers/treatment 10). I

11. Feeding Tetramin food, fed 1.5 mL (4 g/L stock) daily to
each test chamber starting on day -1

12. Aeration None unless D0 in overlying water drops below
2.5 mg/L

13. Overlying water Preferably culture water. Well water, surface
water, or site water may also be used.

14. Test chamber cleaning Occasionally the outside of the screen may be
brushed gently with a toothbrush.

15. Overlying water quality Hardness, alkalinity, conductivity, and ammonia
at the beginning and end of a test. Temperature
daily. Dissolved oxygen (DO) and pH three
times/week. Concentrations of DO should be
measured more often if DO drops more than 1
mg/L since the previous measurement.

16. Test duration About 40 to 50 d

17. Endpoints 20 day survival and growth; female and male
emergence, adult mortality, number of egg
masses deposited, number of eggs produced, and
number of hatched eggs.

18. Test acceptability Mean weights of organisms in controls at 20 (1

must be at least 0.6 mg/individual as dry wt. Or
0/48 mg/ind. as ash-free dry weight and should
be at least 70% survival.

86

Table 2. General activity schedule for conducting a whole-life cycle assay with
Chironomus tentans.
Day Activity

-4 Start reproduction ﬂask with cultured adults (1 :3 malezfemale ratio).
-3 Collect egg masses and incubate at 23°C.
-1 Check egg cases for batch and development. Add sediment into each test chamber,

place chambers into exposure system, and start renewing overlying water. Add 1.5
mL Tetramin suspension to each chamber.

0 Measure total water quality (pH, temperature, DO, hardness, alkalinity, conductivity,
ammonia). Transfer all egg cases to a crystallizing dish containing control water.
Discard larvae that have already left the egg cases in the incubation dishes. Add 1.5
mL Tetramin suspension to each chamber before adding larvae. Add 12 larvae to
each replicate beaker.

1- Add 1.5 mL Tetramin suspension to each beaker. Measure temperature daily.

End Measure pH and DO three times a week. If DO declines more than 1 mg/L between
measurements, increase the number of water replacements or aerate all beakers.
Measure hardness, alkalinity, conductivity, ammonia weekly.

6 For auxiliary male production, start reproduction ﬂask with culture adults.

7-10 Set up schedule for auxiliary male beakers (4 replicates/treatment) same as that
described above for day -3 through 0.

19 Ash weigh boats at 550°C for 2h in preparation for weight measurements. Store boats
in a dessicator before use.
20 Randomly select 4 replicates from each treatment and sieve the sediment to recover

larvae for growth and survival determination. Pool all living larvae per replicate and
dry the sample at 60°C for 24 h. Install emergence traps on each of the remaining
reproduction replicate beakers.

21 Samples with dried larvae are brought to room temperature in a dessicator and
weighed to the nearest 0.01 mg. The pans are then ashed at 550°C for 2 h. Pans are
then reweighed and the tissue mass of the larvae determined as the difference between
the weight of thedried larvae plus pan and the weight of the ashed larvae plus pan.

23- On a daily basis, record emergence of males and females, pupal, and adult mortality,

End and time to death for previously collected adults. Each day, transfer adults from each
replicate to a corresponding reproduction/ovoposition (RIO) chamber. Transfer each
primary egg case from the RIO chamber to a corresponding petri dish to monitor
incubation and hatch. Record each egg case oviposited number of eggs produced, and
number of hatched eggs. If it is difﬁcult to estimate the number of eggs in an egg
case, used a direct count to determine the number of eggs; however the hatchability
data will not be obtained for this egg case.

30 Place emergence traps on auxiliary male replicate beakers.

33- Transfer males emerging from the auxiliary male replicates to individual inverted petri

End dishes. The auxiliary males are used for mating with females from corresponding
treatments from which most of the males are likely to have already emerged or in
which no males emerged.

40- After 7 d of no recorded emergence in the control treatment, the test can be terminated

End by sieving the sediment to recover larvae, pupae, or pupal exuviae. Alternatively,
each treatment can be ended independently when no emergence has occurred in that
treatment for 7 d.

87

Table 3. Test acceptability requirements for a whole-life cycle assay with Chironomus
tentans.

 

B. The following performance criteria for the whole life stage test with C. tentans
have been recommended by the EPA:

 

H
o

Tests must be started with less than 1 (1 old larvae.

2. Average survival of midges in the control sediment must be greater than or
equal to 70% at the end of the test. Average dry weights should be 2_0.6
mg/individual, or _>_ 0.48 mg/individual AFDW.

3. Based on round-robin and EPA lab results, larval survival is usually > 76%,
adult survival is > 96%, and emergence is > 60%. Time to death after
emergence is < 6.5 d for males and < 5.1 d for females. The mean number of
eggs per egg mass range from 906 to 1107 and the percent hatch ranges
from 91 to 96%.

4. Hardness, alkalinity, and ammonia in the overlying water typically should not

vary by more than 50% during the test.

 

. Additional Requirements

 

B

1 All organisms in a test must be from the same source.

2. Negative-control sediment must be included in a test.

3 Test organisms should be cultured and tested at 23°C.

4 The mean of the daily test temperature must be 23i1°C. The instantaneous
temperature must be 23;3°C.

88

APPENDIX 5: STANDARD PROCEDURE FOR A WHOLE LIFE CYCLE
ASSAY WITH THE AMPHIPOD H YALELLA AZTECA

Aquatic Toxicology Laboratory Standard Operating Procedure
Department of Zoology
Michigan State University

Title: Whole life cycle assay with the amphipod Hyalella azteca.

Description of Method:

The sediment exposure portion of this assay starts with 7- to 8-d-old amphipods.

On Day 28, amphipods are isolated from the sediment and placed in water-only chambers

where reproduction is measured on Day 35 and 42. Typically, amphipods are ﬁrst in

amplexus at about Day 21 to 28 with release of the ﬁrst brood between Day 28 to 42.

Endpoints measured include survival (Day 28, 35 and 42), growth (as length or weight

measured on Day 28 and 42), and reproduction (number of young/female produced from

Day 28 to 42).

Referenced Documents:

Benoit, D.A., Phipps, GA., and Ankley, GT. 1993. A sediment testing intermittent
renewal system for the automated renewal of overlying water in toxicity tests with
contaminated sediments. Water Res. 27:1403-1412.

Ingersoll, CG, and MK. Nelson. 1990. Testing sediment toxicity with Hyalella azteca
(Amphipods) and Chironomus riparius (Diptera). In: W.G. Landis and W.H. van
der Schalie (eds), Aquatic Toxicology and Risk Assessment, 13th volume. ASTM

STP 1096. Philadelphia, PA, pp. 93-109.

89

Ingersoll, C.G., E.L. Brunson, F.J. Dwyer, D.K. Hardesty, and NE. Kemble. 1998. Use of
sublethal endpoints in sediment toxicity tests with the amphipod Hyalella azteca.
Environ. Toxicol. Chem. 17: 1508-1523.

Kubitz, J.A., J.M. Besser, and JP. Giesy. 1996. A two-step experimental design for a
sediment bioassay using growth of the amphipod Hyalella azteca for the test
endpoint. Environ. Toxicol. Chem. 15: 1783-1792.

Reynoldson, T.B., K.E. Day, C.Clarke, and D. Milani. 1994. Effect of indigenous animals
on chronic endpoints in freshwater sediment toxicity tests. Environ. Toxicol.
Chem. 10:973-977.

USEPA. 1999. Methods for measuring the toxicity and bioaccumulation of sediment-
associated contaminants with freshwater invertebrates: Second edition.
Environmental Research Laboratory, Duluth, Minnesota. Draft document.

Procedure:

Test Conditions
Conditions for evaluating sublethal endpoints in a sediment toxicity test with H.

azteca are summarized in Table l. A general activity schedule is outlined in Table 2. The

42-d sediment toxicity test with H. azteca is conducted at 23°C with a 16L:8D

photoperiod at an illuminance of about 500 to 100 lux. Test chambers are 300-mL high-

form lipless beakers containing 100 mL of sediment and 175 mL of overlying water. Ten
amphipods in each test chamber are fed 1.0 mL of yeast, cerophyll, trout chow suspension

(YCT) daily. Each test chamber receives 2 volume additions/d of overlying water. Water

renewals are performed manually or using an automatic dispenser system. Overlying

water should be from a source that has been shown to support reproduction of H. azteca

90

in culture. In our lab we usually use 30% well water and 70% reverse osmosis or
ultrapure water. Reconstituted water should be avoided when possible.

The number of replicates and concentrations tested depends in part on the
signiﬁcance level selected and the type of statistical analysis. A total of 12 replicates, each
containing ten 7- to 8-d-old amphipods, are usually tested for each sample. Starting the
test with substantially younger or older organisms may compromise the reproductive
endpoint. For the total of 12 replicates the assignment of beakers is as follows: 12
replicates are set up on Day -1 of which 4 replicates are used for 28-d growth and survival
endpoints and 8 replicates for measurement of survival and reproduction on Day 35, and
survival, reproduction, and growth on Day 42.

Preparation of Test Chambers

The day before the test is started (day -1) each sediment should be thoroughly
homogenized and added to the test chambers. Sediment homogenization is best
performed by mixing the sediments in a bucket with a drill-mounted stainless steel paddle.
A stainless steel spoon should then be used to transfer 100 mL of sediment into each
beaker. Any sediment which is accidentally smeared on the sides of the beaker above the
100 mL mark should be removed with a paper towel. Otherwise this sediment may serve
as refuge for the invertebrates if they try to avoid the main mass of sediment in the bottom
of the beaker.

Overlying water is added to the chamber in a manner that minimizes suspension of
sediment. This is best accomplished by slowly pouring the water onto a Styrofoam disk
placed on top of the sediment to dissipate the force of the water. Renewal of overlying

water is started on day -l by performing a water exchange after all replicates for the test

91

have been set up. A test begins when the organisms are added to the test chambers (day
0).
Renewal of Overlying Water

Two volume additions of overlying water should be delivered to each test chamber
daily over the course of the test. Routine water quality parameters will be measured at
regular intervals during the test. The number of replacements per day should be increased
if hardness, alkalinity or ammonia concentrations in the overlying water vary by more than
50% among treatments, or if oxygen drops to less than 30% saturation. Although
contaminant concentrations are reduced in the overlying water in water-renewal tests,
organisms in direct contact with sediments will generally still receive a substantial
proportion of a contaminant dose directly from either the whole sediment or from the
interstitial water. The water delivery system should be calibrated and run before a test is
started to verify that the system is functioning properly.
Acclimation of Test Organisms and Placement into Test Chambers

Test organisms must be cultured and tested at 23°C in the same test water.
However, if a sufﬁcient supply of culture water is not available for the exposure,
acclimation of test organisms to the test water is not required. Performance standards for
this test are calibrated to the response of amphipods at 23°C. If the test must be run at a
different temperature, then the organisms should be acclimated at a rate of 1°C every 2 h
to prevent thermal shock. In addition, testing at temperatures other than 23°C need to be
preceded by studies to determine expected performance under other than standard

conditions.

92

Test organisms should be disturbed as little as possible. To start a test, amphipods
are collected with a Pasteur pipette from the bottom of a crystallizing dish with the aid of
a dissecting microsc0pe. Test organisms are pipetted directly into the overlying water of
the exposure chambers. Care should be taken to release them under the surface of the
water.

Feeding

Each beaker receives a daily addition of 1.0 mL of YCT. Suspensions of food
should be thoroughly mixed before aliquots are taken. If excess food collects on the
sediment, a fungal or bacterial growth may develop on the sediment surface, in which case
feeding should be suspended for one or more days. A drop in dissolved oxygen below 2.5
mg/L during a test may indicate that the food added is not being consumed. Feeding
should be suspended for the amount of time necessary to increase the dissolved oxygen
concentration. If feeding is suspended in one treatment, it should be suspended in all
treatments.

Overlying Water Quality

All chambers should be checked daily and observations made to assess test
organism behavior such as sediment avoidance. However, monitoring effects on
burrowing activity of test organisms may be difficult because the test organisms are often
not visible during the exposure. The operation of the exposure system should be
monitored daily.

Conductivity, hardness, alkalinity, and ammonia should be measured in all
treatments at the beginning of the test and at least weekly until the end of the test. Total

water quality should also be measured at the beginning and end of the reproductive phase

93

(day 29 and day 42). Conductivity should be measured weekly and DO and pH three
times/week. Overlying water should be sampled just before water renewal from about 1
to 2 cm above the sediment surface using a pipet. It may be necessary to pool water
samples from individual replicates. The pipet should be checked to make sure no
organisms are removed during sampling of overlying water.

Routine chemistries on day 0 should be taken before organisms are placed in the
test beakers. DO and pH can be measured directly in the overlying water in an effort to
accurately measure concentrations of DO. The DO probe should be inspected between
samples to make sure that organisms are not attached and should be rinsed between
samples to minimize cross contamination.

It is likely that DO concentrations will drop signiﬁcantly during the test, but should
not run below 2.5 mg/L. If the DO dr0ps below 2.5 ppm for more than one day, then
water replacements should be increased or aeration of all exposure chambers should begin.
Occasional brushing of the outsides of the screens on the exposure chambers will help
maintain the exchange of water during renewals.

Temperature should be checked at least daily in at least one exposure chamber
from each treatment. The temperature of the water bath or the exposure chamber should
be continuously monitored with a max/min thermometer. The daily mean test temperature
must be 23 i 1°C. The instantaneous temperature must always be 23 i 3°C.

Test Termination

Endpoints monitored include 28-d survival and growth of amphipods and 35-d and

42-d survival, growth, and reproduction (number of young/female) of amphipods. On day

28, 4 of the replicate beakers/sediment are sieved with a #40 mesh sieve (425-um mesh;

94

U.S. standard size sieve) to remove surviving amphipods. Animals in the water column or
on the surface of the sediment can be pipetted from the beaker before sieving the
sediment. The sediment in each beaker should be sieved in two separate aliquots (i.e.
most of the amphipods will probably be found in the surface aliquot). Immobile organisms
isolated from the sediment surface or from sieved material should be considered dead.
Surviving amphipods from these 4 replicates are preserved in separate vials containing 8%
sugar formalin solution if length of amphipods is to be measured. We do not always do
length in our lab because of the ﬂeeting availability of a digitizing microscope. The sugar
formalin solution is prepared by adding 120 g of sucrose to 80 mL of formalin which is
then brought to a volume of 1 L using deionized water. This stock solution is mixed with
an equal volume of deionized water when used to preserve organisms. Sugar formalin is
preferable to alcohol for preservation because it does not cause as much shrinkage as
EtOH.

Growth of amphipods can be reported as either length or weight. Amphipod body
length (+0.1 m) can be measured from the base of the ﬁrst antenna to the tip of the third
uropod along the curve of the dorsal surface. Dry weight of amphipods in each replicate
can be determined on day 42 and for the archived samples on day 28. If both weight and
length are to be determined, weight should be measured after length on the preserved
samples. Dry weight of amphipods can be determined by: (1) transferring the archived
amphipods from a replicate out of the sugar formalin solution into a crystallizing dish; (2)
rinsing amphipods with deionized water; (3) transfering these rinsed amphipods to a pre-
weighed aluminum micro-weigh boat (1 cm in diameter); (4) drying these samples for 24 h

at 60 degrees C; and (5) weighing the pan and dried amphipods on a balance to the nearest

95

0.01 mg. Average dry weight of individual amphipods in each replicate is calculated from
these data. Due to the small size of amphipods, caution should be taken during weighing
(10 dried amphipods after a 28-d sediment exposure may weigh less than 2.5 to 3.5 mg).
Weigh pans need to be carefully handled using powderless gloves and the balance should
be calibrated with standard weights with each use. Use of small, homemade aluminum
pans (e.g., 1 cm in diameter) will help reduce variability in measurements of dry weight.
These may be constructed out of sheets of aluminum foil.

Historically, dry weight has been used for the growth endpoint for C. tentans and
H. azteca. However, for C. tentans, this recommendation was changed to ash-free dry
weight (AFDW) instead of dry weight, with the intent of reducing bias introduced by gut
contents (Sibley et al. 1997). However, this recommendation was not extended to include
H. azteca because the ash content of H. azteca is not greatly decreased by purging
organisms in clean water before weighing, suggesting that sediment does not comprise a
large portion of the overall dry weight. In addition, using AFDW further decreases an
already small mass, potentially increasing measurement error.

On Day 28, the remaining 8 of the original 12 beakers/sediment are also sieved and
the surviving amphipods in each sediment beaker are placed in 300 mL water-only beakers
containing 150 to 275 mL of overlying water and a 5-cm x 5-cm piece of Nitex screen.
Each water-only beaker receives 1.0 mL of YCI' stock solution and about two volume
additions of water daily.

Reproduction of amphipods is measured on Day 35 and Day 42 in the water-only
beakers by removing and counting the adults and young in each beaker. On Day 35, the

adults are then returned to the same water-only beakers. Adult amphipods surviving on

96

Day 42 are preserved in sugar formalin. The number of adult females is determined by
simply counting the adult males (mature male amphipods will have an enlarged second
gnathopod) and assuming all other adults are females. The number of females is used to
determine number of young/female/beaker from Day 28 to Day 42. Growth can also be
measured for these adult amphipods.
Results Interpretation
Grain Size

Hyallela azteca tolerate a wide range in sediment grain size and organic matter.
During development of this test by the EPA,USFWS, and during the round robin assays,
no signiﬁcant correlations were observed between the survival, growth, or reproduction of
H. azteca and the physical characteristics of the sediment (grain size ranging from
predominantly silt to predominantly sand), TOC (ranging from 0.3 to 9.6%), water
content (ranging from 19 to 81%; Ingersoll et al. 1998). Additionally, no signiﬁcant
correlations were observed between these biological endpoints and water quality
characteristics (i.e., hardness, alkalinity, ammonia) of pore water or overlying water in the
sediments evaluated by Ingersoll et al. (1998). Weak trends were observed between
reproduction of amphipod percent clay, percent silt, and percent sand. The weak
relationship between the sediment grain size and reproduction in that study may have been
due to the fact that samples with higher amounts of sand also had higher concentrations of
organic contaminants compared to other samples evaluated in Ingersoll et a]. (1998).
Inﬂuence of Indigenous Organisms

Survival of H. azteca in 28—d tests was not reduced in the presence of oligochaetes

in sediment samples (Reynoldson et al. 1994). However, growth of amphipods was

97

reduced when high numbers of oligochaetes were placed in a sample. Therefore, it is
important to determine the number and biomass of indigenous organisms in ﬁeld-collected
sediments in order to better interpret growth data (Reynoldson et a1. 1994). Furthermore,
presence of predators may also inﬂuence response of test organisms in sediment (Ingersoll
and Nelson 1990).
Number of Replicates

Although 8 replicates are recommended for evaluating reproduction in the long-
term sediment tests with H. azteca, there may be instances where either additional of
fewer replicates might be evaluated to monitor reproduction (Ingersoll et a1. 1998). For
example, Kubitz et al. (1996) recommended a two step process for assessing growth in
sediment tests with H. azteca. Using this process, a limited number of replicates would be
tested in a screening step. Samples identiﬁed as possibly affecting reproduction could then
be tested in a conﬁrmatory step with additional replicates. This two-step analysis
conserves laboratory resources and increases statistical power when needed to identify
sublethal effects. A similar approach could be applied to evaluate reproductive effects of
contaminants in sediment where a limited number of replicates could be initially tested to
evaluate potential effects. Samples identiﬁed as possibly toxic based on reproduction
could then be reevaluated using an increased number or replicates. It should be noted that
results of reproduction have been mixed, and the value of this endpoint is still under

consideration.

98

Table 1. Test conditions for conducting a whole-life cycle assay with Hyalella azteca.

 

 

Parameter Conditions

1. Test Type Whole-sedirnent toxicity test with renewal of
overlying water

2. Temperature 23:1°C

3. Illuminance About 500 to 1000 lux

4. Photoperiod 16L:8D

5. Test chamber 300 mL high-form lipless beakers

6. Sediment volume 100 mL

7. Overlying water volume 175 mL in the sediment exposure from day 0 to

8.
9.

10.

11.

12.

l3.

14.

15.

16.
17.

18.

Age of organisms

Number of organisms/chamber
Number of replicate
chambers/treatment
Feeding
Aeration
Overlying water

Test chamber cleaning

Overlying water quality

Test duration
Endpoints

Test acceptability

day 28 and 175-275 mL in the water only
exposure from day 28 to day 42

7 to 8 (1 old at the start of the test

10

12 (4 for 28 d survival and growth and 8 for 35
and 42 d survival, growth, and reproduction).
YCT food, fed 1.0 mL (1800 mg/L stock) daily
to each test chamber

None unless D0 in overlying water drops below
2.5 mg/L

Preferably culture water. Well water, surface
water, or site water may also be used.
Occasionally the outside of the screen may be
brushed gently with a toothbrush.

Hardness, alkalinity, conductivity, and ammonia
at the beginning and end of a test (day 0 and day
28). Temperature daily. Conductivity weekly.
Dissolved oxygen (DO) and pH three
times/week. Concentrations of DO should be
measured more often if DO drops more than 1
mg/L since the previous measurement.

42 d

28 day survival and growth; 35 day survival and
reproduction, 42 day survival, growth,
reproduction, and number of adult males and
females.

Minimum mean control survival of 80% on day
28 and suggested perfomance-based developed
following round-robin testing

 

99

Table 2. General activity schedule for conducting a whole-life cycle assay with Hyalella

azteca. .

 

Day
-8

-7
-6to-l
-1

0

lto 27

28

29 to 35

35

36 to 41

41

42

Activity

Isolate several hundred adult amphipods from culture and place into breeding
chambers.

Remove adults from breeding chambers and isolate <24h old amphipods

Feed and observe isolated amphipods.

Add sediment into each test chamber, place chambers into exposure system,
and start renewing overlying water

Measure total water quality (pH, temperature, DO, hardness, alkalinity,
conductivity, ammonia). Transfer ten 7 to 8 (1 old amphipods into each test
chamber. Release organisms under the surface of the water. Add 1.0 mL of
YCT into each test chamber.

Add 1.0 mL of YCT to each test beaker. Measrue temperature daily,
conductivity weekly, and DO and pH three time/wk. Observe and record
behavior of test organisms.

Measure temperature, DO, pH, hardness, alkalinity, conductivity and
ammonia. End the sediment exposure portion of the test by collecting the
amphipods with a #40 mesh sieve (425 um mesh). Use four replicates for
growth measurements: count survivors and preserve organisms in sugar
formalin for growth measurements. Eight replicates for reproduction
measurements: place survivors in individual replicate water-only beakers and
add 1.0 mL of YCT to each test beaker/d and 2 volume additions/d of
overlying water.

Feed daily. Measure temperature daily, conductivity weekly, DO and pH three
times a week. Measure hardness and alkalinity weekly. Observe behavior of
test organisms.

Record the number of surviving adults and remove offspring. Return adults to
their original individual beakers and add food.

Fee daily. Measure temperature daily, conductivity weekly, DO and pH three
times a week. Measure hardness and alkalinity weekly. Observe behavior of
test organisms.

Same as day 1. Measure total water quality (pH, temperature, DI, hardness,
alkalinity, conductivity, ammonia).

Record the number of surviving adults and offspring. Surviving adult
amphipods on day 42 are preserved in sugar formalin solution. The number of
adult males in each beaker is determined from this archived sample. This
information is used to calculate the number of young produced per female per
replicate from day 28 to day 42.

100

Table 3. Test acceptability requirements for a whole-life cycle assay with Hyalella azteca.

 

C. The following performance criteria for the whole life stage test with H. azteca have
been recommended by the EPA:

 

1. Age of H. azteca at the start of the test should be 7 to 8 (1 old. Starting a test with
substantially younger or older organisms may compromise the reproductive
endpoint.

2. Average survival of H. azteca in the control sediment on day 28 should be greater
than or equal to 80%.

3. Laboratories participating in round-robin testing reported after 28 d sediment

exposures in a control sediment (West Bearskin Lake), survival >80% for >88%
of the labs; length > 3.2 mm/individual for > 83% of the labs; and weight > 0.15
mg/individual for 67% of the labs. Reproduction from day 28 to day 42 was >2
young/female for 63% of the labs. Reproduction was more variable within and
among laboratories; hence, more replicates might be needed to establish statistical
differences among treatments with this endpoint.

4. Hardness, alkalinity, and ammonia in the overlying water typically should not vary
by more than 50% during the sediment exposure.

 

. Additional Requirements

 

B

1 All organisms in a test must be from the same source.

2. Negative-control sediment must be included in a test.

3 Test organisms should be cultured and tested at 23°C.

4 The mean of the daily test temperature must be 23il°C. The instantaneous
temperature must be 23;3°C.

 

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