CBP/TRS 89/93
December 1992
903R92017
Development of a Chronic
Sediment Toxicity Test
for Marine Benthic
Amphipods
December 1992
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225
.C54
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Chesapeake Bay Program
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Development of a Chronic
Sediment Toxicity Test
For Marine Benthic
Amphipods
Theodore H. DeWitt
Michele S. Redmond
John E. Sewall
Richard C. Swartz
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—jn Resource
• - .-.i'A JSJ07
U.S. EPA-ERLIN
Pacific Ecosystems Branch
2111 S.E. Marine Science Dr.
Newport, OR 97365-5260
Cooperative Agreement #CR-816299010
and
Contract #68-CO-0051
Project Officer
Robert C. Randall
Office of Research and Development
Pacific Ecosystems Branch
2111 S.E. Marine Science Dr.
Newport, OR 97365-5260
Printed by the U.S. Environmental Protection Agency for the Chesapeake Bay Program
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DISCLAIMER
This material has been funded in part by the U.S. Environmental Protection Agency under
Contract #68-CO-0051 and Cooperative Agreement #CR-816299010. It has been subjected to
the Agency's review, and it has been approved for publication as an EPA document. Mention
of trade names or commercial products does not constitute endorsement or recommendation for
use.
This report is Contribution No. N-240 from EPA's Environmental Research Laboratory-
Narragansett.
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TABLE OF CONTENTS
EXECUTIVE SUMMARY iii
INTRODUCTION x
ACKNOWLEDGEMENTS xiv
CHAPTER I: Collection, Handling, and Culture of the Amphipods
Leptocheirus plumulosus, Ampelisca abdita, Lepidactylus
dytiscus, and Monoculodes edwardsi
Introduction 1-1
Leptocheirus plumulosus 1-3
Ampelisca abdita 1-12
Lepidactylus dytiscus 1-23
Monoculodes edwardsi 1-27
Figures 1-33
CHAPTER II: The Acute and Chronic Sensitivity of the Estuarine
Benthic Amphipod, Leptocheirus plumulosus, to Chemically-
Contaminated Sediments
Introduction 2-01
Materials and Methods 2-03
Results 2-17
Discussion 2-25
Conclusions 2-40
Figures and Tables 2-43
CHAPTER III: Development of a Chronic Sediment Bioassay with
Ampelisca abdita
Introduction 3-01
Materials and Methods 3-02
Results and Discussion 3-09
Figures and Tables 3-23
APPENDIX A: Literature Review of Selected Chesapeake Bay
Amphipods A-l
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APPENDIX B: Procedures to Minimize the Risk of Releasing Non-
Indigenous Amphipods, Pathogens, Waters, or Sediments into
Local Waters or Watersheds B-l
APPENDIX C: Leptocheirus plumulosus Annex to the ASTM E1367-90
Document C-l
APPENDIX D: Research Methodology to Assess Chronic Toxicity of
Marine and Estuarine Sediments with the Benthic Amphipod,
Leptocheirus plumulosus D-l
APPENDIX E: Ampelisca abdita: Generic Life Cycle Test Design E-l
REFERENCES R-l
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Ill
DEVELOPMENT OF A CHRONIC SEDIMENT TOXICITY TEST
FOR MARINE BENTHIC AMPHIPODS
EXECUTIVE SUMMARY
Most marine sediment toxicity bioassays presently test only the acute mortality of
benthic organisms exposed for short periods of time to contaminated sediment. However, the
contaminant concentration needed to induce mortality may be considerably greater than the
concentration needed to slow somatic growth, or reproductive output. Benthic organisms in
the field are generally chronically (not acutely) exposed to contaminated sediments, and
benthic populations may be exposed to contaminants for more than one generation. Response
criteria are needed that reflect both the lethal and sublethal consequences of long-term
exposure to contaminated sediment.
Research to develop a chronic sediment test for marine benthic amphipods was
initiated in fall 1989 as a cooperative effort between researchers at the U.S. Environmental
Protection Agency and Oregon State University. A workplan for this research was developed
in conjunction with the EPA Chesapeake Bay Liaison Office, the EPA Office of Puget Sound,
the EPA Office of Science and Technology, and researchers from several laboratories in the
Chesapeake Bay and Pacific Northwest regions. The sequence of work proposed was to (1)
select several amphipod species that were abundant in Chesapeake Bay and showed promise
of being good candidates for use in toxicity tests based on previous research, geographic
distributions relative to urban or industrial centers, or taxonomic affinity with other
toxicologically sensitive amphipods; (2) collect these amphipods from Chesapeake Bay, ship
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them to the EPA Environmental Research Laboratory in Newport, OR, and attempt to culture
each species; (3) conduct short-term, comparative toxicity experiments to select the most
sensitive species; (4) select one or more species for further development based on ease of
culture and handling and toxicological sensitivity; (5) develop a chronic toxicity test method,
including appropriate controls; (6) conduct chronic, concentration-response, sediment toxicity
experiments with chemical-spiked sediment; and (7) conduct a chronic sediment toxicity test
with field-collected, chemically contaminated sediment from Chesapeake Bay.
We report here the results of this research effort which culminated in the development
of a research method for assessing the chronic toxicity of contaminated marine and estuarine
sediments using the benthic amphi pod, Leptocheirus plumulosus. The report is presented
in three chapters followed by five appendices. The first chapter describes our efforts at
collecting, handling, and culturing four estuarine amphipods from Chesapeake Bay, including
L_. plumulosus. This chapter includes maps of the distribution and abundance of these
amphipods within Chesapeake Bay and methodologies for establishing cultures of amphipods
which could be readily adopted by other laboratories. The second chapter reports the
development of acute and chronic sediment toxicity test methods for L. plumulosus, its
sensitivity to non-contaminant environmental variables, cadmium, two polynuclear aromatic
hydrocarbons, and contaminated sediment from Baltimore Harbor, MD. The third chapter
reports our attempts to develop a chronic sediment toxicity test with Ampelisca abdita. This
effort was not as successful as that with L. plumulosus, primarily because we could not
determine satisfactory conditions for its reproduction. The L. plumulosus and A. abdita
chronic sediment toxicity tests were developed independently, and, thus, different conditions
were necessary under which the experiments and final test protocols were conducted. The
different experimental conditions reflected the different ecologies of the two amphipods.
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Chapter 1: Collection, Shipping, Culture, and Handling of the Amphipods
Leptocheirus plumulosus, Ampelisca abdita, Lepidactylus dytiscus. and
Monoculodes edwardsi.
Leptocheirus plumulosus: This burrow-building aorid amphipod was found throughout
Chesapeake Bay in medium- to fine-grained sediments in waters of ca. 5-25%c. It was
tolerant of handling and shipping, and quite receptive to culturing. Cultures started in
March, 1990, were still thriving and expanding in August, 1992. Static-renewal cultures
were maintained in plastic dishpans with a <1 cm layer of sediment and 10-15 cm layer of
seawater at 20%o and 20°C. The water was replaced three times per week, at which time the
amphipods were fed with a mixture of cultured phytoplankton and a small amount of a dry
food mixture. Cultures were thinned when the density of adults exceeded ca. 1.5 cm"2, and
new cultures started with ca. 100 adults and 200 juveniles. Generation time for L.
plumulosus was approximately 4 wk and females produced multiple broods. High numbers
of sub-adult and newborn age-classes were available for sediment toxicity tests at all times
of the year.
Ampelisca abdita: This tube-building ampeliscid amphipod was found in relatively saline
waters (i.e., >20%c) of Chesapeake Bay adjacent to seagrass beds in sandy-mud sediments.
High densities of A. abdita were difficult to obtain in Chesapeake Bay, so animals from
Narragansett, RI, were used also. This species was more difficult to ship, handle, or culture
than L/. plumulosus. Culture conditions were similar to those for L. plumulosus, with the
exceptions that A. abdita were fed only algae (i.e., no dry food), the salinity was maintained
at 30%o, and the depth of the substrate was 4 cm. Our success in culturing A. abdita was
highly variable: some cultures thrived, but most had little reproduction. No environmental
factors could be identified that consistently regulated culture success.
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Lepidactylus dytiscus: Only one population of this haustoriid amphipod was found in
Chesapeake Bay during our efforts to collect the animal. It was tolerant of being shipped
across the country, and was maintained in laboratory culture from March, 1990, through
August, 1992 under conditions described for L. plumulosus, with the exception that the
substrate was fine sand and the salinity was 32%o. Reproduction occurred in spring and
summer, and was reduced or absent in fall and winter. Additionally, the generation time for
this species was approximately 1 yr. For these two reasons, cultures of L. dytiscus were not
sufficiently productive to supply the numbers of animals, on a year-round basis, that were
needed for sediment toxicity tests.
Monoculodes edwardsi: Populations of this oedicerotid amphipod were found in sandy
sediments at mesohaline salinities (i.e. 10-20%o) in Chesapeake Bay. Mortality during
shipping was high. M. edwardsi was cultured under conditions similar to .L. plumulosus,
with the exception that fine sand was used for the substrate, and small cultures have been
maintained for over 2 yr in the laboratory. However, generation time appears to be >1 mo.,
and only low and highly variable population densities (ca. 10-40 animals/pan) were sustained
under these conditions. Relative to L. plumulosus. it was not practical to attempt to produce
numbers of .M. edwardsi as were needed for the experiments.
Chapter 2: The Acute and Chronic Sensitivity of the Estuarine Benthic Amphipod,
Leptocheirus plumulosus, to Chemically-Contaminated Sediments
Two sediment toxicity test methodologies, one for acute exposures and the other for
chronic exposures, were designed using the benthic, estuarine amphipod, Leptocheirus
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plumulosus. Both methods used animals from laboratory cultures as were described in
Chapter 1. The 10-d acute sediment toxicity test utilized sub-adult sized animals (i.e., 2-4
mm) under static conditions, whereas the 28-d method used 0-d old newborn JL plumulosus
under static-renewal conditions. The procedures for both tests were very similar to sediment
toxicity test procedures established for other marine and estuarine amphipods (ASTM,
1990b). Procedures for reference toxicant controls and negative controls were also established
for both toxicity tests.
The mortality, growth, and fertility of F0 L. plumulosus were affected by 28-d
exposures to high concentrations of sediment-associated phenanthrene and field-collected
sediment from a highly contaminated site in Chesapeake Bay. Shorter-term exposures (i.e.,
10-d) of sub-adult L. plumulosus to sediment-associated acenaphthene, phenanthrene and the
polluted Chesapeake Bay sediment also affected mortality and growth; reproduction was not
recorded in the 10-d exposures since the test was designed to minimize the likelihood that
broods would be released during the exposure. The sensitivity of the 10-d and 28-d tests were
similar, particularly with respect to mortality and growth. Fertility, the number of juveniles
produced per female in an exposure chamber, was considerably more sensitive than mortality
or growth in one experiment, but not in a second experiment.
The acute and chronic K plumulosus sediment toxicity tests are sufficiently developed
to be used to assist in the evaluation of sediment quality, but the methodologies should be
viewed as interim in development until their limitations are better defined. This is
particularly true for the 28-d test method for which several uncertainties remain. Chief
among these is the interaction between nutrition and toxicological sensitivity, but also
requiring attention are the effects of salinity, temperature, and grain size on response
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sensitivity, its sensitivity to other chemicals, and its sensitivity relative to other toxicity tests.
Reference toxicity tests also need to be developed for the sub-lethal responses of growth and
fertility.
Chapter 3: Development of a Chronic Sediment Bioassay with Ampelisca abdita
Research with Ampelisca abdita sought to develop culture methods and a chronic
bioassay for this species. Bioassay development built on the research of Scott and Redmond
(1989), who showed that A. abdita could be used to test chronic and population endpoints.
Culturing methods and results are described in Chapter I. The approach to chronic test
development was to 1) establish cultures, 2) estimate optimum temperature and salinity
regimes, 3) outline a proposed chronic test design, 4) evaluate the chronic test design with
uncontaminated sediment, and 5) evaluate the chronic test design with contaminated
sediment. The experiments conducted addressed points 2-4. Both cultures and the controlled
experiments described in this section utilized amphipods from Narragansett, RI.
A workable draft test protocol for a generic, 35-day chronic sediment toxicity test with
this species was developed. However, successful reproduction in laboratory-held A. abdita
was inconsistent. Juvenile amphipods of a known age were successfully isolated from
brooding females held in seawater only. Although it was feasible to initiate a test with
newly-released juveniles, 8-10 day old amphipods were easier to work with. Sex ratio of
juvenile amphipods used to start a test was determined from daily and final observations.
A survival curve for an acceptable test control could be distinguished from that showing
unacceptable control mortality. Significant differences in growth were also detected under
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control conditions in 10-d to 14-d exposures for this species, and a short-term growth test may
be a viable sublethal toxicity test for A. abdita.
There are still unresolved problems with the culture and chronic testing of this
species. In controlled experiments with uncontaminated sediment, the amphipods grew,
looked healthy, and produced eggs and sperm, but rarely reproduced. Replicate culture
containers with the same density, light cycle, salinity, temperature, sediment, and renewal
and feeding regimes performed drastically differently, regardless of container type. Shipping
and handling stress may be particularly important in determining the success of subsequent
toxicity test responses. Offspring of field-collected and shipped females with broods showed
poorer survival after 10-d than did offspring from cultured females. Juveniles developing in
the maternal brood pouch may be a very sensitive life stage for A. abdita. This species may
require a flow-through system with frequent volume replacements, a different photoperiod
and temperature regime, or may not be culturable in some waters.
To complete the development of a sublethal sediment tests with this species, the low
reproduction problem must be resolved, successful life cycle tests conducted in
uncontaminated sediment to firmly establish performance under control conditions, and
finally chronic and short-term growth tests conducted with contaminated material.
Intel-laboratory comparisons of the test methodology will be vital to ensure that this chronic
test can be conducted successfully in other regions of the country.
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INTRODUCTION
Nearly all chemical contaminants entering coastal waters eventually accumulate in
the sediments. Most toxic contaminants (including heavy metals, chlorinated pesticides,
PAH's, PCB's, TBT, oil and grease) bind to particulate matter in the water column and sink
to the sediment surface. Thus, concentrations of toxic chemicals in sediments can be several
orders of magnitude higher than in the water column. As many of these chemical
contaminants are persistent and can exert toxic effects to both benthic and demersal biota
for years after their initial discharge, sediments have become both a sink and a source of
contamination in many marine ecosystems.
The impact of sediment pollution on marine ecosystems is reflected in changes in
macrobenthic community structure and function (Pearson and Rosenberg, 1978; Swartz et al,
1985b, 1986a; and others). Benthic mfauna are good indicators of sediment contamination
because of their proximity and long-term exposure (as residents of sediments) to toxic
materials in polluted sediments. Benthic community responses to organic enrichment are
predictable (Pearson and Rosenberg, 1978; Mearns and Word, 1982), but comparable
predictive models of the response of the benthos to chemical contaminants have not been
developed. Major obstacles to the development of such models are (1) our inability to
discriminate between organic enrichme;nt and contaminant effects and (2) uncertainty in the
long-term responses of benthic fauna to chemical contaminants.
Little is known of the toxicological responses of most marine benthic taxa to
contaminated sediment. Only a few marine species have been examined, and most data
concern only acute mortality (Swartz, 1987). Nonetheless, the acute mortality of certain
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benthic species is being used as an assessment of the environmental impact of contaminated
sediment (Chapman and Long, 1983). Surveys of the relative toxicological sensitivities of
taxonomically diverse benthic species are necessary to validate the current choices of
sediment toxicity test species. Furthermore, virtually nothing is known of the long-term
effects of contaminated sediments on benthic populations, such as effects on individual
growth rates, reproductive output, or rate of population growth. New sediment toxicity tests
are needed to predict the long-term and sublethal impacts of chronic exposure of benthic
invertebrates to low levels of sediment contamination. These issues must be addressed if the
effects of contaminated sediments on marine ecosystems are to be assessed and protective
sediment quality criteria developed.
Most marine sediment toxicity bioassays presently test only the acute mortality of
benthic organisms exposed for short periods of time to contaminated sediment. However, the
contaminant concentration needed to induce mortality may be considerably greater than the
concentration needed to slow somatic growth, or reproductive output. Benthic organisms in
the field are generally chronically (not acutely) exposed to contaminated sediments, and
benthic populations may be exposed to contaminants for more than one generation. Response
criteria are needed that reflect both the lethal and sublethal consequences of long-term
exposure to contaminated sediment.
Research to develop a chronic sediment test for marine benthic amphipods was
initiated in fall 1989 as a cooperative effort between researchers at the U.S. Environmental
Protection Agency and Oregon State University. This project was funded in part by the EPA
Chesapeake Bay Liaison Office, EPA Office of Puget Sound, and EPA Office of Science and
Technology with the understanding that the new sediment toxicity test would be directly
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applicable to sediments from Atlantic coast and Pacific Northwest estuaries. A workplan for
this research was developed in conjunction with these EPA offices and researchers from
several laboratories in the Chesapeake Bay and Pacific Northwest regions. The sequence of
work proposed was to (1) select several amphipod species that were abundant in mid-Atlantic
estuaries, particularly Chesapeake Bay, and showed promise of being good candidates for use
in toxicity tests based on previous research, geographic distributions relative to urban or
industrial centers, or taxonomic affinity with other toxicologically sensitive amphipods; (2)
collect these amphipods from Chesapeake Bay, ship them to the EPA Environmental
Research Laboratory in Newport, OR, and attempt to culture each species; (3) conduct short-
term, comparative toxicity experiments to select the most sensitive species; (4) select one or
more species for further development based on ease of culture and handling and toxicological
sensitivity; (5) develop a chronic toxicity test method, including appropriate controls; (6)
conduct chronic, concentration-response, sediment toxicity experiments with chemical-spiked
sediment; and (7) conduct a chronic sediment toxicity test with field-collected, chemically
contaminated sediment from Chesapeake Bay.
We report here the results of this research effort which culminated in the development
of a research method for assessing the chronic toxicity of contaminated marine and estuarine
sediments using the benthic amphipod, Leptocheirus plumulosus. The report is presented
in three chapters followed by five appendices. The first chapter describes our efforts at
collecting, handling, and culturingfour estuarine amphipods from Chesapeake Bay, including
L_. plumulosus. This chapter includes maps of the distribution and abundance of these
amphipods within Chesapeake Bay and methodologies for establishing cultures of amphipods
which could be readily adopted by other laboratories. The second chapter reports the
development of acute and chronic sediment toxicity test methods for L,. plumulosus, its
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sensitivity to non-contaminant environmental variables, cadmium, two polynuclear aromatic
hydrocarbons, and contaminated sediment from Baltimore Harbor, MD. We believe these
methods will find wide utility in Chesapeake Bay and throughout much of the country. The
third chapter reports our attempts to develop a chronic sediment toxicity test with Ampelisca
abdita. This effort was not as successful as that with L. plumulosus, primarily because we
could not determine satisfactory conditions for its reproduction. The appendices include (A)
a literature review of the biology and ecology of the amphipods initially considered for the
sediment toxicity test development; (B) a protocol for handling and disposing materials that
come into contact with non-indigenous amphipods, sediments or waters; (C) a methodology
for testing the acute toxicity of contaminated sediment with L. plumulosus; (D) a methodology
for conducting chronic sediment toxicity tests with L/. plumulosus; and (E) the design of a life
cycle sediment toxicity test with A. abdita.
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ACKNOWLEDGEMENTS
Many aspects of this project could not have been completed without the help of several
people. We thank Ray Alden, Rich Batiuk, Faith Cole, Janet Lamberson, Beth McGee, Chris
Schlekat and John Scott for helping to improve this manuscript through their thoughtful
reviews. We also thank Richard Batiuk, Jack Gakstatter, Chris Zarba, Barry Burgan, Ray
Alden, Bob Diaz, Tuck Hines, Fred Holland, Rom Lipcius, Harriet Phelps, Eli Reinharz, and
John Scott for guidance in developing the research plan and for providing logistical support
during the collection of amphipods from Chesapeake Bay and Narragansett Bay. We thank
John Brezina, Emily Deaver, Paul Gerdes, Tammy Tonare, and Tom White for their
assistance in the field. We thank Beth McGee and Chris Schlekat for collecting sediment and
preparing the sediment dilution series and Claudia Walters for providing QA/QC assistance
in the experiment with field sediment. We thank Dave Hansen for providing AVS and SEM
analyses. We also thank Mary Culver, Linda Liptrap, and Sharon Nieukirk for assistance
in preparation of species distribution maps. And finally, we wish to thank our US EPA and
AScI colleagues in Newport, OR, for their tireless assistance in the culturing, toxicology, and
chemistry laboratories: Michael Becerra, Wally DeBen, George Ditsworth, Steve Ferraro,
John Frazier, Laura Hoselton, Jill Jones, Janet Lamberson, Bob Ozretich, Don Schults, and
Rob Singleton. This project was supported in part by US EPA cooperative agreement
CR816299010 to Oregon State University. Funds for this research were provided, in part,
by the EPA Chesapeake Bay Liaison Office, the EPA Region 10 Office of Puget Sound, and
the EPA Office of Science and Technology.
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Chapter I
COLLECTION, HANDLING, CULTURE OF THE AMPHIPODS
LEPTOCHEIRUS PLUMULOSUS, AMPELISCA ABDITA,
LEPIDACTYLUS DYTISCUS, AND MONOCULODES EDWARDSI
1.1 INTRODUCTION
The development of acute and chronic sediment toxicity tests with Chesapeake Bay
amphipods began with the identification and collection of candidate species and quickly
followed with the culture of these species. Six species were selected for consideration for
sediment toxicity test development from a list of 60 amphipod species (Appendix A) based on
their distribution relative to sediment contamination, taxonomic and ecological similarity to
other amphipods currently used in sediment toxicity tests, and their relative abundance and
ecological importance in Chesapeake Bay. Of these, five species were collected from
Chesapeake Bay sediments in March, 1989, and shipped west to the EPA ORD ERL-N
Newport laboratory for development as sediment toxicity test species. Culture techniques
needed to be developed for each species so that sufficient numbers of animals could be
available for experiments. Methods for field collection, handling and shipping, and culturing
four species are presented below.
These five amphipod species (i.e., Leptocheirus plumulosus, Ampelisca abdita,
Lepidactylus dytiscus, Monoculodes edwardsi, and Corophium lacustre) were collected from
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various localities between Baltimore, MD, and Virginia Beach, VA, by T.H. DeWitt and
assistants during a trip that extended from 19 to 28 March, 1990. Shortly after collection,
each species was sorted, held in native habitat sediment in running (or frequently changed)
seawater, placed in shipping containers and shipped via Federal Express (overnight delivery)
to the EPA laboratory in Newport, OR. A sediment sample from each collecting site was also
sent to Newport for grain size analysis. One of the original candidate species (Neohaustorius
schmitzi) was not collected for logistical reasons (i.e., uncertainty of collecting locations), and
an alternate species, C_. lacustre, was substituted. However, JC. lacustre failed in culture,
subsequent attempts to have it collected and shipped to Oregon failed, and it was deleted
from the remainder of the research effort. Ampelisca abdita was also collected from
Narragansett, RI, by personnel from the EPA Environmental Research Laboratory.
Leptocheirus plumulosus showed the best promise for mass culturing, although
populations of Ampelisca abdita, Monoculodes edwardsi and Lepidactylus dytiscus have been
sustained for nearly two years in the laboratory. Static-renewal culture conditions were
employed for all species because of previous success at culturing amphipods in this manner
(DeWitt, 1987) and in order to minimize the amount of waste-water which had to be
sterilized. Because these species were not indigenous to Oregon estuaries, special efforts
were made to chlorine-bleach-sterilize or autoclave all materials (e.g., sediment, water,
glassware, adsorbent materials, culturing and handling equipment, etc.) that came into
contact with the amphipods before otherwise used or discarded. These materials were
sterilized to minimize the risk of accidentally introducing non-indigenous amphipods or
pathogens into local waters. Procedures for handling and sterilization of these material are
presented in Appendix B. We strongly believe that these quarantine handling and culturing
practices must be adopted by all laboratories using any non-indigenous toxicity test organism
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or sediment. Additionally, some States require permits and special containment procedures
for handling and culturing non-indigenous species; the procedures described below may or
may not be sufficient for another laboratory's permit to work with such materials.
1.2 Leptocheirus plumulosus
1.2.1 SUMMARY
Leptocheirus plumulosus was collected in large numbers from the field and was easily mass-
cultured in the laboratory. This species was very tolerant of being handled and survived
shipping well. Cultures were reared under static renewal cultures using equipment
commonly available in aquatic laboratories. Further research into the nutritional
requirements of this species and simplification of handling methods should further reduce the
cost and effort involved in culturing these amphipods, and may make culturing preferable to
field collections even in areas with large natural populations.
1.2.2 OVERVIEW OF THE SPECIES
Leptocheirus plumulosus is a euryhaline amphipod of the family Aoridae found
throughout the mesohaline portions of Chesapeake Bay (Fig. 1-1). Under laboratory
conditions, it is fast growing and can mature in less than 25 days at 25°C. The size of the
first brood is typically 10-20 young for a healthy female; larger mature females can produce
over 40 young in a single brood (see Chapter 2). At 25°C, the minimum interval of time
between broods is less than 10 days. Females may live for over 100 days and produce at
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least 6 broods in that period, and potentially may live longer and produce more broods.
Reproduction of animals in the cultures continues uninterrupted year-round under constant
culture conditions. The sex ratio of the young is assumed to be approximately 1.0
(females/males). The newly released young are about 1.5 mm long, and, at 25°C, they can
double in size in about 10 days and triple their size (or more) in 14 days. Large adults are
over 1 cm long. Eggs first appear in the ovaries of females by age 12-d (i.e., days since
leaving the maternal brood pouch), and eggs may be seen in the brood pouch by age 14-d.
L^ plumulosus constructs a U-shaped burrow in the soft, organically rich sediment that
it seems to prefer. The burrow walls have little cohesive structure to them, and the burrows
disintegrate during sieving. The animal pumps water through the burrow and may filter out
suspended particles for food. It also pulls in sedimentary particles surrounding the tube
opening, apparently scraping the surface of mineral particles for food or tearing pieces of
organic material into small enough pieces to ingest. Animals will occasionally leave their
tubes to roam the sediment surface, apparently picking up pieces of food material or
searching for mates. This feeding mode may allow individuals to live in water without
sediment for extended periods of time if particulate food is available. The regular spacing
of burrow openings suggests that this species may be territorial. Males may compete for
mates as evidenced by their high mortality, due to fighting, when held in culture containers
in the absence of females. Survival of both sexes may be >90% in mixed sex culture between
0-4 wk of age.
Please refer to Appendix A (Literature Review of Selected Chesapeake Bay
Amphipods) for further details and references concerning the natural history of Leptocheirus
plumulosus.
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1.2.3 FIELD COLLECTION
Leptocheirus plumulosus were collected in shallow water (1-2 m deep) by hand using
a small grab sampler and a suction dredge; the latter was far more efficient at collecting this
amphipod. Approximately 50 amphipods were collected in a 2 h period by T.H. DeWitt on
3/21/90 from muddy sediment at the end of the fishing pier at Fort Armitage Park in
Baltimore Harbor, MD, with a small Ponar sampler in 1-2 m depth (salinity ca 5%o). This
was not a good site for collecting L. plumulosus. Approximately 1000 animals were collected
rapidly (i.e., < 0.5 h) by T.H. DeWitt and Tom White (Virginia Institute of Marine Science)
on 3/24/90 from muddy substrate in Queens Creek, York River, near Williamsburg, VA, in
shallow water (ca. 1 m deep) by suction dredge (salinity ca 14%c); this was a very good site
to collect L_. plumulosus in March, 1990. Scientists at the Maryland Department of
Environment have routinely collected this species from Corsica River and Magothy River
subestuaries in northeastern Chesapeake Bay (B. McGee and C. Schlekat, personal
communication). However, the abundance of this amphipod can be variable at any site
(including Queens Cr. and Corsica R.), ranging from highly abundant to absent (Emily
Deaver and Ray Alden, Old Dominion University, pers. comm.).
Leptocheirus plumulosus is widely distributed in Chesapeake Bay (Fig. 1-1), and is
therefore potentially widely available year-round. However, this amphipod is highly motile
especially nocturnally, at which time many individuals may be found swimming in the nekton
(Dauer et al, 1982). Furthermore, L. plumulosus populations boom in the early spring and
bust in the summer following the return of predatory fish to Chesapeake Bay (Hines et al,
1986). Thus, population densities at specific collection sites will probably vary considerably
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throughout the year, and some sites will be ephemeral in quality while others (such as
Corsica R., apparently) may be quite dependable as sources of this amphipod.
1.2.4 SHIPPING
Leptocheirus plumulosus were shipped successfully from Chesapeake Bay to Newport,
OR, in March, 1990, and from Newport, OR, to Narragansett, RI, in June, 1991. Field-
collected animals were held overnight or longer in running or frequently changed bay water
at a salinity and temperature close to that where the animals were collected. Dead or injured
animals were removed prior to packing. The choice of salinity depended on the ambient
salinity from which the animals were taken: ~15%c from the field, and 20%e from the
laboratory cultures. Amphipods were shipped at densities of 50-100 per plastic container (i.e.,
a 250-1000 ml sandwich box or ice cream tub) containing water and substrate (i.e., 0.2-1.0
cm layer of marsh grass detritus) or just water (i.e., 15-25%o). Field-collected animals were
shipped with substrate and cultured amphipods without substrate.
Several plastic containers were placed in an insulated cooler along with 3 or 4 freezer
packs (such as blue ice) to keep the temperature cool, but above freezing, and then the cooler
was sealed and immediately shipped by overnight delivery. The field-collected L. plumulosus
suffered approximately 25% mortality, whereas very few of the laboratory-cultured animals
died during shipment. Mortality may have been due to lack of oxygen caused by the BOD
of the higher organic-content substrate included with the field-collected amphipods. Future
shipments of L/. plumulosus should (1) minimize the amount of substrate included in the
shipping containers, (2) use low organic-content or sterile substrates, or (3) omit substrate
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altogether. Shipping success might be further enhanced if the amphipods were shipped in
oxygen-saturated water.
Upon arrival at the laboratory, the unopened containers of amphipods were placed in
a water bath and slowly acclimated to 20°C. The containers were then opened, the overlying
water was decanted for chlorination and disposal, and the containers were refilled with 20°C
water at the same salinity as the packing water. Later, the amphipods were sieved from the
packing substrate and transferred to tubs for culturing.
1.2.5 CULTURING
Leptocheirus plumulosus was cultured in inexpensive polyethylene tubs measuring
29.2 cm x 34.3 cm x 13.3 cm (depth) (i.e., 11.5" x 13.5" x 5.5"), holding about 13 L (3.5 gal)
of seawater (11-12 cm deep) with a <1 cm thick sediment layer. This configuration held
several hundred mature animals and facilitated the handling of individual culture containers
for sieving, water replenishment and moving. The tubs were held in shallow seawater-table
trays which served the dual purposes of catch basins for any water spilled from the tubs and
water baths to maintain the appropriate culture temperature. The amphipods were also
cultured in the tubs placed on shelves with the room air temperature maintained at 20°C.
The cultures were maintained at 20°C and a salinity of 20%o. The culture sediment was
a muddy sand from South Beach, Yaquina Bay, OR, that had been sieved through a 0.5 mm
or 0.25 mm mesh sieve. Photoperiod was maintained at 16:8 hr light:dark. The tubs were
gently aerated constantly. About 60% of the water in each tub was changed every other day,
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except on weekends. This was accomplished by pouring off the old water and refilling the
tub from a plastic pitcher with fresh seawater laden with the algal food. The stream of
incoming water was directed onto a ilat piece of glass held at the water surface to disburse
the flow and minimize disturbance of the sediment bed. The renewal water consisted of
seawater (ca. 32%o), cultured phytoplankton and deionized water which were combined to a
salinity of 20%e and ca. 106 algal cells per ml. The algae used are Pseudoisochrysis paradoxa
and Phaeodactylum tricornutum in equal portions by volume. The cultures were also fed
about 0.5 gr. of a dry food (i.e., "gorp') just after the water change. Gorp consisted of 48.5%
Tetra min®, 24% dried alfalfa, 24% dried wheat leaves and 4.5% Neo-Novum® (a maturation
feed for shrimp mariculture; Argent Chemical Laboratories, Redmond, WA), combined and
ground to a fine powder. The gorp was sprinkled on the water surface.
An important aspect of maintaining culture health was to prevent overcrowding.
Densities should be maintained below 1500 per tub (e.g., <1.5 cm"2). The occurrence of
overcrowding was marked by cultures with a large number of animals of small size and few
gravid females. Those females that were gravid bore only a very small number of eggs (e.g.
<5). Under these conditions, newly released young were very difficult to obtain, and the
animals in the tubs appeared to be stressed from food or space limitation. The number of
adults (i.e., animals >4 mm long) should not exceed ca. 400 per tub. To avoid overcrowding,
cultures were thinned approximately every two months by sieving through a 1 mm mesh
sieve. This allowed the young to pass through and remain in the sediment. Only about 100
healthy adults were returned to the culture tub. The rest were used to start new cultures
or disinfected and discarded.
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L. plumulosus has been in culture in the Newport, OR, EPA laboratory since March, 1990.
The performance of the cultures and the potential of this species for mass production of
animals are excellent. Twenty culture tubs are currently maintained at 20°C which has
consistently provided sufficient numbers of animals to conduct 50- to 80-beaker acute
sediment toxicity experiments. These cultures provided up to 1000 newly released young per
month for chronic sediment toxicity tests. Densities of >1500 animals per tub (i.e., >1.5 cm'2)
were readily achieved in the culture tubs, but this was an overcrowded density leading to
reduced growth and fecundity as described previously. L. plumulosus does not seem sensitive
to seasonal changes when maintained under constant culture conditions, and can provide
animals for toxicity tests year-round.
1.2.6 HANDLING
The contents of culture tubs were gently sieved through a 0.5 mm mesh to obtain
subadult L_. plumulosus (i.e., 2-4 mm long) for acute toxicity tests. This allowed some of the
very smallest animals to pass through, but retained all of the animals over a few days old.
Larger animals were excluded by gently sieving the animals through a 1 mm screen.
Animals were rinsed free of sediment and washed into a shallow glass picking dish.
Subadults 2-5 mm long were transferred by pipette into a smaller glass dish for acclimation
to the test temperature and salinity. Water for sieving and rinsing were the same
temperature and salinity as the cultures (e.g., 20%c) to minimize stress. Unused cultured
animals were returned to the culture tub after the sediment bed had settled for a few hours.
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Cultured amphipods used in the 28-d chronic sediment toxicity test were of uniform
age, 1- to 2-d post brood-release. To obtain newly-released juveniles, contents of the culture
bins were sieved through a 1 mm screen to isolate adults, and gravid females were selected
from the mass of adults and transferred to fresh culture tubs with sediment so that they
could acclimate to the temperature and salinity of the toxicity test. The females were fed in
the same manner as the general cultures during this acclimation period. Gravid females
were isolated for 8 days before the start of the toxicity test if the test was run at 20°C, or for
5 days in advance if the test was conducted at 25°C. Gravid females were easily recognized
by the dark egg mass in the brood pouch; as the eggs approach hatch, the mass turned a tan
color and became somewhat translucent which was more difficult to see without a dissecting
microscope. Three days before the start of a toxicity test, the females were sieved from the
isolation tub using a 1 mm mesh, rinsed well and placed in a glass dish with water only (e.g.,
no sediment) at the test temperature and salinity and fed with the algal suspension used to
feed the general cultures. The females were inspected to insure that no previously released
young were transferred to the glass dish. All debris was removed from the bottom of the dish
since this could conceal young. The following day, the adult females were separated from the
young they had released using a 1 mm screen, and the young left behind were ready for
experimental use. This process was repeated on the succeeding two days in order to provide
additional animals in the event that insufficient numbers of young were produced on the day
of the toxicity test. The isolated juveniles were maintained in a glass dish with sediment pre-
sieved to <0.25mm and were fed with the algal suspension. Exposures were initiated only
with the <24hr-old juveniles. If insufficient numbers of these young were available, some
replicates of the experiment were set aside to be started the following day with newly isolated
juveniles. This procedure was judged to be superior to mixing together juveniles produced
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on different days on the basis that variability in sensitivity or growth within replicates would
be reduced if all test animals were of the same age.
The performance control (e.g., culture sediment) survival of these young was >90%.
Although each female can produce up to 40 young, we isolated one gravid female for each
newborn that is needed for a toxicity test. This seeming excess was required because of the
uncertainty of the timing of brood release for each female. After producing young, the
females were returned to the cultures; the processes of culture thinning and isolating newly
released young were often combined into a single effort.
1.2.7 CONCLUSION
Leptocheirus plumulosus was well adapted for laboratory culture. Cultures consistently
produced large numbers of animals within narrow age brackets required for research or
routine acute or chronic toxicity tests. Field-collected animals may have also been used for
acute and chronic sediment toxicity tests, although the utility of these approaches was not
evaluated in this study. Further work in improving culture techniques should be directed
at defining the minimum diet required to maintain highly productive cultures, obtaining
greater synchrony in brood release among gravid females, and simplifying the process of
obtaining newly released young needed for chronic toxicity tests.
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1.3 Ampelisca abdita
1.3.1 SUMMARY
This species has been maintained in culture for several generations, but with great
variability in culturing success. Periodic reference toxicant tests with cadmium chloride
showed that the sensitivity of cultured animals was comparable to that of field-collected
animals from the source population in Narragansett, RI. Several hundred animals were
maintained at 20°C in each of several plastic dish bins, with about 4 cm of Yaquina Bay,
Oregon, sediment and about 13 cm of overlying seawater. Half of the overlying seawater was
renewed 3-5 times per week with a mixture of seawater (30%c) and algae. Amphipods were
sieved from the culture bins when needed for testing, or when density exceeded 2000 animals
per bin. With more research, this species may hold the potential to be routinely cultured.
However, the productivity of these cultures was too variable to consistently provide the large
numbers of animals needed for frequent toxicity tests.
1.3.2 OVERVIEW OF THE SPECIES
Ampelisca abdita is a tube-dwelling amphipod belonging to the family Ampeliscidae,
found mainly in protected areas from the low intertidal zone to depths of 60 m. It ranges
from central Maine to south-central Florida and the eastern Gulf of Mexico (Mills 1964,
Bousfield 1973), and has also been introduced into San Francisco Bay, CA (Nichols and
Thompson 1985). In Chesapeake Bay, A. abdita has been reported from moderate- to high-
salinity waters (e.g., >20%o) (Fig. 1-2). Where A. abdita are present, they are often dominant
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members of the benthic community, with densities up to 110,000 m"2 (Nichols and Thompson
1985, Stickney and Stringer 1957, Santos and Simon 1980). This species generally inhabits
sediments from fine sand to mud and silt without shell, although it may also be found in
relatively coarser sediments with a high organic content (Stickney and Stringer 1957).
Ampelisca abdita is a particle feeder, feeding both on particles in suspension and on
those from the surface of the sediment surrounding its tube. Gut contents of field-collected
specimens have been found to include algal material, sediment grains, and organic detritus
(Mills 1967, Stickney and Stringer 1957).
In the colder waters of its range, A. abdita produces two generations per year, an
overwintering generation which breeds in the spring and a second which reproduces in mid
to late summer (Mills 1967, Nichols and Thompson 1985). Each female produces one brood,
and males die shortly after mating. Sex ratio of the population at breeding times is
approximately 1:1. In New England, breeding of the overwintering generation begins when
the water temperature is about 8°C, but in warmer waters south of Cape Hatteras, NC,
breeding might be continuous throughout the year. Adults mate in the water column, and
intense breeding activity is correlated with the full moon and spring tides. Juveniles are
released after approximately two weeks in the brood pouch, at about 1.5 mm in length. It
then takes 40-80 days for newly released juveniles to become breeding adults (Mills 1967).
Females in a population from Barnstable Harbor, MA, were found to carry a mean of 26 eggs
(Mills 1967), and a population from North Carolina a mean of 13.7 (Nelson 1980). In the
laboratory, A. abdita will breed all year, although large numbers of individuals are needed
to ensure the ability to harvest sufficient numbers for testing. At 20°C, its full life cycle is
approximately 6 to 8 weeks (Scott and Redmond 1989).
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A. abdita has been collected in waters of -2°C to 27°C (Redmond and Scott,
unpublished data). It is euryhaline, and has been reported in waters which range from fully
marine to 10%c salinity (Bousfield 1973). This species is photonegative, and has been found
to have a strong mortality response when exposed to sunlight (Redmond and Scott,
unpublished data).
Please refer to Appendix A (Literature Review of Selected Chesapeake Bay
Amphipods) for further details and references concerning the natural history of A. abdita.
1.3.3 FIELD COLLECTION
Ampelisca abdita were collected by hand (e.g., shovel and sieve) and with a suction
dredge from intertidal and shallow subtidal sediments, respectively. Personnel from the EPA
Environmental Research Laboratory and SAIC in Narragansett, RI, collected several hundred
amphipods by shovel and sieve from the intertidal in Pettaquamscutt Cove (Pettaquamscutt
River, Narragansett, RI) on 1/30/90, 8/29/90, 3/26/91, and 7/31/91. They regularly collect A.
abdita in this manner from this site, although winter collections sometimes require chopping
through ice to access the sediment. T.H. DeWitt (with help from Paul Gerdes, Virginia
Institute of Marine Science) collected approximately 150 A. abdita on 3/24/90 with a suction
dredge from muddy sand substrate in Zostera beds off the lee side of Allen's Island, VA, near
the mouth of York River (salinity ca 25%c). This method allowed rapid collection of many
amphipods over a large area of the benthos. However, many other amphipod species were
simultaneously collected requiring considerable post-collection sorting. Subtidal populations
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(3-10 m) were collected from several sites in San Francisco Bay on 7/30-31/91 with a small
grab sampler and bucket dredge from a boat (John Brezina, personal communication).
A. abdita is widely distributed along the U.S. eastern seaboard (including the lower
reaches of Chesapeake Bay, Fig. 1-2) and San Francisco Bay, CA, and is therefore potentially
widely available for field collection. Intertidal populations may be seasonally ephemeral, with
mass emigrations occurring in response to disturbance from other macrofauna or storms
(Mills, 1967; Grant, 1965). However, the intertidal and shallow subtidal population at
Pettaquamscutt R. (Narragansett, RI) has proven to be a dependable, year-round source of
A. abdita for several years (M.S. Redmond, unpubl. data).
1.3.4 SHIPPING
Ampelisca abdita have been shipped around the country in several different ways,
each with mixed success. Field-collected animals were held overnight or longer in running
or frequently changed bay water at a salinity and temperature close to that where the
animals were collected. Dead or injured animals were removed prior to packing. Amphipods
were shipped from Chesapeake Bay in March, 1990, packed 25-50 individuals per water-filled
(20%o) plastic container (i.e., 250-1000 ml sandwich boxes or ice cream tubs) with a 3-5 mm
thick, silty-sand substrate. Amphipods were shipped from Narragansett, RI, in (1) sandwich
boxes or cubitainers with A. abdita in seawater only, (2) sandwich boxes with a 2 cm layer
of mud, (3) sandwich boxes filled with mud and >1 cm layer of seawater, or (4) 4-1 jars with
a 2 cm layer of mud. The various containers were topped-off with water or had a 1 cm
headspace of air. Several plastic containers were typically placed in an insulated cooler along
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with 3 or 4 freezer packs (such as blue ice) to keep the temperature cool, but well above
freezing, and the cooler was sealed and immediately shipped by overnight delivery. A. abdita
in their tubes only were shipped from San Francisco Bay in large plastic bags filled to 1/2
capacity with water, with a 5-6" air headspace. Each bag was packed in an insulated
cardboard box with a single ice pack for cooling.
Survival of shipped A. abdita was variable. Inclusion of ice packs was very important:
most of the amphipods died in one shipment that omitted cooling. Shipping success might
be further enhanced if the amphipods were shipped in oxygen-saturated water.
Upon arrival at the laboratory, the containers of amphipods were opened, placed in
a water bath with aeration, and slowly acclimated to 20°C. The overlying water in the
containers was decanted for chlorination and disposal, and the containers were refilled with
20°C water at the same salinity as the packing water. Later, the amphipods were sieved
from the packing substrate and transferred to tubs for culturing.
1.3.5 CULTURING
Cultures of Ampelisca abdita were initially maintained in 1-gal glass jars which held
ca. 4 cm sediment and ca. 20 cm overlying water. The water was constantly gently aerated
from a glass pipette attached to a filtered air supply. More recently, we have maintained
cultures in plastic dish tubs (ca. 27cm x 30cm x 17cm deep) with a 4 cm layer of sediment
and 13 cm column (ca. 10.5 L) of overlying water. The water is also constantly aerated. The
tubs are held in shallow seawater-table trays which serve the dual purposes of catch basins
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for any water spilled from the tubs and water baths to maintain the appropriate culture
temperature.
The culture sediment was collected from tide flats in Yaquina Bay, OR, and wet-sieved
through a 250 um sieve before use. The fine sediment facilitated recovery of small juvenile
animals when cultures were sieved through a 0.25 mm screen. The photoperiod was
maintained at 16 h light: 8 h dark. This mid-summer photoperiod has been found to sustain
reproductive activity in other amphipod species (Arthur, 1980).
The culture water was renewed 3-5 times per week, at which time the cultures were
also fed. Prior to renewal, aeration was stopped, the sides of the tubs rinsed down, and any
amphipods trapped on the water surface pushed underwater with a glass rod. The floaters
were given some minutes to burrow into the sediment. Then, ca. 1/3 to 1/2 of the overlying
seawater in each bin was carefully poured off and replaced with an algae-seawater mixture.
The renewal water was added using a turbulence reducer (glass dish attached to a glass rod)
held just at the water surface, so that the sediment was not disturbed. The renewal water
was a mixture of filtered seawater (28-35%c), a culture of the diatom, Phaeodactylum
tricornutum, and a culture of the golden-brown flagellate, Pseudoisochrysis paradoxa, in the
ratio of approximately 1.5:1:1. The salinity of the mixture was adjusted to ca. 30%o with
deionized water if necessary. About 3.5 liters of the renewal water were added to each
amphipod culture bin.
Cultures were maintained at 20°C and 28-35%o which was the routine sediment
toxicity test temperature and salinity for A. abdita (Scott and Redmond 1989). A. abdita will
tolerate lower temperatures, but it grows more slowly and will probably not reproduce until
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the temperature reaches about 10°C. We tried to culture this species at 25°C, hoping this
would shorten the life-cycle and increase productivity. This is a temperature that shallow-
water populations would encounter in summer. However, after a few generations, culture
production declined. Similarly, while Bousfield (1973) reported A. abdita present at 10%o, we
had some data that suggested that culture productivity was worse at 20%o than at 30%o. We
do not know if the higher temperature or lower salinity was the cause of poor culture
performance since there was considerable variation among replicates. Our decision to use
20°C and 28-35%o as routine culture conditions was based on the results of a preliminary
experiment and our judgement that the higher temperature and lower salinity might
represent marginal environmental conditions for populations in the field.
From January - August, 1990. most A. abdita were maintained at 25°C in glass gallon
jars with screened overflows and fed only Pseudoisochrysis paradoxa, mixed with seawater,
3 times per week. These cultures were productive initially, but reproduction decreased by
the fourth generation. Temperature was then decreased to 20°C, and the amphipods were
fed a mixture of three algal species (e.g., P. paradoxa. Phaeodactylum tricornutum. and
Chaetoceros calcitrans) 5 times per v/eek. Experimental data (see Chapter 3) indicated that
A. abdita grew better when fed a mixture of algal species than if they were fed P_. paradoxa
alone. Samples taken from culture jars in October 1990 (after ca. 6 generations), indicated
that reasonable numbers of individuals were again being produced. However, the cultures
crashed in late November, possibly due to a late-fall water quality problem (e.g., natural
release of toxic compounds during the decomposition of dying macrophytes or a bloom of
dinoflagellates in Yaquina Bay), and the cultures had to be replenished with animals shipped
from Narragansett, RI.
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Since the January, 1991, shipment of anaphipods, A., abdita have been cultured in
plastic tubs instead of gallon jars, because tubs are more convenient to work with and do not
need to be thinned as often. A culture tub is usually started with 400-1000 animals (e.g., 0.5-
1.2 cm"2). In one instance, approximately 4000 healthy animals (e.g., 5 cm"2) were recovered
from one tub. This is the maximum density we have obtained in our cultures; field densities
of 110,000 m"2 (e.g., 11 cm'2) have been reported (Santos and Simon, 1980).
Regular estimates of female fecundity were not obtained from the cultures due to the
sensitivity of the animals to sieving. The brood sizes of eight first generation females ranged
from 1-27 eggs, with a mean clutch of 13.5 (7.2 SD). Field-collected A. abdita had average
brood sizes of 26 (Mills, 1967) and 13.7 (Nelson, 1980). Scott and Redmond (1989) obtained
means of 13.6 and 15.8 eggs/female in A. abdita produced and reared to maturity in the
laboratory. Thus, it is possible to achieve reasonably natural brood sizes in cultured animals,
albeit inconsistently.
Production data from our A. abdita cultures are ambiguous or inconsistent. Maximum
production was about 5 times the original number of animals added to a tub, as was
described previously. Populations crashed beyond recovery in other culture tubs, despite all
attempts to maintain consistent and constant conditions among all tubs. Variation in
production of cultured A. abdita did not appear to be correlated to the density or life stages
of the animals with which cultures were initiated, season (except in case of possible late-fall
water quality problems), type of container, or sediment source. In experiments with
uncontaminated sediment, animals survive, grow, look healthy, and produce eggs and sperm
but rarely produced offspring (see Chapter 3). There appears to be some unidentified
factor(s) which causes reproduction to be inconsistent in this species. One possibility is a
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natural microbial or viral pathogen present in low density in Yaquina Bay water or sediment
to which A. abdita is sensitive, although we have no direct evidence to support this
hypothesis.
Ampelisca abdita may be sensitive to some aspect of our culturing regime, such as the
lack of flowing water in the culture tubs, the omission of a critical nutritional ingredient in
the diet, or the lack of seasonal changes in photoperiod or temperature. No factors were
discovered that distinguished healthy cultures from mediocre or failed cultures. Some culture
containers were quite productive, but the majority were not. A flowing seawater system for
delivery of seawater and algal food daily would be less labor-intensive, and this species might
perform better in a flow-through system. A. abdita has sometimes shown increased
sensitivity to toxicants in a static system compared to flow-through (Word et al., 1989). In
previous efforts to develop a chronic sediment toxicity test with A., abdita, Scott and Redmond
(1989) got this species to reproduce with a 14 h light: 10 h dark photoperiod, which might be
a better approximation of their summer breeding photoperiod in Rhode Island. It also may
be possible to stimulate higher production and synchronize reproduction by mimicking
overwintering: maintain low temperature (e.g., <10°C) and a shorter diurnal period (e.g., 8
h light: 16 h dark) for a few weeks until reproduction is desired, and then gradually increase
the temperature and diurnal period to simulate the onset of spring and, hopefully, stimulate
reproduction.
Further efforts to culture A. abdita might be best conducted at a laboratory near a
natural supply of the species, since one could use flow-through culture conditions and
constantly administer suspended-particulate food. Neither were practical at our laboratory
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due to the large volume of water that would have required treatment to prevent the
accidental release of pathogens or non-indigenous amphipods into the local environment.
1.3.6 HANDLING
Animals for acute sediment toxicity tests were obtained by sieving the contents of each
culture tub through a 0.5 mm screen. Larger adults were excluded by sieving with a 1 mm
sieve. Animals swimming or crawling in the culture tub immediately prior to sieving were
removed by dip net; these were probably either reproductively active adults or stressed
animals. Sieving extracted only about half of the amphipods in a tub; the rest remained in
their tubes. More animals could be coaxed from the tubes by allowing the tube mat to "rest"
for 20-30 min between bouts of sieving. Animals also could be forced to leave from individual
tubes by gently working a probe along the tube toward the opening. This was done under
a dissecting microscope and was very time consuming. The animals were then gently washed
from the sieve into a shallow, flat bottomed glass dish for picking using 28-30%o seawater at
20°C for sieving and washing to minimize stressing the animals.
Healthy A. abdita were light pink and often remained tightly curled. Unhealthy
animals tended to be translucent white and uncurled. As with most other amphipods,
individual amphipods were transferred from the picking dish using a wide bore pipette
(ASTM, 1991). Handling and time-held-without-sediment were minimized. Before use in a
test, amphipods had food available on a daily basis. General observations suggested that A.
abdita were stressed by repeated sieving, and cultures were not used if the culture tub had
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been sieved recently. Detailed handling information on setting up an acute bioassay with
this species is provided in ASTM (1991) and Redmond et al. (1991).
1.3.7 CONCLUSION
Due to some unidentified factor(s) that seem to inhibit reproduction, we have been
unable to produce large numbers of A. abdita on a consistent basis. We have had spectacular
successes followed by complete failure of cultures, with no discernable pattern to explain
these inconsistencies. Further research on culture methods for this organism may be
worthwhile, since it is widely used ir acute sediment toxicity tests, and it may potentially be
used to test the chronic toxicity of sediments (see Section III of this report). Successful
culture methods would allow the development of multiple sources of the animals for toxicity
tests, and eliminate the difficulty of obtaining animals in winter, when they are available but
frequently difficult to collect. Culture research might be more successfully conducted in a lab
near a naturally occurring population of this species, since until the problems in culturing
are identified, large numbers of field-collected animals are required to support investigations.
1.4 Lepidactylus dytiscus
1.4.1 SUMMARY
Lepidactylus dytiscus was collected in high densities from one location near Virginia
Beach, VA, and may be available in high densities in other locations within Chesapeake Bay.
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It was possible to maintain _L. dytiscus in the laboratory, but cultures expanded only very
slowly due to the long time to first reproduction in this species. Furthermore, reproductive
activity was apparently suspended during the winter months despite maintenance of constant
temperature and photoperiod in the laboratory cultures. This increased the time required
for culture expansion and limited the rate at which animals could be harvested for
experiments. Small cultures have been maintained in our laboratory since March, 1990. The
purpose of culturing JL. dytiscus was to assist the development of chronic sediment toxicity
tests, but the prolonged life cycle of this species made L. dytiscus less convenient than other
amphipod species for that purpose. However, this species may be suitable for acute sediment
toxicity tests, and animals from our collections were used successfully in a comparative acute
toxicity-test experiment. Adequate numbers of young JL dytiscus may be produced in the
spring and summer, but reproductive activity was substantially lower or non-existent in fall
and winter. While further research may reveal conditions that would enhance the production
rate of this amphipod, the culturing approach used in this project did not successfully produce
sufficient numbers of animals for experiments on a year-round basis.
1.4.2 OVERVIEW OF THE SPECIES
Lepidactylus dytiscus is a free-burrowing, estuarine, haustoriid amphipod found in fine
sand sediments. They are widely but sporadically distributed in Chesapeake Bay (Fig. 1-3),
and are tolerant of salinities ranging from ca. 5-30%o (Ray Alden & Emily Deaver (Old
Dominion Univ.), pers. comm.). In laboratory cultures, L,. dytiscus preferred a sandy
sediment to organic-rich mud. They appear to be deposit feeders, and do not form tubes in
the sediment or filter particles from the overlying water.
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L. dytiscus is a slow growing species and not amenable to the rapid production of large
numbers of offspring. Time to first reproduction seems to be greater than 6 months: a group
of 19 cultured newborns held for six months had 100% survival, but little growth and no
indication of sexual development. Adult (i.e., sexually mature) animals vary greatly in size
(e.g., 4-12+ mm) and may live for more than a year. Although we have not carefully
documented the life-cycle of this amphipod, females can apparently produce more than one
brood of offspring, but the period between broods is probably several weeks or months.
Reproduction in the cultures ceased during winter months even though constant physical
conditions were maintained (e.g., temperature, salinity and photoperiod).
1.4.3 FIELD COLLECTION
Several hundred L;. dytiscus were collected by T.H. DeWitt and Emily Deaver (Old
Dominion Univ.) on 3/26/90 with a shovel and sieve from intertidal and shallow subtidal (ca -
.25 m) sandy sediments in the Lynnhaven River estuary near Virginia Beach, VA (salinity
ca 28%c). Recent searches for other populations of L. dytiscus in the lower Chesapeake Bay
area have not been successful (E. Deaver and R. Alden, personal communication), but high
densities of this amphipod have been reported elsewhere in the estuary, particularly near the
Calvert Cliffs in Maryland (Fig 1-3).
1.4.4 SHIPPING
Lepidactylus dytiscus were shipped successfully from Chesapeake Bay to Newport, OR,
in March, 1990. Approximately 50-100 animals were packed per plastic container (i.e., a 250-
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1000 ml sandwich box or ice cream tub), which held a 1-2 cm layer of fine sand substrate and
filled to the top with 28%
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the overlying water in each tub was changed every other day, except on weekends. This was
accomplished by pouring off the overlying water and flocculent organic matter on the
sediment surface, and refilling the tub from a plastic pitcher with fresh seawater laden with
the algal food. The stream of incoming water was moved across the bottom of the tub to
slightly agitate and aerate the sediment. The fresh, renewal water consisted of seawater (ca.
32%e), cultured phytoplankton and deionized water which were combined to achieve a salinity
of 20%o and ca 106 algal cells per ml. The algae used were Pseudoisochrysis paradoxa and
Phaeodactylum tricornutum in equal portions by volume. The cultures were also fed about
0.5 g of a dry food (i.e., "gorp") during the water change. Gorp consisted of 48.5% Tetra min®,
24% dried alfalfa, 24% dried wheat leaves and 4.5% Neo-Novum® (a maturation feed for
shrimp mariculture; Argent Chemical Laboratories, Redmond, WA), combined and ground to
a fine powder. The gorp was sprinkled on the sediment surface after the old water was
poured off and before the tubs were refilled.
1.4.6 HANDLING
Culture bins were gently sieved through a 0.5 mm mesh to obtain subadult L_. dytiscus
(i.e., 2-4 mm long) for acute toxicity tests. This screen size retained all of the animals in a
culture. Larger animals were excluded by gently sieving the animals through a 1.5 mm
screen. Animals were rinsed free of sediment and washed into a shallow glass counting dish.
Subadults of a uniform size were selected and transferred by pipette into a smaller glass dish
for acclimation to the test temperature and salinity. Water for sieving and rinsing was
maintained at the same temperature and salinity as the cultures to minimize stress to the
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animals. Unused animals were returned to the culture tub after the sediment bed had
settled.
1.4.7 CONCLUSION
Lepidactylus dytiscus was a hardy amphipod, well suited for acute sediment toxicity
tests in most respects. It is distributed throughout Chesapeake Bay with apparent high
densities near the Calvert Cliffs of Maryland (Fig. 1-3). L_. dytiscus can be cultured, but their
slow somatic growth and long time to first reproduction resulted in low culture productivity.
The culturing approach described here was not successful in producing sufficient numbers
of L/. dytiscus for routine use in sediment toxicity tests. Furthermore, the long lifespan and
slow growth of this species preclude its utility in chronic sediment toxicity tests for which
growth or reproduction are desirable endpoints. Further research might reveal factors to
enhance culture productivity, such as the discovery of a limiting nutrient, or the simulation
of a shortened annual cycle (e.g., changing temperature, photoperiod and, possibly, salinity
to mimic seasonality) to stimulate more frequent reproduction.
1.5 Monoculodes edwardsi
1.5.1 SUMMARY
Monoculodes edwardsi was easily collected in shallow water in Chesapeake Bay and
has some potential for being cultured. Several small culture tubs of M- edwardsi were
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maintained from March, 1990, through September, 1992, but only at a relatively low
population density (e.g., 0.01 cm"2). The animals were very active burrowers and swimmers,
and appeared to require a large amount of space. Females produced eggs frequently but did
not always produce juveniles; the eggs possibly were not fertilized or the embryos did not
develop. Further research might reveal means to enhance productivity, but culturing M.
edwardsi under current conditions failed to produce adequate numbers of animals for routine
sediment toxicity tests.
1.5.2 OVERVIEW OF THE SPECIES
Monoculodes edwardsi is a very active, free-burrowing oedicerotid estuarine amphipod
found in subtidal sandy sediments. It leaves the sediment at night, as evidenced by trails
made on the surface of the sediment. M. edwardsi is distributed from the Gulf of St.
Lawrence to Georgia/N. Florida, and is found also in the Gulf of Mexico. It is an omnivorous
predator, that will opportunistically feed on living or dead animal prey as well as microalgae
and possibly detritus. It has been observed feeding on the remains of conspecifics, but it is
not known if this is evidence of cannibalism (i.e., killing conspecifics for nourishment) or
indiscriminant scavenging.
First reproduction occurs approximately at age 32-41 days, and average brood size is
5.7. Females can apparently produce several broods. Individuals probably do not live longer
than a year, but the life-cycle of this species has not been fully documented.
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1.5.3 FIELD COLLECTION
Approximately 150 Monoculodes edwardsi were collected with a dipnet and sieve in
shallow water (l-2 m)
off Calvert Cliffs, MD, by Tammy Tonare (VERSAR, Columbia, MD) on 3/12/90. The dipnet
procedure was quite simple: the net was rapidly scraped across the sediment surface, just
skimming the top 1-3 mm of substrate. The contents of the net were sieved through the
mesh to concentrate the amphipods. Attempts at collecting the amphipods in shallow water
with a shovel were unsuccessful (relative to the dipnet approach) because the animals were
winnowed from the upper millimeters of sediment as the shovel was drawn to the surface.
M. edwardsi is widely distributed in Chesapeake Bay and should be available for field
collection at many locations (Fig. 1-4). However, M. edwardsi is highly motile, especially
nocturnally, and local population densities might fluctuate substantially within short periods
of time.
1.5.4 SHIPPING
Monoculodes edwardsi were shipped successfully from Chesapeake Bay to Newport,
OR, in March, 1990. Approximately 50-100 animals were packed per plastic container (i.e.,
a 250-1000 ml sandwich box or ice cream tub), which held a 1-2 cm layer of fine sand
substrate and filled to the top with 10%o estuarine water. Field-collected animals were held
overnight or longer in running or frequently changed bay water at a salinity and temperature
close to that where the animals were collected. Dead or injured animals were removed prior
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to packing. Three plastic containers were placed in an insulated cooler along with 3 or 4
freezer packs (such as blue ice) to keep the temperature cool, but above freezing, and then
the cooler was sealed and immediately shipped by overnight delivery. All animals died in one
of the three shipping containers, but survival was high in the remaining containers. The
reason for the failure of one container could not be determined, but this container held mostly
gravid females. Shipping success might be further enhanced if the amphipods were shipped
in oxygen-saturated water.
1.5.5 CULTURING
Monoculodes edwardsi cultures were maintained under static/renewal conditions at
20°C and 20%c seawater on a 16hr light : 8hr dark photoperiod. Each culture was kept in a
29 cm x 34 cm x 13 cm (depth) (i.e., 11.5" x 13.5" x 5.5") plastic tub filled with a 10 cm deep
layer of seawater and a 1.5 cm layer of sand (sieved to <0.25 mm diameter) on the bottom.
The water was constantly aerated with a gentle flow of filtered air. Feeding and water
renewal were conducted simultaneously three times per week. This consisted of a 50-75%
replacement of the old water column with a 1:1 mixture (v/v) of the cultured microalgae
Pseudoisocrysis paradoxa and Phaeodactvlum tricornutum at a density of ca. 106cells/ml.
Cultures were initially also provided with 15 ml of frozen Artemia nauplii, but this was
discontinued with no apparent ill effect. In addition, 0.5 g of "gorp" (e.g., 48.5% Tetra min®,
24% dried alfalfa, 24% dried wheat leaves and 4.5% Neo-Novum® [a maturation feed for
shrimp mariculture; Argent Chemical Laboratories, Redmond, WA] combined and ground to
a fine powder) was sprinkled on the water's surface once a week. It is not known if this was
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an optimal or even sufficient diet for the cultures, but the cultures have been maintained for
longer than 18 mo on this diet.
Culture densities of M. edwardsi were low relative to the other amphipod species
considered in this study. The highest density observed was 0.1 cm"2 (e.g., 130 animals per
bin), but typical densities averaged an order of magnitude lower (e.g., 0.01 - 0.03 cm'2; 10-40
animals per bin). The sex ratio in December, 1991, was 1.4 females/males. Brood size ranged
from 1-12 eggs per gravid female, and females could produce several clutches. Offspring
became sexually mature within 5-7 weeks. Sometimes clutches of eggs appeared to die
within a female's brood pouch: eggs that appeared healthy (e.g., green) one week may turn
black the next. Those eggs may have been unfertilized or did not develop. Juvenile mortality
also may have been high in these cultures, possibly due to cannibalism by adults.
Maintaining cultures that could produce large numbers of animals for routine sediment
toxicity tests (e.g., several hundred juveniles per week) would seem to require a large amount
of space. Further work is needed to better define the culture conditions for this species:
cultures should be able to attain higher densities with this level of fecundity and the
relatively short time to first reproduction. In addition to providing large amounts of space
per individual, better methods for isolating juveniles from adults might reduce juvenile
mortality.
1.5.6 HANDLING
Juvenile M. edwardsi for toxicity tests were extracted from the cultures by sieving the
sediment through a 0.25 mm screen; adults were retained on a 0.5 mm screen. Animals were
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rinsed free of sediment and washed into a shallow counting glass dish from which individuals
of a uniform size were selected and trainsferred by pipette to a smaller glass acclimation dish.
Water for sieving and rinsing was maintained at the same temperature and salinity as the
cultures to minimize the stress to the animals. M. edwardsi were very active, often
swimming and flicking around the counting dish at high speeds. Negatively phototaxic and
individuals could be coaxed to one side of a dish for collection by placing a light source at the
opposite side. Unused animals were returned to the culture tub after the sediment bed had
settled.
1.5.7 CONCLUSION
Monoculodes edwardsi should be readily available for collection in many parts of
Chesapeake Bay. However, laboratory cultures were unable to sustain high population
densities, and they apparently suffered from high rates of juvenile mortality. Current culture
conditions were not capable of producing adequate numbers of animals for routine sediment
toxicity tests. Culture productivity might be improved with knowledge of (1) whether
survival or reproduction were really density dependent as observations suggested, (2) whether
this density dependence was due to space or food limitation, and (3) less stressful means of
separating juveniles from adults. Secondarily, methods should be developed to constrain or
at least slow-down the animals once they are removed from the cultures in order to assist the
distribution of animals to toxicity-test chambers.
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"WASHINGTON, DC
» 00
DISTRIBUTION AND ABUNDANCE OF
Leptocheirus plumulosus
No. Individuals/mS
None
O w-
100-1,000
1,000-10,000
Present
Abundant
1-33
Figure 1-1. Distribution and abundance of Leptocheirus plumulosus in Chesapeake Bay
estuary. Data compiled from several sources, including the US Environmental Protection
Agency's Maryland and Virginia Chesapeake Bay Benthic Monitoring Programs, Dauer et al
(1987), Diaz (1989), Feeley and Wass (1971), Hines and Comtois (1985), Holland (1985),
Holland et al (1977, 1987, 1988), Jordan and Button (1984), Marsh (1988), Mountford et al
(1983), Reinharz and O'Connell (1983), and Schaffner et al (1987). Data from the USEPA's
Maryland and Virginia Chesapeake Bay Benthic Monitoring Programs have been condensed
to average densities per site over the 1984-1988 sampling period. The "Present" and
"Abundant" population density designations were subjectively assigned to sites based on
qualitative descriptions from the literature, personal communications or experience.
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DISTRIBUTION AND ABUNDANCE OF
Ampelisca abdita
No. Individuals/m2
1-34
Figure 1-2. Distribution and abundance of Ampelisca abdita in Chesapeake Bay estuary.
Data compiled from several sources, including the US Environmental Protection Agency's
Maryland and Virginia Chesapeake Bay Benthic Monitoring Programs, Boesh (1973), Dauer
et al (1984), Feeley and Wass (1971), Holland et al (1988), Marsh (1973), Orth (1973),
Reinharz and O'Connell (1983), and Schaffner et al (1987). Data from the USEPA's Maryland
and Virginia Chesapeake Bay Benthic Monitoring Programs have been condensed to average
densities per site over the 1984-1988 sampling period. The "Present" and "Abundant"
population density designations were subjectively assigned to sites based on qualitative
descriptions from the literature, personal communications or experience.
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WASHINGTON, DC
» CK
DISTRIBUTION AND ABUNDANCE OF
Lepidactylus dytiscus
No. Individuals/m2
100-1,000
0)1,000-10,000
f] Present
1-35
Figure 1-3. Distribution and abundance of Lepidactvlus dvtiscus in Chesapeake Bay estuary.
Data compiled from several sources, including the US Environmental Protection Agency's
Maryland and Virginia Chesapeake Bay Benthic Monitoring Programs, Diaz (1989), Jordon
and Button (1984), Lippson et al (1979), and Mountford et al (1977). Data from the USEPA's
Maryland and Virginia Chesapeake Bay Benthic Monitoring Programs have been condensed
to average densities per site over the 1984-1988 sampling period. The "Present" and
"Abundant" population density designations were subjectively assigned to sites based on
qualitative descriptions from the literature, personal communications or experience.
-------
DISTRIBUTION AND ABUNDANCE OF
Monoculodes edwardsi
No. Individuals/m2
Kilometers
A None
10-100
100-1,000
1,000-10,000
Present
Abundant
1-36
Figure 1-4. Distribution and abundance of Monoculodes edwardsi in Chesapeake Bay
estuary. Data compiled from several sources, including the US Environmental Protection
Agency's Maryland and Virginia Chesapeake Bay Benthic Monitoring Programs, Ewing and
Dauer (1982), Feeley and Wass (1971), Holland et al (1987), Loi and Wilson (1979), and
Moutford et al (1977). Data from the USEPA's Maryland and Virginia Chesapeake Bay
Benthic Monitoring Programs have been condensed to average densities per site over the
1984-1988 sampling period. The "Present" and "Abundant" population density designations
were subjectively assigned to sites based on qualitative descriptions from the literature,
personal communications or experience.
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CHAPTER II
THE ACUTE AND CHRONIC SENSITIVITY
OF THE ESTUARINE BENTHIC AMPHIPOD,
LEPTOCHEIRUS PLUMULOSUS,
TO CHEMICALLY-CONTAMINATED SEDIMENTS
2.1 INTRODUCTION
Sediment toxicity tests are a widely used method for estimating the response of
benthic organisms to contaminated sediments. While some researchers have examined
sublethal responses of various benthic taxa to contaminated sediments, most sediment
toxicity tests presently evaluate only the acute mortality of benthic organisms exposed for
short periods of time to contaminated sediment (Swartz, 1987). However, the contaminant
concentration needed to induce mortality may be considerably greater than that needed to
slow somatic growth, reproductive output, or population growth. Benthic organisms living
in contaminated sediments are usually exposed to chemical toxicants for much of their life
cycle, if not for generations. Toxicity tests that reflect both the lethal and sublethal
consequences of long-term exposure to contaminated sediment would thus provide important
information to assist in environmental risk assessment of this polluted sediments.
2-1
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We describe here the development of a chronic sediment toxicity test using a benthic
estuarine amphipod. We focused our attention on benthic amphipod crustaceans because of
their known sensitivity to sediment contaminants and their intimate association with the
substrate. Amphipods are among the most toxicologically sensitive of taxa used to assay
sediments (Nebeker et al., 1984; Reish, 1987; Swartz et al., 1982), and amphipod population
densities decline along pollution gradients in the field (Bellan-Santini, 1980; Chasse, 1978;
Notrini, 1978; Sanders et al., 1980; Seng et al., 1987; Swartz et al., 1982, 1985b). Five
amphipod species were selected for consideration for sediment toxicity test development based
on an extensive literature review of the most abundant amphipods species of mid-Atlantic
estuaries, especially Chesapeake Bay (see Appendix A). Of these species, the burrowing
aorid, Leptocheirus plumulosus, showed especially great promise based on its abundance,
short life cycle (Marsh, 1988), and its distribution relative to major sources of chemical
contamination (see Appendix A). After preliminary tests with all of the amphipod species,
we selected L. plumulosus, for further development based on its culturability, hardiness in
the laboratory, broad salinity tolerance, and apparent sensitivity to chemical contaminants
in the field. The progression of the research program to develop a chronic sediment toxicity
test with L. plumulosus was to (1) determine the appropriate conditions under which toxicity
tests could be conducted with this species by measuring its sensitivity to non-contaminant
variables, such as sediment grain size, TOC, and absence of food; (2) measure its short-term
(i.e., acute) and long-term (i.e., chronic) sensitivities to chemical contaminants spiked into
sediment; and (3) compare its acute and chronic sensitivities to real-world contaminated
sediment. Two 10-d acute responses (i.e., mortality and size) and four 28-d chronic responses
(i.e., mortality, size, fertility, and sex ratio) were examined. We report here the results of
experiments to determine the (1) acute sensitivity of L. plumulosus to sediment geophysical
variables (i.e., grain size, organic carbon, water content, and Eh), different feeding regimes,
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2-3
and three common sediment contaminants (i.e., cadmium, phenanthrene, and acenaphthene);
and (2) its chronic sensitivity to phenanthrene-spiked sediment and a dilution series of
contaminated sediments from Baltimore Harbor. The acute and chronic toxicity-test
methodologies were based on the widely used 10-d amphipod sediment toxicity tests described
in Swartz et al. (1985a), DeWitt et al. (1989), and ASTM (1990b), and the new techniques
should be straight forward for other laboratories to adopt.
2.2 MATERIALS AND METHODS
Toxicity-Test Procedures:
The amphipods were obtained from cultures derived from field-collected Leptocheirus
plumulosus (see Chapter 1). For the 10-d exposures, pre-reproductive individuals 2-4mm in
length were isolated from the cultures by first sieving the amphipods through a 0.5mm screen
(e.g., to remove smaller juveniles), then through a 1mm screen (e.g., to remove larger adults),
and finally selecting smaller animals from the remainder for toxicity testing. For the 28-d
exposures, newly released 1-d old juveniles were obtained from gravid females (see Chapter
1 for handling procedures).
In both types of exposures (i.e., 10-d and 28-d), 20 randomly selected amphipods were
distributed to holding dishes from which the animals were transferred to the exposure
chambers. The amphipods were always double counted prior to the initiation of exposure.
One to three subsets of 20 amphipods were set aside during set-up for measurement (i.e., size
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at T0); these animals were relaxed with carbonated water and preserved in 70% EtOH. The
static exposures were conducted in 1-L beakers containing 175 ml of test sediment and 725
ml of overlying water. Unless other-wise noted, the interstitial and overlying water salinities
were adjusted to 20%o. The compositions of the test substrates varied with each experiment
and are described below. The exposure chambers were placed in temperature-controlled
water baths within vented cabinets. The exposures were conducted at 25°C with a 16-h:8-h
light:dark photoperiod. Each exposure chamber was aerated constantly. Each exposure
beaker was monitored daily for proper temperature, aeration, and amphipod emergence, and
to submerge any amphipods trapped at the water surface.
Amphipods in the 10-d and 28-d exposures were fed 0, 3, 5, or 7 times per week,
depending on the experiment. Feeding included either 400 ml of an algal suspension (106
algal cells/ml, 1:1 v/v mixture of Pseudoisochrysis paradoxa and Phaeodactylum tricornutum)
or 10 ml of a finely-ground dry food (i.e., "gorp": 48.5% TetraMin®, 24% dried alfalfa, 24%
dried wheat leaves, and 4.5% Neo-Novum® [a maturation feed for shrimp mariculture; Argent
Chemical Laboratories, Redmond, WA]) in suspension in 20%o seawater, or both. The
amphipods were fed algae at the time each beaker's overlying water was renewed; 400 ml of
the old water was siphoned off and 400 ml of the algal suspension siphoned in. The gorp was
added as 1 ml of a suspension of dry gorp in seawater at a concentration of 10 mg/ml.
After 10-d exposure, the sediment from each beaker was sieved through a 0.5mm
screen to collect the remaining amphipods. These animals were transferred to glass sorting
dishes from which survivors were counted, relaxed with ca. 10% carbonated water, and
preserved in 70% EtOH for later measurement. After 28-d exposures, the sediments were
sieved through 0.5mm and 0.25mm screens to retain adults and juveniles, respectively.
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Adults were counted and preserved as in the 10-d exposures. Juveniles were too numerous
to count at the time of bioassay breakdown, so the material remaining on 0.25mm screen and
in the sorting dishes was transferred to a vial, stained overnight with a few ml of
concentrated rose bengal in 20%e seawater, preserved the next day in 70% EtOH, and held
until juveniles could be counted at a later time.
Amphipod size was determined as the length of the curved line running dorsally from
the base of the first antennae to the base of the urosome (i.e., posterior end of the third
abdominal segment; Fig. 2-1). The measurements were made with a computer-based image
analyzer connected to a dissecting microscope-mounted video camera. Even though size of
each survivor was measured, only the mean size of all survivors from a replicate (i.e., one
exposure chamber) was used as the size-response endpoint.
Adult amphipods in the 28-d exposures were sexed at the time of measurements.
Revealing sexual characteristics were the presence of eggs in the oviducts or brood pouch
(females), brood plates (females), gnathopod morphology (i.e., a notched palm on the dactyl
and stout 5th and 6th segments) (male), or the presence of penile papillae (males; only visible
in dead animals) (Bousfield, 1973; Fig. 2-1).
The reproductive response of the cohort in the 28-d exposures was reported as fertility,
or the average number of daughters produced per surviving mother. This was calculated
from the number of juveniles and female adults retrieved at the termination of the 28-d
exposures using the following equation:
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Fertility = No. Juveniles/2
No. Surviving Females
This equation presupposes a juvenile sex ratio of 1.0 (females/males). Since juveniles cannot
be sexed presently, the sex ratio must be based on an estimate for the population and will
be a constant in this equation. Thus, the choice of the sex ratio parameter will only affect
the absolute estimate of fertility, and will not affect the relative comparison of fertilities
among experimental treatments.
Control Treatments:
Three types of control treatments were used in the sediment toxicity tests. The first
type was a performance control which tested the response of the amphipods in the absence
of contaminant stress and under the best possible conditions for the amphipods. The
performance control used culture sediment as the test substrate and maintained the same
temperature and salinity as the experimental treatments (i.e., 20%c or 28%o, and 25°C).
Culture sediment was collected from a sandflat adjacent to the lab in Yaquina Bay (South
Beach, OR), sieved to <0.25mm, and stored at 4°C; sediment for the performance controls was
obtained from cold storage, not from the culture bins. The exposure periods were 10- and 28-
d for the acute and chronic toxicity test performance controls, respectively. Performance
controls were used for QA/QC, to assure that the test organisms were healthy.
The second control was a reference toxicant control which tested the sensitivity of the
animals to a single toxicant under repeatable exposure conditions. The reference toxicant
control consisted of 96-h, water-only exposures to cadmium chloride at 20%o and 25°C.
Animals for these controls were selected from the same population as the test animals. The
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reference toxicant controls for the acute sediment toxicity tests were conducted within 4 d of
the start of the sediment toxicity test. The reference toxicant controls for the chronic tests
were initiated 1 wk after the start of the sediment toxicity test because the newborn
amphipods could not survive 96-h without sediment or food, having been released from their
mothers' marsupium for less than 1 d. In this case, the newborns were placed in culture
sediment and fed in the same manner as the amphipods in culture (see Chapter 1) for this
1 wk period, at which time they were sieved from the sediment and randomly allocated to the
different cadmium concentrations. The cadmium concentrations for the these controls ranged
from 0.19 - 6 mg/L, although 30 mg/L was used as the highest Cd concentration for one
experiment. The reference toxicant control was also employed for QA/QC, to determine
whether the sensitivity of the test animals was consistent among experiments.
The third control was a carrier or site control in which the substrate was not spiked
with contaminants, but was manipulated in all other ways the same as the other
experimental treatments. This included sieving, salinity adjustment, addition of the toxicant
carrier, rolling, and storage. These controls were included as the uncontaminated treatment
against which the toxicity of the other experimental treatments were compared statistically.
Geotechnical Analyses:
Sediment particle-size was measured by the sieve/pipette method (Buchanan, 1984), and
sediment water content was measured as the percentage of weight lost upon drying overnight
at 90°C.
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Chemical Analyses:
Substrates for chemical analyses were spooned into separate 1-L beakers at the same
time as the toxicity-test beakers were filled. Chemistry beakers were removed from the
exposure tables on day 0 of the exposures immediately prior to adding amphipods to the
toxicity-test beakers, and the water overlying the sediment was aspirated off. Approximately
25 g of wet substrate was collected and stored frozen in glass vials for later measurement of
total-sediment organic contaminants and total organic carbon (TOC) concentrations. For
metals analyses in experiments using field-collected contaminated sediment, a sample of the
substrate for acid volatile sulfide (AVS) and simultaneously extracted metals (SEM) was
withdrawn from the beaker into the open barrel of a 10 cc plastic syringe. Parafilm was
secured over the open end of the syringe, and the sample was frozen. The sample was
shipped frozen to the EPA laboratory in Narragansett, RI, for analysis of AVS and SEM.
Total-sediment organic contaminants were extracted from stored sediment samples
by the method of Ozretich and Schroeder (1986) utilizing acetonitrile, sonication and cleanup
on C-18 solid phase extraction cartridges. Sediment samples were spiked prior to extraction
with d10-acenaphthene, d10-phenanthrene, or other deuterated organic compounds (depending
on the experiment and chemicals being measured) allowing quantitation by the method of
surrogate internal standards. Quantitation of PAH was accomplished using a Hewlett-
Packard 70B Gas Chromatograph-Mass Selective detector equipped with a 0.25mm ID x 30m
or 60m fused silica, DB-5 coated, capillary column (J & W Scientific).
Substrate TOC was determined using high temperature combustion thermal
conductivity detection with a Perkin Elmer Model 2400 CHN analyzer. Samples were
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acidified to pH <2 prior to measurement (to liberate inorganic carbonate as C02), and the
TOC measurements were calibrated using NBS acetanilide as the standard.
The AVS was determined by converting the solid-phase sulfide to hydrogen sulfide
(H2S) using cold 6 M HC1. The released H2S was trapped in sulfide anti-oxidant buffer, and
the sulfide measured with a sulfide-specific electrode. The SEM were determined by
inductively coupled plasma spectrometry from a filtered sample of the sediment/acid solution
after the AVS was released (Di Toro et al., 1990).
Substrate Eh was measured at the beginning and end of the exposure period in each
treatment to determine if the sediment remained aerobic. Eh was measured with a platinum
redox electrode which was inserted ca. 1 cm below the sediment surface and allowed to
equilibrate for 1-2 minutes, until the reading stabilized. Dissolved oxygen levels of the water
overlying the sediment was also measured at the beginning and end of the exposure using
a DO electrode.
EXPERIMENTS:
Sensitivities of Sub-Adults and Newborns to Non-Contaminant Variables
Two sets of experiments examined the effects of sediment variables and feeding
regimes on survival and growth of Leptocheirus plumulosus. The first experiment examined
the sensitivity of sub-adult amphipods (i.e., those used in 10-d acute sediment toxicity
exposures) to sediments collected at 12 sites in the Yaquina R. and Alsea R. estuaries in
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Lincoln Co., OR. These sediments ranged from a fine sand to very silty mud, encompassing
TOG concentrations from 0.4% to >4%. Each treatment was replicated three times, and the
exposure period was 10-d. Sensitivity of mortality to several sediment variables (i.e., median
grain size, percent fines [< 63 um], water content, TOC content) was assessed by correlation
analysis.
The second experiment examined the sensitivity of 1-d old newly released juveniles
(i.e., the age class used in the 28-d chronic sediment toxicity exposures) to sediment from five
sources (e.g., culture sediment [from a sandflat in South Beach, ORJ, Eckman Slough,
McKinney Slough, East Log Pond, and Curt's Mud Hole [the latter two were from South
Beach, Yaquina Bay]) and five feeding regimes (e.g., 106 algal cells/ml, 105 algal cells/ml, 104
algal cells/ml, 5 mg gorp, and no food). The field-collected sediments were sieved through a
0.25mm screen; additionally, sediment from Eckman Slough and Curt's Mud Hole was sieved
through only a 1.0mm screen, creating two more sediment treatments. This difference in
processing allowed us to examine whether forcing sediment through the 0.25mm screen
altered its suitability to the amphipods. Culture sediment sieved through a 0.25mm screen
was used in the feeding treatments. Each sediment source, handling, and feeding treatment
was replicated three times. The reference toxicant control LC50 could not be determined
since all newborns died. In this experiment, 1-d old newborns were added to the water-only
beakers; later experiments called for newborns to be held for 1 wk under culture conditions
prior to exposure to the reference toxicant control conditions.
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The Life History and Demography of Leptocheirus plumulosus
This experiment followed the full lifespan of a cohort of JL. plumulosus in order to
chronicle the life history of this amphipod. One hundred newborn amphipods were randomly
distributed among 5 beakers holding culture sediment. These beakers were fed microalgae
and gorp ad libidum daily. After 14 d, and every 7 d thereafter, the amphipods were sieved
through a 0.5mm sieve and a 0.25mm sieve to separate adults and newborns, respectively.
Newborns were preserved in 70% EtOH. The adults were measured alive with an image
analyzer connected to a dissecting-scope-mounted video camera, and then returned to the
beaker with the same culture sediment from which they had been sieved. In this way,
weekly mortality, production of offspring, and size distribution of adults was monitored.
Acute Toxicity of Cadmium
The first experiment compared the acute cadmium sensitivity of sub-adult
Leptocheirus plumulosus with two Pacific coast amphipods (Rhepoxynius abronius and
Eohaustorius estuarius) and three Atlantic coast amphipods (Ampelisca abdita, Lepidactylus
dytiscus, and Monoculodes edwardsi) in simultaneous, static, 96-h, water-only exposures. The
purpose of the water-only exposure was to compare these species' relative sensitivities to a
reference toxicant under similar contaminant bioavailability regimes. Since salinity is known
to modify the bioavailability of cadmium through regulation of free ion concentration, three
species (Eohaustorius, Lepidactylus, and Leptocheirus) were exposed to cadmium at two
salinities (i.e., 28%c and 20%c); the other species were exposed only at 28%o salinity.
Furthermore, since temperature may modify a species' sensitivity by altering its metabolic
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rates, Leptocheirus was exposed to cadmium at two temperatures (i.e., 15°C and 20°C); all
other species were exposed at 15°C.
The second experiment examined the acute Cd sensitivity of newborn L,. plumulosus
that had been held for 1 wk after release from their mothers' marsupium. In essence, this
experiment established the protocol for the 28-d reference toxicant control for the chronic
sediment toxicity test. The experimental conditions were the same as described previously
for that control treatment. The nominal Cd concentrations were 0, 18.8, 37.5, 75, 150, 300,
and 600 ug/L. Two replicates were run for each concentration.
Acute Toxicity of Acenaphthene
These experiments examined the sensitivity of L. plumulosus to acenaphthene-spiked
sediment in 10-d static exposures. This polynuclear aromatic hydrocarbon (PAH) is a
common contaminant of sediments near urban and industrial areas, being derived from
petroleum or the combustion of organic materials. In the first sediment exposure, L.
plumulosus was exposed to seven treatments (six nominal acenaphthene concentrations and
a carrier control) in each of three sediments. The three sediments covered a range of
sediment textures and TOG concentrations, and were selected to examine the effect of TOC
on the bioavailability of acenaphthene. The three sediments were collected from the Yaquina
R. and Alsea R. estuaries in Lincoln Co., OR: South Beach (SB: very fine sand, poorly sorted;
TOC = 0.8-1.6%), McKinney Slough (medium silt, poorly sorted; TOC = 2.4-2.5% ), and
Eckman Slough (medium silt, very poorly sorted; TOC = 3.0-3.7%). Each sediment was sieved
through a 500 um screen to remove macrofauna, adjusted to 28%c, and stored at 4°C for 6-d
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at which time acenaphthene was added. Preliminary flow-through, water-only exposures at
25°C and 28%c with acenaphthene indicated that the 10-d LC50 for L,. plumulosus in
seawater was 678 ug/L (Swartz et al., unpublished data). The spiking concentrations for each
sediment were calculated from equilibrium partitioning equations (assuming log Koc= 3.511)
so that the sediment interstitial water (IW) concentration of the median treatment
concentration was approximately the LC50. Each sediment was split into aliquots and spiked
with acenaphthene to achieve the following nominal sediment concentrations: South Beach:
0, 7.0, 11.7, 19.4, 32.4, 54.0, and 90.0 mg/dry kg; McKinney SI: 0, 19.4, 32.4, 54, 90, 150, 250
ing/dry kg; Eckman SI.: 0, 32.4, 54, 90, 150, 250, 416 mg/dry kg. Spiking was accomplished
using the methods of Ditsworth et al. (1990) by rolling sediment at room temperature
intermittently over a 24-hr period in ca. 2-L glass jars which had the requisite acenaphthene
plated onto the inside walls of the jars. The spiked sediments were then stored for 8 d at 4°C
to allow acenaphthene to equilibrate between particulate-sorbed and IW phases. Each
treatment had two replicates for toxicity and one for chemistry. The amphipods were fed
gorp daily in this experiment. It would have been a better decision not to feed the amphipods
in this experiment for more direct comparison with the other acenaphthene and
phenanthrene acute toxicity experiments.
A second acenaphthene-spiked sediment exposure was conducted in which L,.
plumulosus was exposed to higher concentrations of acenaphthene-spiked sediment since high
mortalities were not observed in the initial experiment. The sediments and nominal total
sediment acenaphthene concentrations were: South Beach (0, 150 and 250 mg/dry kg),
McKinney SI. (0 and 416 mg/dry kg), and Eckman SI. (0 and 693 mg/dry kg). The amphipods
were not fed in this experiment.
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Acute Toxicity of Phenanthrene
These experiments examined the sensitivity of L. plumulosus to phenanthrene-spiked
sediments in 10-d static exposures. The experimental design was virtually identical to the
acenaphthene-sensitivity experiment described above, including exposure to sediments from
South Beach (TOC = 0.8-2.0%), McKinney Slough (TOC = 2.4-2.5%), and Eckman Slough
(TOC = 3.0-3.6%). The phenanthrene concentrations were likewise selected to bracket L.
plumulosus's 10-d, flow-through LC50 for phenanthrene at 25°C and 28%o (i.e., 180 ug/L;
Swartz et al., unpublished data) using the equilibrium partitioning model (Di Toro et al.,
1991), the TOC concentration for each sediment, and a log Koc = 4.065 for phenanthrene. The
three sediments were spiked to the following nominal, total-sediment phenanthrene
concentrations: South Beach (0, 7.0, 11.7, 19.4, 32.4, 54, and 90 mg/dry kg), McKinney SI. (0,
19.4, 32.4, 54, 90, 150, and 250 mg/dry kg), and Eckman SI. (0, 32.4, 54, 90, 150, 250, and 416
mg/dry kg). Sediments were sieved through a 0.5mm screen and adjusted to 28%o, spiked
with phenanthrene, stored at 4°C for 13-15 d to equilibrate, at which time the exposure
beakers were loaded with substrate. Each experimental treatment had two replicates for
toxicity and one for chemistry. The amphipods were not fed in this experiment.
A second phenanthrene-spiked sediment experiment was conducted in which L.
plumulosus was exposed to higher concentrations of the PAH since high mortalities were not
observed in the initial experiment. The sediments and nominal total sediment phenanthrene
concentrations were: South Beach (0 and 150 mg/dry kg), McKinney SI. (0 and 416 mg/dry
kg), and Eckman SI. (0, 416, and 693 mg/dry kg). The amphipods were not fed in this
experiment.
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Acute and Chronic Toxicity of Phenanthrene-Spiked Sediment
The objective of this experiment was to compare the 10-d and 28-d responses of
Leptocheirus plumulosus to phenanthrene-spiked sediment and to determine whether
handling amphipods after 10-d of exposure affected their sensitivity to phenanthrene after
28-d of exposure. One day-old amphipods were exposed to 9 concentrations of phenanthrene
spiked into fine-grained sediment from Eckman Slough (Alsea R. estuary, Lincoln Co., OR;
TOG = 2.44%) for 10 d or 28 d. The nominal total-sediment phenanthrene concentrations
were 0, 25, 35, 50, 72, 103, 147, 210, and 300 mg/dry kg. Three replicate beakers were set
up for each concentration. The IW and overlying water were maintained at 20%e and 25°C.
The amphipods were fed algae and gorp every other day during this experiment.
One replicate of each of the 9 concentrations was exposed for 10 d to measure acute
toxicity, after which the survivors were transferred to replacement beakers containing fresh
sediment for an additional 18-d exposure to the same phenanthrene concentrations. The
replacement beakers were set up at the same time as all other beakers, but amphipods were
not added until day 10. L. plumulosus in these beakers would be used to measure chronic
toxicity plus the effects of handling on toxicity. L_. plumulosus in two other sets of replicate
beakers were exposed undisturbed for 28 days to measure chronic toxicity. At the end of the
10-d and 28-d exposures, the amphipods were sieved from each beaker through a 0.25mm
screen, and the following data were collected: 1) survival of the initial cohort, 2) sizes and
sexes of the survivors, and 3) the number of offspring. Sediment chemistry samples were
collected on day 0 for all concentrations (except 210 mg/dry kg at 28 d). Chemical analyses
were conducted to measure total sediment and IW phenanthrene, TOC, Eh, and overlying
water DO.
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Acute and Chronic Toxicity of Field-Collected Sediment
The purpose of this experiment was to compare the sensitivities of the 10- and 28-d
responses of L. plumulosus to field-collected, contaminated sediment from Chesapeake Bay.
Since the toxicities of field sediments in Chesapeake Bay were generally unknown, a dilution
series was prepared from mixtures of a chemically contaminated sediment from Curtis Cr.,
Baltimore Hbr., MD, and uncontarninated sediment from Corsica R., MD. The Curtis Cr.
sediment was known to be heavily contaminated with a complex mixture of metals and
organic chemicals and acutely toxic to field-collected L. plumulosus, and the IW salinity and
median sediment grain size of Curtis Cr. sediment was expected to be similar to that of the
Corsica R. sediment (Schlekat et al., 1992; McGee et al., in press; E. Reinharz, B. McGee (MD
Dept. of Environment) pers. comm.). Personnel from the Maryland Department of
Environment prepared six concentrations for the dilution series: 100% (Curtis Cr.), 50%, 25%,
12.5%, 6.25%, and 0% (= 100% Corsica R.). The sediment IW salinity was 11%0 for all
dilution treatments. The substrates were color coded by Claudia Walters (EPA Quality
Assurance officer) so that the test was conducted in a blind fashion; only she knew the cipher
until the termination of all toxicity tests. Sufficient sediment to test three replicate samples
of each substrate with each toxicitjr test was shipped on ice to our laboratory and stored at
4°C. Substrate was added undisturbed (i.e., no homogenization) to each exposure chamber
within 7-d of dilution; the overlying water was ll%c to match the sediment IW salinity. The
performance and reference toxicant controls were conducted at 20%c, and the site control (i.e.,
the 0% Curtis Cr. treatment) and dilution treatments were conducted at ll%c. The dilution
treatments and controls were conducted at 25°C, with the exception of the reference toxicant
controls which were conducted at 2()°C. The reference toxicant controls for both the 10-d and
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28-d sediment toxicity tests consisted of 96-h, water-only exposures at 0, 0.18, 0.50, 1.40,
3.90, 10.8, 30.0 mg/L Cd. L/. plumulosus in the 10-d exposures were not fed, but those used
in the 28-d exposures were fed algae and gorp three times per week.
One additional beaker for each dilution treatment was loaded with sediment for
chemistry samples. Samples for total-sediment organic chemicals, AVS, and SEM analyses
were collected from the chemistry beakers immediately prior to starting the exposures.
Sediment IW chemical analyses were not conducted.
2.3 RESULTS
Sensitivities of Sub-Adults and Newborns to Non-Contaminant Variables
Although there was some variability in the survival of Leptocheirus plumulosus sub-
adults among sediments from 12 sites in the Yaquina R. and Alsea R. estuaries (Table 2-1),
mortality after 10-d was not significantly correlated with any sediment variable (Table 2-2).
Mortality was >15% in 14 of 36 replicates. Mean mortality was >15% in 5 sediments (i.e.,
East Long Pond 3 and 5, Eckman SI. 2, McKinney SI., and South Beach "Old Log"). Mean
mortality in all 12 sediments was 17.9%. All sites were believed to be substantially free of
chemical contamination, based on previous chemical analyses (R. Ozretich, unpubl. data) and
the lack of local industrial activity. The cause of the higher mortality was not apparent, and
was not explained by the sediment variables we measured. The reference toxicant control
LC50 for sub-adults = 3.15 mg/L Cd (95% CI: 2.33-4.25 mg/L).
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Mortality of newborn _L. plumulosus was <17% for all but one of the seven substrates
in the second sediment source and handling experiment, but mortality was not significantly
different among the treatments (Table 2-3). The size of the newborn amphipods
approximately doubled during the 10-d exposure, but also was not significantly different
among the sediment source and handling treatments. Neither mortality nor growth was
correlated with any sediment variable, although size was negatively correlated with mortality
(Table 2-4). The highest and most variable mortality and slowest and most variable growth
was found in the sandiest substrate, Curt's Mud Hole
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The Life History and Demography of Leptocheirus plumulosus
The five replicates of the cohort of .L. plumulosus showed very similar life history
behavior over 12 wk (Fig. 2-2), which was somewhat surprising since feeding was
unintentionally suspended after week 8. Mortality was very low over the first 3-5 wk, then
senescence, and possibly starvation, led to the slow but steady decline in abundance. Growth
was very rapid over the first 4 wk, then asymptoted at approximately 6.5mm. The first brood
was produced at approximately 3 wk of age, and fertility (i.e., the number of female offspring
born per female) increased with the age of the amphipods. Despite the discontinuation of
feeding, L. plumulosus continued to produce large broods through weeks 10 and 11. It is
probable that longer lifespans and prolonged reproductive periods could have been achieved
had feeding been continued.
Acute Toxicity of Cadmium
Six marine and estuarine amphipod species, including sub-adult L. plumulosus, had
comparable acute sensitivity (i.e., within an order of magnitude) to cadmium in seawater
under certain temperature-salinity conditions when the toxicant was adjusted to its free ion
concentration (Table 2-6). L. plumulosus, Ampelisca abdita. Rhepoxynius abronius. and
Monoculodes edwardsi all had free ion LCSO's between 0.01 and 0.09 mg/L Cd2+, although the
environmental conditions differed under which these highest sensitivities were achieved for
each species. One-week old Li. plumulosus were approximately ten times more acutely
sensitive to Cd in water than were sub-adult L. plumulosus (compare Tables 2-6 and 2-7),
although the exposure with the 1-wk-old amphipods was conducted at a higher temperature
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(i.e., 25°C vs 20°C). Note that the sources of the species varied (i.e., R. abronius and K
estuarius were collected from the field within 4 d of testing, A. abdita and 1-wk-old L.
plumulosus were taken from cultures, and sub-adult L. plumulosus. M. edwardsi. and L.
dytiscus were taken from the stock used to start cultures, several weeks after collection from
the field) and the toxicological comparison was made only once. The relative sensitivities of
these species might vary seasonally or between field-collected and cultured animals.
Acute Toxicity of Acenaphthene
In 10-d exposures with sub-adult L. plumulosus. mortality increased and size
decreased as acenaphthene concentrations increased in three sediments (Tables 2-8 and 2-9).
Mortality and size showed comparable statistical sensitivity to acenaphthene concentration:
in each sediment, one or two acenaphthene concentrations were found to cause significantly
higher mortality or lower growth than the carrier control treatment. The LC50 of
acenaphthene increased with sediment TOG content for at least two of the sediments (i.e.,
McKinney Slough and Eckman Slough), but the LC50 for acenaphthene in South Beach
sediment could not be calculated (Fig. 2-3). Body size declined as a function of acenaphthene
concentration, but as the TOG content of the sediment increased, higher concentrations of
acenaphthene were required to cause a decrease in growth (Fig. 2-4).
The reference toxicant control LC50 for the first experiment could not be calculated
due to high mortality in all concentrations. The reference toxicant control LC50 for the
second experiment was 0.69 mg/L Cd (95% CI: 0.49-0.97 mg/L).
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Acute Toxicity of Phenanthrene
As with acenaphthene, mortality was enhanced and size depressed as the
concentration of phenanthrene increased (Tables 2-11 to 2-13). Mortality and size showed
comparable statistical sensitivity to phenanthrene concentration in two sediments (i.e.,
McKinney Slough and Eckman Slough; Tables 2-12 and 2-13), but size was significantly
depressed in four of seven phenanthrene concentrations in the South Beach sediment while
mortality was significantly higher in only one concentration (Table 2-11). As with
acenaphthene, The LC50 of phenanthrene increased with sediment TOC content (Fig. 2-5)
as was expected from equilibrium partitioning models. Body size declined as a function of
phenanthrene concentration, but relatively higher concentrations of the PAH were needed
to elicit a decrease in growth as the TOC content of the sediment increased (Fig. 2-6).
The reference toxicant control LC50 for the first experiment was 0.90 mg/L Cd (95%
CI: 0.61-1.33 mg/L), and 0.69 mg/L (95% CI: 0.49-0.97 mg/L) for the second experiment. Note
that the reference toxicant control for the second acenaphthene acute sediment toxicity test
was the same control used for the second phenanthrene acute sediment toxicity test.
Acute and Chronic Toxicity of Phenanthrene-Spiked Sediment
Mortality of newborn JL. plumulosus increased as a function of the concentration of
phenanthrene after 10-d and 28-d exposures to spiked culture sediment (Table 2-14). The
concentration-mortality responses of the amphipods were very similar despite the nearly 3-
fold difference in exposure period: the 10-d and 28-d LC50's were 161.20 mg/dry kg (95% CI:
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106.4-244.3) and 177.02 mg/dry kg (95% CI: 165.60-189.22). Most, if not all, of the mortality
apparently occurred during the first 10 d, since mortality did not change between day 10 and
28 in those replicates that were observed on day 10, transferred to new sediment, and
observed again on day 28 (Fig. 2-7). Mortality after 28-d was significantly different from the
carrier control mortality only in the highest phenanthrene concentration (i.e., 184 mg/ dry
kg) (Table 2-14).
Size was affected very little by the concentration of phenanthrene after either 10-d or
28-d of exposure (Table 2-14, Fig. 2-7), and was not significantly different in the spiked-
sediment treatments relative to the carrier control. Under the performance and carrier
control conditions, the newborn amptiipods doubled in length after 10-d and more than tripled
in length after 28-d. The slopes of concentration-size response were virtually flat for both the
10-d and 28-d exposures and were not significantly different, although the y-intercepts
differed because animals in the 28-d exposures had time to grow larger.
Fertility was significantly lower in all phenanthrene concentrations, relative to the
carrier control (Table 2-14). Fertility was reduced by approximately 30-40% in the lower four
phenanthrene concentrations and by 45-60% in the highest four phenanthrene concentrations,
but no distinct concentration-response was observed beyond that (Fig. 2-7). Fertility was
nearly an order of magnitude more sensitive to phenanthrene than was mortality or growth.
Sex ratio did not vary significantly with phenanthrene concentration (Table 2-14), nor
was sex ratio significantly different from 1.0 within any treatment. However, females
outnumbered males in 66% of the 32 beakers used for the whole experiment for a grand mean
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sex ratio of 1.45 (0.79 SD) which was significantly different from 1.0 (G-test: pooled G = 6.64,
1 df, p<0.05; Sokal and Rohlf, 1981).
Handling the amphipods did not affect the toxicological sensitivity of the amphipods.
Sieving, counting, and measuring the amphipods after 10 d exposure did not change the slope
or y-intercept of the concentration-mortality, -growth or -fertility regressions relative to
animals that were not handled during the experiment (Fig. 2-8; analysis of covariance:
p»0.05).
An LC50 for the reference toxicant control could not be calculated in this experiment
because mortality was >80% for all concentrations including 0 mg/L Cd. The high mortality
was probably due to starvation, since the newborns used in these water-only exposures were
not provided contact with sediment subsequent to their release from the maternal brood
pouch. Subsequent to this experience, newborn L. plumulosus were allowed to grow for 1 wk
under culture conditions prior to conducting the reference toxicant exposure.
Acute and Chronic Toxicity of Field-Collected Sediment
Curtis Cr. sediment was considerably more heterogeneous in texture and chemical
content than was the Corsica R. sediment that was used as a dilutant in this experiment.
The Curtis Cr. sediment was characterized as a very poorly sorted medium silt punctuated
with gravel, a high TOC content (Table 2-15), a strong diesel oil smell, and an oily sheen.
The Corsica R. sediment was characterized as very fine silt lacking gravel, a moderate TOC
content (Table 2-15), and no chemical odor or appearance. Extremely high concentrations of
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PAHs (Table 2-16) and metals (Table 2-17) were found in the 100% Curtis Cr. sediment,
while the dilutant sediment from Corsica R. (i.e., 0% Curtis Cr. treatment) had much lower
concentrations of both classes of contaminants. The concentrations of the PAHs in the
remaining substrates increased in direct proportion to the amount of Curtis Cr. sediment that
had been added. The SEM concentrations increased as a function of the concentration of
Curtis Cr. sediment except in the 0% and 6.25% treatments. The total-SEM/AVS ratio was
well below 1.0 for all six experimental substrates, and thus would not be expected to exert
substantial toxicological impact (Di Toro et al, 1990).
For both the acute and chronic sediment toxicity tests, mortality increased and both
size and fertility decreased as a function of the percentage of Curtis Cr. sediment and
chemical contaminants in the substrate (Fig. 2-9). In the 10-d exposure, L. plumulosus
mortality was significantly higher in the 50% and 100% Curtis Cr. substrates, relative to the
site control (i.e., 0% Curtis Cr. = 100% Corsica R. sediment), but size was not significantly
different among the treatments (Table 2-18). In the 28-d exposure, the 100% Curtis Cr.
sediment caused significantly higher mortality and decreased size and fertility of
Leptocheirus plumulosus relative to the site control (Table 2-18). However, no other chronic
test responses were significantly different from the site control for any of the other
treatments, with the exception of decreased size in 50% Curtis Cr. sediment. Since there was
an obvious and significant trend between Curtis Cr. sediment concentration and 28-d
mortality, size, and fertility (Fig. 2-9), the failure of anova to detect differences among
treatments was probably partly due to low statistical power due to insufficient replication
(i.e., N=3) and high variability in the responses, particularly for mortality and fertility which
had coefficients of variation (CV) ranging from 10.3-87% and 12.7-710%, respectively.
Fertility in the carrier control was only half that of the performance control. The two
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treatments differed with respect to sediment source (i.e., Chesapeake Bay sediment and
South Beach, OR, sandflat sediment, respectively) and interstitial salinity (i.e., ll%o and
20%c, respectively).
The sex ratio of _L, plumulosus exposed for 28-d did not differ significantly among the
sediment dilution treatments (Table 2-18), nor was sex ratio different from 1.0 within any
treatment. However, as in the 28-d phenanthrene-spiked sediment experiment, females
outnumbered males in 73% of the 22 exposure chambers, and the grand mean sex ratio for
this experiment was 1.64 (0.89 SD) which was significantly different from a 1:1 ratio of
females to males (G-test: pooled G = 10.42, 1 df, p«0.05).
The reference toxicant control, 96-h LC50 for the 10-d exposure using sub-adults was
0.61 mg/L Cd (95% CI: 0.43-0.66 mg/L). The reference toxicant control LC50 for the 28-d
exposure was 0.27 mg/L Cd (95% CI: 0.20-0.38). The 28-d reference toxicant control used
newborns from the same cohort used in the sediment exposures which were then held for 1
wk in culture sediment prior to the water-only reference toxicant control exposure.
2.4 DISCUSSION
The mortality, growth, and fertility of newborn Leptocheirus plumulosus were affected
by 28-d exposures to high concentrations of sediment-associated phenanthrene and field-
collected sediment from a highly contaminated site in Chesapeake Bay. Shorter-term
exposures (i.e., 10-d) of sub-adult L. plumulosus to sediment-associated acenaphthene,
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phenanthrene and the polluted Chesapeake Bay sediment also affected mortality and growth;
reproduction was not recorded in the 10-d exposures since the test was designed to minimize
the likelihood that broods would be released during the exposure. The sensitivity of the 10-d
and 28-d tests were similar, particularly with respect to mortality and growth. Fertility, the
number of juveniles produced per female in an exposure chamber, was considerably more
sensitive than mortality or growth in one experiment, but not in a second experiment. These
findings culminate in the establishment of acute and chronic sediment toxicity tests for the
estuarine, benthic amphipod, Leptocheirus plumulosus.
The Sediment Toxicity Test Methodologies:
Procedures to conduct 10-d acute- and 28-d chronic sediment toxicity tests with
Leptocheirus plumulosus were developed from the research reported here. The 10-d acute
sediment toxicity test method has since been developed as an appendix to the ASTM
"Standard guide for conducting solid-phase 10-day static sediment toxicity tests with marine
and estuarine amphipods" (ASTM, 1990b). This protocol was coauthored by B.L. McGee and
C.E. Schlekat (Maryland State Department of Environment, Baltimore, MD) with assistance
from T.H. DeWitt, (Oregon State University, Newport, OR), M.S. Redmond and J.E. Sewall
(AScI, Newport, OR), and J.O. Lamberson (U.S. EPA, Newport, OR), and is presented in
Appendix C of this report. The 28-d chronic sediment toxicity test method has been
developed as a "Research Methodology to Assess the Chronic Toxicity of Marine and
Estuarine Sediments with the Benthic Amphipod, Leptocheirus plumulosus" (Appendix D).
Both methods used animals from laboratory cultures, and the procedures for culturing L_.
plumulosus were described in Chapter 1. The bioassay procedures used within some of 10-d
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and 28-d experiments reported here varied somewhat from these proposed "Research
Methodologies" with respect to the salinity and feeding regime, but these differences reflect
the evolution of the techniques which was the goal of this research.
The bioassay procedures using the 10-d and 28-d experiments were simple
modifications of the standard amphipod sediment toxicity test procedures (ASTM, 1990b).
The 10-d sediment toxicity test primarily used sub-adult amphipods (i.e., 2-4mm long) from
cultures (although newborn juveniles were used instead in some experiments), in static
exposures, usually without feeding. Schlekat et al. (1992) described a similar 10-d sediment
toxicity test with L,. plumulosus, except that they used only 10 animals that were larger (i.e.,
4-8mm) and collected from the field, 1 qt jars instead of 1 L beakers were used for the
exposure chambers, the exposure temperature was 20°C, and their Cd 96-hr water-only
reference toxicant controls were conducted at 6%o. Our two laboratories have designed a
single 10-d sediment toxicity test which has been approved by ASTM and will be included in
the next issue of the ASTM amphipod sediment toxicity test guidelines, i.e., ASTM E-1267-92,
which should be published in 1993. The 28-d sediment toxicity test required 0-d old newborn
juveniles in static-renewal exposures, and the animals were fed three to seven times per week
on a mixture of cultured phytoplankton and/or dried food (i.e., "gorp"). The sediment from
the 28-d bioassay was sieved through a 1.0mm and a 0.25mm screen to capture the surviving
adults (regeneration) and their offspring, respectively, whereas the sediments in the 10-d
test were only washed through a 0.5 mm screen to capture survivors. Juveniles captured on
the 0.25mm screen were relaxed, stained, and preserved prior to counting, whereas the adults
were counted live before preservation. The body lengths of the initial cohort used to seed the
exposure beakers and the adult survivors were measured in most of the 10-d and 28-d
experiments. In the 28-d exposures, the F0 survivors were also sexed. These methods
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2-28
entailed modest modifications of the standard amphipod sediment toxicity tests, and should
not be difficult to implement in other laboratories that have experience with amphipod
sediment toxicity tests.
Response to Control Conditions
Performance Controls: Mean mortality in the performance controls ranged from 0% to 11.7%
in the 10-d exposures and from 8% to 12% in the 28-d exposures (Table 2-19). Mortality was
as high as 15% in one replicate in almost all of the performance control trials, and as high
as 20% to 30% in one replicate each of two 28-d exposures. Performance control mortality
did not vary significantly as a function of salinity, feeding regime, exposure period, or age at
T0 (anova; p>0.05) Preliminary studies suggest that newborn Leptocheirus plumulosus were
very sensitive to temperature changes; (J.E. Sewall, AScI, pers. comm.). Thus, maintaining
a constant 25°C while selecting and handling the amphipods during bioassay set up may
reduce performance control mortality for the 28-d test. The performance control mortality
of the 10-d and 28-d toxicity tests should decrease as more experience is gained with these
tests and this species, as has frequently been the case for other amphipod sediment toxicity
tests (J.O. Lamberson, EPA, and M.S. Redmond, AScI, pers. comm.). ASTM guidelines for
amphipod acute sediment toxicity tests allow for up to 20% performance control mortality in
individual replicates as long as the mean mortality among the replicates is <10% (ASTM,
1990b). No general guidelines have been established for chronic amphipod bioassays, and
while it is not possible to predict whether the criteria used for the 10-d test will apply to the
28-d test, it seems likely that low control mortality can be achieved with the chronic L.
plumulosus bioassay.
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2-29
Size of amphipods in the performance controls varied from 2.8 to 7.6mm (Table 2-19),
which was largely a reflection of the initial size of the amphipods1 and the duration of the
exposure. Either salinity or the addition of food (irrespective of quantity, quality, or
frequency) significantly affected 10-d performance control growth (i.e., the difference in size
between amphipods at the start and end of the exposure) (Fig. 2-10 and 2-11). Salinity and
food presence were confounded (i.e., 28%e = No Food, 20%e = Food Added) in most of the 10-d
experiments (Table 2-19), so it was not possible to identify the dominant factor affecting
growth. In the 28-d performance controls, salinity was 28%c and food was added.
Performance control fertility was quite consistent among the three, 28-d exposures in which
it was measured (Table 2-19) and not obviously affected by any of the environmental
variables measured.
The performance control is used as a QA/QC control for the health of the amphipods.
Obviously, L. plumulosus was sensitive to changes in either the feeding regime or the
salinity, and could also be affected by any of several other factors. In future experiments, the
environmental conditions of the performance control must be held constant, as is specified
in Appendices C and D, in order that the background mortality, growth, and fertility may be
compared among experiments. More experience is needed with these two sediment toxicity
tests before absolute criteria can be set for performance control mortality, growth, or fertility
for either the 10-d or 28-d tests.
Reference Toxicant Controls: Juvenile K plumulosus were more sensitive than sub-adults to
Cd in the 96-h water-only exposures that comprised the reference toxicant control treatments
for the 28-d and 10-d sediment toxicity tests, respectively. In preliminary experiments with
subadult L/. plumulosus (i.e., sediment grain size sensitivity and cadmium sensitivity), the
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2-30
reference toxicant LCSOs ranged from 2.75-2.79 mg/L Cd. Salinity and temperature were
maintained at 28%c and 20-25°C in these two reference toxicant controls. For the rest of the
study, the 10-d sediment toxicity reference toxicant controls were conducted at 20%e and 25°C,
and the reference toxicant LCSOs for subadults dropped to 0.61-0.90 mg/L Cd. The reference
toxicant controls for the 28-d sediment toxicity test used newborns that were held for 7-d
under culture conditions prior to exposure to the 96-h water-only conditions (also 20%c and
25°C). The LC50 for these tests were 0.27 and 0.28 mg/L Cd. More reference toxicant control
exposures need to be conducted before quality control criteria can be established for either
the 10-d or 28-d L_. plumulosus sediment toxicity tests. However, these preliminary runs
provide initial guidance for the expected ranges of response of subsequent reference toxicant
control runs.
No reference toxicant control procedures have been developed for growth or fertility,
but this would be a useful avenue for further research.
Sensitivity of Leptocheirus plumulosus to Non-Contaminant Variables:
Sub-adults were tolerant of a broad variety of sediment types from fine sand to silty,
high TOC muds. They build burrows more readily in mud than in sand, and more
individuals may be seen out of their burrows in sandy sediments than in mud. Similar wide
tolerance for different sediment types was also seen for field-collected L. plumulosus
(Schlekat et al, 1992).
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2-31
This amphipod also has a broad salinity tolerance, ranging from 1.5%o to 32%&
(Schlekat et al., 1992). No experiments were conducted in this study to determine the
salinity tolerance of the cultured amphipods, but informal trials and other observations
confirm that adults and sub-adults tolerated salinities from <5%c to >35%e, and withstood
rapid changes in salinity without acclimation. This may not be true for newborns. A
posteriori analysis of the performance control data from all 10-d exposures indicated that
reduced growth may have been associated with high salinity (i.e., 28%o) (Fig. 2-10), although
high salinity was confounded by the absence of food (Table 2-19). It was also possible that
low salinity (i.e., ll%c) was responsible for reduced fertility in uncontaminated Corsica R.
sediment relative to fertility in the culture sediment performance control which had an
interstitial salinity of 20%c (Table 2-18). Clearly, further examination of the effect of salinity
on growth, fecundity, juvenile mortality, and contaminant sensitivity is needed.
Sub-adult amphipods tolerated 4 to 10-d periods without food or sediment (i.e., the
reference toxicant controls and unpubl. data) with very little mortality (i.e., <15%), but
newborns required food and/or sediment to survive even 4-d. If fed, sub-adults grew as much
as 35% in 10-d, but grew very little (i.e., <3%) if they were not fed. If fed, newborn L.
plumulosus doubled in size after 10-d and tripled in size after 28-d. Growth of sub-adults
and newborns in 10-d performance controls was possibly affected by the presence of food (Fig.
2-11), but since the presence of food was confounded with salinity, it was not possible to
distinguish which of these was more important in stimulating growth. However, based on
other experiments with food quantity and quality, we suspect that food was probably the
controlling variable. Size increased with the density of phytoplankton (i.e., food) provided,
and the dry food, gorp, also promoted rapid growth and high survival. However, gorp can
stimulate patches of bacterial mat growth on the sediment surface if it is not eaten, and for
-------
2-32
a
this reason, our laboratory continues to feed cultures and experimental amphipods with
mixture of gorp and cultured phytoplankton. More work should be directed to defining
simple-to-prepare, nutritious diet for L. plumulosus.
This species was also amenable to considerable handling during the course of an
exposure with no apparent effect on its concentration-sensitivity. Newborns exposed to a
wide concentration range of phenanthrene were sieved from the sediment in their exposure
chambers, counted, transferred to depression slides and measured under an microscope, and
returned to sediment without any change in their mortality-, growth- or fertility-
concentration relationship relative to amphipods that were not handled. Furthermore, it was
remarkable that amphipods as young as <24 h old could be used reliably to start sediment
toxicity tests with low control mortality. This tolerance of handling will help make L.
plumulosus a durable test organism.
Mortality as an Endpoint:
Mortality demonstrated consistent concentration-responsiveness in 4-d, 10-d and 28-d
exposures for a variety of chemical contaminants, including acenaphthene, phenanthrene,
cadmium, and a complex mixture of chemicals from a heavily polluted site in Chesapeake
Bay. The sensitivity of L,. plumulosus mortality to acenaphthene and phenanthrene
decreased as the organic content of the sediment increased, as has been seen for other PAHs
(Swartz et al., 1991) and unpolarized hydrophobic organic chemicals (DiToro et al., 1990).
Mortality was approximately as sensitive to contaminant concentrations as growth (Table 2-
20). As compared to fertility, mortality was less sensitive to phenanthrene but equally
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2-33
sensitive to Curtis Cr. (Baltimore Hbr.) sediment. Mortality might have been more sensitive
if variability within-treatments been lower or had more replicates been used. For example,
the average, among-treatment coefficient of variation for 28-d mortality in the Chesapeake
Bay dilution series experiment was 55.1%, which was approximately 12 times higher than
the mean CV of growth in that experiment.
The duration of the exposure did not greatly affect the sensitivity of mortality in two
experiments that compared 10-d and 28-d exposure periods (i.e., the phenanthrene-spiked
culture sediment and the Chesapeake Bay sediment dilution series experiments). If
anything, amphipod mortality in 10-d exposures was slightly more sensitive than mortality
of newborns in 28-d exposures. In the phenanthrene-spiked culture sediment experiment,
newborn amphipods were used for both exposure periods, and the survivors of the 10-d
exposure were returned to their respective experimental treatments for the remaining 18-d
for comparison with amphipods that were not handled during the 28-d exposure. The
phenanthrene sensitivity of the handled amphipods did not differ from that of the
unmanipulated amphipods after 28-d of exposure, and was not different from their
phenanthrene sensitivity after 10-d of exposure. Thus, it appears that the lethal toxicity of
phenanthrene was exerted within the first 10-d of exposure, and perhaps within a shorter
period of time. Mortality of sub-adults in 10-d exposures was more sensitive than newborn
mortality in 28-d exposures when exposed to a dilution-series of Baltimore Hbr. sediment.
However, the sub-adults in the 10-d exposure were not fed, whereas the newborns in the 28-d
exposure were fed. It seems likely that the sub-adults may have been stressed from
nutritional deficiency which may have rendered ~L. plumulosus more sensitive to chemical
contaminants, or the amphipods consumed more contaminated sediment when offered less
-------
2-34
food. As will be shown below for growth, the interaction between nutrition and contaminant
stress is clearly an important issue to examine in future research.
Size as an Endpoint:
Growth of Leptocheirus plumulosus decreased in a concentration-dependant manner
in response to PAHs and contaminated sediment from Baltimore Hbr. As with mortality, the
concentration of PAH necessary to reduce growth increased with the sediment TOC content,
as is predicted by equilibrium partitioning (DiToro et al., 1990). Body length (i.e., size) at the
termination of 10-d and 28-d exposures showed comparable sensitivity to chemical
concentration as mortality (Table 2-20), but was equally or less sensitive than fertility.
Within treatments, size was not highly variable, particularly in comparison with mortality
or fertility. Thus, increasing the number of replicates probably would not have led to an
appreciable increase in the concentration-sensitivity of growth. However, the "failure" of size
or growth to exceed mortality or fertility in sensitivity could be a reflection of the toxicants
used in these experiments (i.e., predominantly PAHs), and growth should continue to be
measured in exposures with other chemicals and field-collected sediments. Furthermore,
since males seem to grow faster than females (T.H. DeWitt and R. Singleton, AScI, unpubl.
data), some of the sensitivity of size may have been masked by differential growth rates.
Future research should examine whether the contaminant-sensitivity of growth is sex-
dependent, or whether differential growth rates of the two sexes masks the concentration-
response of growth for the two sexes combined.
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2-35
The chemical-sensitivity of size did not change consistently with the duration of
exposure to contaminants, and the differences in sensitivity that were observed may have
been due to nutritional deficiency. In the phenanthrene-spiked sediment experiment, size
after 10-d and 28-d exposures declined with PAH concentration (Table 2-14); however, due
to low statistical power (i.e., low replication), size was not significantly smaller at any
concentration relative to the carrier control. In the Chesapeake Bay sediment dilution
experiment, size at the 50% and 100% Curtis Cr. sediment treatments was significantly
reduced in the 28-d exposure, but not in the 10-d exposure (Table 2-18). Furthermore, size
did not decrease appreciably with concentration in the Chesapeake Bay 10-d exposure.
However, the sub-adults in this acute exposure were not fed and very little growth was
observed in either the carrier control (i.e., Corsica R. sediment) or the performance control
(i.e., culture sediment). Conversely, the newborn L. plumulosus in the 28-d exposure were
fed and size was reduced as the concentration of Curtis Cr. sediment increased. Since there
was little growth to begin with in the non-fed, 10-d exposure, there was little opportunity for
chemical contamination to reduce growth, unless it were to cause the amphipods to shrink,
which has been observed (unpubl. data). Thus, as with mortality, nutrition apparently
interacts with the concentration-response and/or sensitivity of size in both 10-d and 28-d
exposures.
Fertility as an Endpoint:
The fertility of female JL. plumulosus decreased in a concentration-responsive manner
in experimental exposures to phenanthrene-spiked culture sediment and a dilution series of
two Chesapeake Bay sediments. Fertility ranged from 10.3 to 0.08 female offspring per
-------
2-36
female survivor. The highest fertility was observed in a performance control (i.e.,
approximately culture conditions), and the lowest was observed in 100% Curtis Cr. sediment
from Baltimore Hbr. Fertility was more sensitive to phenanthrene-spiked sediment than
either mortality or growth, but equal in sensitivity to both of these endpoints after 28-d
exposure to the Chesapeake Bay sediment dilution series. Fertility was highly variable
within treatments, relative to size: for the Chesapeake Bay sediment dilution experiment, the
average, within treatment CV was 132% (or 36% if the 100% treatment is excluded) (Table
2-18). The statistical power of fertility as a sub-lethal response would clearly increase if more
than 3 replicates were used per treatment.
Fertility was sensitive to one or more uncontrolled factors. In the Chesapeake Bay
sediment dilution experiment, fertility in the site control (i.e., the 0% Curtis Cr. treatment
= 100% Corsica R. sediment) was half that of the performance control. Both treatments
received food equally. The Corsica R. sediment was collected from a site sustaining a year-
round population of L. plumulosus (B. McGee, MD Dept. Environment, pers. comm.).
Although metal and PAH contaminants were low in this sediment, it is possible that
unmeasured chemicals were present that inhibited reproduction. Alternatively, fertility may
have responded to the different interstitial salinities present in the two sediments (i.e., ll%c
in the Corsica R. sediment and 20%o in the South Beach, OR, sediment). The sensitivity of
fertility to salinity is uncertain: Schlekat et al. (1992) found reduced reproductive production
at 5-15%o relative to 25-32%c after 20-d exposure under non-contaminant conditions, but the
difference was not significant after 28-d of exposure. The effect that food quantity or quality
has on JL plumulosus fertility could not be determined since the amphipods were fed in all
of the 28-d experiments. However, the feeding regime should be expected to affect the
magnitude and possible contaminant-sensitivity of fertility as it apparently does mortality
-------
2-37
and growth. Clearly, the effects of non-contaminant environmental variables and food on
reproduction of L,. plumulosus must be high priority for further research in the development
of a chronic sediment toxicity test protocol with this amphipod.
Finally, the mechanism(s) responsible for reduced fertility need to be determined. As
measured here, fertility was the number of juvenile females produced per surviving female.
Reduction of fertility could have been caused by reduced egg production, death of embryos
held in the maternal brood pouch, or death and decomposition of offspring shortly after
release from the brood pouch. Examination of the number and condition of eggs and embryos
in the brood chambers of females under contaminant-stress and control conditions should the
necessary information.
Sex Ratio as an Endpoint:
The sex ratio of surviving L. plumulosus was not affected by the concentration of
chemical contaminants in either experiment in which it was measured. Thus, it is of no
direct utility as a toxicological endpoint. However, the sex ratio of the surviving members
of the F0-cohort must be measured in every test so that fertility may be accurately
determined. Surprisingly, the sex ratio of L,. plumulosus was not 1.0 as expected, but the
cohorts were comprised of 60-70% females on average (Fig. 2-12). While this difference in
relative abundance of the two sexes appeared in two experiments, possible alternative
hypotheses to explain this sex ratio are 1) females were unconsciously favored in the
supposed random picking of newborns during the setup of the experiment, 2) females left the
maternal brood pouch slightly before males did and were therefore over represented in the
-------
2-38
pool of
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2-39
practical, applied issues are 1) development of reference toxicant control methods for growth
and fertility, 2) establishing the ranges of response of mortality, growth, and fertility under
performance and reference toxicant control conditions, 3) determining the sensitivities of
growth and fertility to natural environmental variables such as sediment grain size, organic
content, and salinity, and 4) determining the sensitivity of all response endpoints to other
classes of chemicals in addition to the PAHs tested in this study. The more fundamental
issues include 1) determining the influence of nutrition on the toxicological sensitivity of L.
plumulosus and methods to either measure or control the nutritional condition of the
bioassay organisms, 2) comparing the relative toxicological sensitivity of L,. plumulosus to
other marine and estuarine species, 3) measuring the relative sensitivities of cultured and
field-collected L. plumulosus, and 4) establishing the ecological significance of the acute and
chronic responses. Some of the practical issues will be resolved as this species is applied in
research and regulatory sediment toxicity tests. The fundamental research questions and the
development of reference toxicant control methods for growth and fertility will require a more
concerted research effort.
The dominant issue to be resolved is the influence of nutrition on the range of
response for mortality, growth, and fertility under controlled conditions, and the interaction
between nutrition and chemical concentration on these response variables. If L. plumulosus
was not fed during a 10-d exposure, it hardly grew. Had the amphipods not been fed in the
28-d exposures, it is likely that growth, fertility, and possibly mortality would also have been
affected. The nutritional quality of sediments is not of great environmental concern,
especially relative to the impact of chemical contaminants, but sediment toxicity tests with
unfed L,. plumulosus might be unable to discriminate between sediments of low nutritional
value and high chemical contamination. However, if the amphipods were to be fed ad
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2-40
libidum, they might reduce their exposure to chemical contamination by preferentially
ingesting nutritious food over polluted sediment, or avoidance of the latter. Such selectivity
has been demonstrated for several benthic deposit feeders, including aorid amphipods
(DeWitt, 1987), and is an important consideration in the determination of bioaccumulation
rates in deposit-feeding bivalves (Lee et al., 1990). There is no obvious solution to this
dilemma short of independently determining the nutritional quality of the sediment or the
nutritional health of the amphipods. This problem is hardly unique to JL plumulosus as it
plagues all bioassay organisms, especially those for which the qualitative and quantitative
nature of the food are unknown. Considerable effort has been expended trying to identify and
quantify the food of deposit-feeding invertebrates (Lopez and Levinton, 1987; Lopez et al.,
1989), and it is not yet possible to independently determine the nutritional quality of
sediments except by the response of a bioassay organism. Furthermore, the nutritional
quality and quantity of the diet has been shown to affect the sensitivity of mysids, another
peracarid crustacean, to zinc (P.M. Vance, OR State Univ., unpubl. data). The influence of
nutrition on bioassay responses has been a latent problem that is now emerging as an
important research issue for chronic water and sediment toxicity tests.
2.5 CONCLUSION
The capacity of the estuarine amphipod, Leptocheirus plumulosus, for use in acute
(i.e., 10-d exposure) and chronic (i.e., 28-d exposure) sediment toxicity tests was
demonstrated. New acute and chronic toxicity-test methodologies developed from the
experiments reported here, and are similar to those described in the ASTM guidelines for
-------
2-41
amphipod sediment toxicity tests (ASTM, 1990b). Laboratory-cultured L. plumulosus were
used in this study, but other researchers have successfully conducted sediment toxicity tests
with field collected animals. Mortality and growth had comparable concentration-sensitivity
for both 10-d and 28-d exposure periods, while fertility was more sensitive than mortality or
growth in one 28-d exposure, but only equally sensitive in another. These responses were
observed in laboratory experiments with chemically-spiked sediments and with a dilution-
series of a highly contaminated sediment from Baltimore Hbr. mixed with an uncontaminated
sediment from the eastern shore of Chesapeake Bay. The 10-d sediment toxicity test was
effective across a wide range of sediment grain sizes and organic contents, and neither
mortality or size were significantly correlated with any sediment parameter. ]L. plumulosus
had a very wide salinity tolerance and probably could be used to test sediments with
interstitial water salinities ranging from ca. 2%c to 30%o, although the effect of salinity on
chronic sensitivity to contaminants has not been examined. The concentration-sensitivity of
L_. plumulosus varied with the bioavailability of the toxicant, as was seen for cadmium in
water of different salinities, and both for acenaphthene and phenanthrene spiked into
sediments with different organic carbon contents. This study provides the bases for
developing acute and chronic sediment toxicity test protocols with L. plumulosus. Several
important issues remain to be resolved, including the determination of the ranges of
responses under control conditions, sensitivities to different contaminants, development of
reference toxicant controls for growth and fertility, and the influence of nutrition on the
sensitivity of L_. plumulosus to contaminated sediments. Experience gained through use of
the new sediment toxicity test methods developed herein will provide much of the necessary
information on the expected ranges of responses under both performance and reference
toxicant control conditions. Future research should focus on the remaining problems, which
are, in order of priority: 1) the influence of non-contaminant variables on toxicological
-------
2-42
sensitivity, particularly nutrition (e.g., quality and quantity), but also temperature, salinity,
sediment grain size, and perhaps ammonia and hydrogen sulfide; 2) development of reference
toxicant method(s) for growth and fertility; 3) determination of the relative sensitivity of
cultured and field-collected animals to chemical contaminants in sediment; 4) simplification
of the culture and feeding methods; 5) comparison of the relative sensitivity of L. plumulosus
to other species (acute and chronic); 6) development of a toxicological database, including both
pure compounds in spiked-sediment exposures and field-collected contaminated sediments;
7) conducting an inter-laboratory comparison study to determine inter-laboratory variability
in toxicity test responses; and 8) conducting field validation studies to determine whether the
methods are predictive of benthic population, community, habitat, or ecosystem responses to
chemical contamination.
-------
2-43
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2-51
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h o
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100
Figure 2-9. Mortality, size, and fertility of _L. plumulosus after 10-d and 28-d exposures to
a dilution series of Curtis Cr. sediment (Baltimore Hbr., MD) mixed with uncontaminated
Corsica R. sediment. The "Control" treatments were the performance controls.
-------
2-52
15
tio
CO
C
o
Newborn
Newborn
SubAdult SubAdult
SubAdult
Newborn SubAdult
20 20 20 20 28 28 28
Newborn
2.5
i
0.5
0
Newborn
Newborn
SubAdult
SubAdult
SubAdult
20
20
20 20 28
Salinity (ppt)
28
Figure 2-10. Influence of salinity on mortality and growth (mean ± SD) of L. plumulosus in
the performance controls of 10-d sediment toxicity tests. The age class of amphipods used
in each experiment is labeled above each bar.
-------
2-53
15
CO
r
o
0
Newborn
Newborn
SubAdult SubAdult
SubAdult
SubAdult
Newborn
No
No
No
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Newborn
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Newborn
SubAdult
O
5
0.5
0
SubAdult SubAdult
SubAdult
1
No No No Yes Yes
Food Added
Yes
Yes
Figure 2-11. Influence of feeding on mortality and growth (mean ± SD) of L. plumulosus in
the performance controls of 10-d sediment toxicity tests. The age class of amphipods used
in each experiment is labeled above each bar.
-------
2-54
.-— N
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o o
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I °
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o
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o
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I
' 0
!_8 °..__€) .0 o -
> O ^"^
o
>
1 1 1 1 1
0
0
20 40 60 80
% Curtis Cr. Sediment
100
Figure 2-12. Sex ratio (#females:#males) of surviving regeneration L,. plumulosus plotted
as a function of contaminant-concentration in (A) culture sediment spiked with phenanthrene,
and (B) a dilution series of two Chesapeake Bay sediments. The "Control" treatments were
the performance controls.
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2-56
1
Diameter
% Fines
% TOG
% Water
.Eh
Mortality
-0.01
0.02
-0.11
-0.15
0.17
Table 2-2. Product-moment correlation coefficients ("r") for mortality of Leptocheirus plumulosus sub-
adults and sediment variables. Data obtained from 12 estuarine sediments from Oregon. None of the
coefficients were statistically significant (i.e., p>0.05).
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-------
2-58
Variables
%Mortality
Amphipod Size
Median Particle
Diameter
%Fines
%Water
%TOC
%Mortality
1.00
-0.61*
0.25
-0.31
-0.29
-0.12
Size
-0.61*
1.00
-0.19
0.17
0.19
0.14
Table 2-4. Pearson product-moment correlations between sediment variables and mortality
and size of Ij. plumulosus newborns;. * = p<0.05.
-------
2-59
Feeding
Treatment
No Food
104 Algal cells/ml
105 Algal cells/ml
106 Algal cells/ml
Gorp Only
%Moi
Mean
26.7A
20.0B
6.7B
11.7AB
6.7B
-tality
SD
7.6
5.0
2.9
2.9
11.5
Size
Mean
2.17A
2.30A
2.87B
4.50D
3.49C
(mm)
SD
0.16
0.20
0.14
0.13
0.06
Table 2-5. Effect of feeding regime on mortality and size of newborn Leptocheirus plumulosus.
Differences in mortality and growth among the treatments were tested with anova and
Tukey's multiple-comparisons t-test; treatment means that were not significantly different
were labeled with the same letter. Mean size at T0 = 2.05 mm.
-------
2-60
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-------
2-61
Cd Cone
(ni{
Nominal
0.0
0.19
0.38
0.75
1.5
3.0
6.0
entration
?/L)
Measured
0.0
0.21
0.45
0.87
1.69
3.47
6.46
%Mo
Rep. 1
10
40
65
100
100
100
100
rtality
Rep. 2
0
45
70
60
100
100
100
LC50NOM1NA1 = 0.25 mg/L Cd (0.16-0.39 = 95% CI)
LC50MEASUKED = 0.28 mg/L Cd (0.18-0.46 = 95% CI)
Table 2-7. Acute mortality of newborn Leptocheirus plumulosus after 96-h static
exposure to Cd in seawater (20%c salinity, 25°C).
-------
2-62
Total Sec
Acenaph
(mg/dr
Nominal
250B
150B
90A
54A
32A
19A
12A
?A
0A
0B
Performance
Control
Performance
Control8
TA
0
rp B
-••o
liment
thene
y kg)
Measured
192.8
121.1
57.5
45.8
22.6
15.8
7.5
3.6
0.0
0.0
0.0
0.0
—
%Mor
Mean
20.0*
2.5
7.5
0.0
2.5
0.0
2.5
0.0
0.0
0.0
7.0
5.0
—
tality
SD
5.7
3.5
10.6
0.0
3.5
0.0
3.5
0.0
0.0
0.0
5.7
6.1
—
Size(
Mean
3.63*
3.74*
4.21
4.26
4.35
4.42
4.60
4.42
4.57
4.30
4.57
4.89
3.37
4.73
mm)
SD
0.11
0.18
0.30
0.04
0.12
0.19
0.04
0.18
0.04
0.09
0.22
0.15
0.59
0.19
Table 2-8. Effects of acenaphthene-spiked South Beach "Old Log" sediment on
mortality and growth of sub-adult Leptocheirus plumulosus after 10-d exposure.
Differences in mortality and size among the treatments, relative to the O mg/kg
treatment, were tested with anova and Dunnett's multiple-comparisons t-test for
exposures conducted in February 1991 (= A) or June 1991 (= B); * = p<0.05. N=2,
except for Performance Controls and T0A. LC50 = >192.8 mg/dry kg.
-------
2-63
Total Se<
Acenaplr
(mg/dr
Nominal
416B
250A
150A
90A
54A
32A
19A
0A
0B
Performance
Control
Performance
Control8
TA
0
rrt B
-"-o
iiment
ithene
P kg)
Measured
422.5
209.7
124.1
67.5
42.2
24.3
14.3
0
0
0
0
_
%Mor
Mean
75.0*
60.0*
2.5
2.5
7.5
2.5
0.0
2.5
0.0
7.0
5.0
—
•tality
SD
7.1
7.1
3.5
3.5
3.5
3.5
0.0
3.5
0.0
5.7
6.1
—
Size (
Mean
2.90*
3.83
4.14
4.40
4.38
4.48
4.47
4.31
6.58
4.57
4.89
3.37
4.73
'mm)
SD
0.09
0.56
0.47
0.08
0.40
0.30
0.11
0.41
0.05
0.22
0.15
0.59
0.19
Table 2-9. Effects of acenaphthene-spiked McRinney Slough sediment on mortality
and growth of sub-adult Leptocheirus plumulosus after 10-d exposure. Differences
in mortality and size among the treatments, relative to the O mg/kg treatment, were
tested with anova and Dunnett's multiple-comparisons t-test for exposures conducted
in February 1991 (= A) or June 1991 (= B); * = p<0.05. N=2, except for Performance
Controls and T0A for which N=5. LC50 = 209.3 mg/dry kg (95% CI: 166.7-262.8
mg/dry kg).
-------
2-64
Total Sec
Acenaph
(mg/dr
Nominal
693B
416A
250A
150A
90A
54A
32A
0A
0B
Performance
Control
Performance
Control6
TA
0
TB
0
liment
tthene
7 kg)
Measured
501.4
356.2
206.4
121.0
63.3
39.7
23.9
0
0
0
0
_
%Mor
Mean
77.5*
45.0*
15.0
15.0
0.0
5.0
15.0
7.5
0.0
7.0
5.0
—
tality
SD
17.7
21.2
7.1
7.1
0.0
0.0
7.1
3.5
0.0
5.7
6.1
_
Size(
Mean
5.41*
3.89
4.02
4.22
4.27
4.48
4.51
4.42
6.30
4.57
4.89
3.37
4.73
mm)
SD
0.14
0.50
0.57
0.26
0.10
0.04
0.06
0.06
0.14
0.22
0.15
0.59
0.19
Table 2-10. Effects of acenaphthene • spiked Eckman Slough sediment on mortality and
growth of sub-adult Leptocheirus plumulosus after 10-d exposure. Differences in
mortality and size among the treatments, relative to the O mg/kg treatment, were
tested with anova and Dunnett's multiple-comparisons t-test for exposures conducted
in February 1991 (= A) or June 1991 (= B); * = p<0.05. LC50 = 373.0 mg/dry kg (95%
CI: 321.2-433.1 mg/dry kg).
-------
2-65
Total Se<
Phenanl
(mg/dr
Nominal
150B
90A
54A
32A
19A
12A
?A
0A
0B
Performance
Control
Performance
Control8
rp A
-"-o
m B
io
iiment
,hrene
y kg)
Measured
107.7
70.6
48.8
27.0
19.1
11.0
6.4
0
0
0
0
_
%Mor
Mean
60.0
35.0*
5.0
0.0
5.0
7.5
2.5
2.5
0.0
8.0
5.0
—
tality
SD
56.7
7.1
7.1
0.0
0.0
3.5
3.5
3.5
0.0
4.5
6.1
_
Size(
Mean
3.09
4.14*
4.45*
4.51*
4.67*
5.09
5.30
5.34
4.30
3.82
4.89
3.74
4.73
mm)
SD
0.00
0.06
0.19
0.12
0.13
0.01
0.23
0.05
0.08
0.38
0.15
0.06
0.19
Table 2-11. Effects of phenanthrene-spiked South Beach "Old Log" sediment on
mortality and growth of sub-adult Leptocheirus plumulosus after 10-d exposure.
Differences in mortality and size among the treatments, relative to the 0 mg/kg
treatment, were tested with anova and Dunnett's multiple-comparisons t-test for
exposures conducted in May 1991 (= A) or June 1991 (=B); * = p<0.05. LC50 = 91.9
(76.2-110.8 = 95% CD mg/dry kg.
-------
2-66
Total Sec
Phenant
(mg/dr
Nominal
416B
250A
150A
90A
54A
32A
19A
0A
0B
Performance
Control
Performance
Control8
rp A
•'•o
rp B
-"-o
iiment
hrene
P kg)
Measured
270.0
173.2
120.0
77.1
49.4
30.3
19.0
0
0
0
0
._
%Mor
Mean
80.0*
60.0*
10.0
5.0
7.5
15.0
5.0
2.5
0.0
8.0
5.0
—
tality
SD
7.1
0.0
7.1
7.1
3.5
0.0
7.1
3.5
0.0
4.5
6.1
—
Size(
Mean
2.99*
2.37
4.07
3.95
4.12
3.79
3.92
3.97
6.58
3.82
4.89
3.74
4.73
mm)
SD
0.44
0.31
0.20
0.01
0.17
0.00
0.05
0.21
0.05
0.38
0.15
0.06
0.19
Table 2-12. Effects of phenanthrene-spiked McKinney Slough sediment on mortality
and growth of sub-adult Leptocheirus plumulosus after 10-d exposure. Differences
in mortality and size among the treatments, relative to the O mg/kg treatment, were
tested with anova and Dunnett's multiple-comparisons t-test for exposures conducted
in May 1991 (= A) or June 1991 (=B); * = p<0.05. LC50 = 170.1 (150.9-191.6 =
95%CI) mg/dry kg.
-------
2-67
Total Sei
Phenanl
(mg/dr
Nominal
693B
416B
416A
250A
150A
90A
54A
32A
0A
0B
Performance
Control
Performance
Control8
rp A
-"-o
TB
0
iiment
;hrene
ykg)
Measured
346.4
273.2
76.6
174.9
105.0
61.0
44.1
27.5
0
0
0
0
—
%Mot
Mean
80.0*
52.5*
12.5
22.5
17.5
17.5
17.5
12.5
7.5
0.0
7.0
5.0
—
•tality
SD
7.1
3.5
10.6
10.6
3.5
10.6
3.5
10.6
3.5
0.0
4.5
6.1
—
Size(
Mean
4.65*
5.36*
4.07
3.93
3.74
4.04
4.04
3.89
4.18
6.31
3.82
4.89
3.74
4.73
mm)
SD
0.07
0.12
0.11
0.19
0.19
0.04
0.16
0.01
0.06
0.13
0.38
0.15
0.06
0.19
Table 2-13. Effects of phenanthrene-spiked Eckman Slough sediment on mortality
and growth of sub-adult Leptocheirus plumulosus after 10-d exposure. Differences
in mortality and size among the treatments, relative to the 0 mg/kg treatment, were
tested with anova and Dunnett's multiple-comparisons t-test for exposures conducted
in May 1991 (= A) or June 1991 (=B); * = p<0.05. LC50 = 254.8 (229.3-283.1 =
95%CI) mg/dry kg.
-------
2-68
Exposure
Duration
(days)
10
10
10
10
10
10
10
10
10
10
28
28
28
28
28
28
28
28
28
28
Total Sec
Phenant
(mg/dr
Nominal
Performance
Control
0
25
35
50
72
103
147
210
300
Performance
Control
0
25
35
50
72
103
147
210
300
liment
hrene
ykg)
Measured
0.00
0.00
20.26
26.34
41.54
51.62
73.32
105.47
141.2:5
183.97
0.00
0.00
20.26
26.34
41.54
51.62
73.32
105.47
141.25
183.97
%Mortality
0.0
5.0
5.0
0.0
0.0
5.0
5.0
0.0
45.0
55.0
8.3
(6.8)
1.7
(2.9)
11.7
(7.6)
1.7
(2.9)
3.3
(5.8)
8.3
(5.8)
3.3
(2.9)
21.7
(22.5)
18.3
(23.6)
60.0*
(5.0)
Size1
(mm)
3.42
3.58
-
-
3.33
-
3.14
3.27
-
2.80
7.60
(0.27)
7.41
(0.05)
7.57
(0.19)
7.41
(0.31)
7.24
(0.44)
7.40
(0.25)
7.51
(0.56)
7.36
(0.06)
7.30
(0.35)
7.07
(0.55)
Fertility
-
-
-
-
-
-
-
-
-
-
9.13
(0.51)
9.68
(1.52)
5.96*
(0.18)
6.44*
(1.54)
6.25*
(1.20)
6.99*
(1.09)
4.42*
(0.45)
4.20*
(1.78)
4.50*
(0.99)
5.36*
(1.83)
Sex Ratio
(P:M)
-
-
-
-
-
-
-
-
-
-
1.05
1.61
0.94
1.61
1.43
1.63
1.95
1.05
1.71
1.42
Table 2-14. Mean (SD) responses of newborn Leptocheirus plumulosus to phenanthrene-
spiked culture sediment after 10 and 28 d of exposure. Statistical significance of responses
among the treatments, relative to the O mg/kg treatment, was tested with anova and
Dunnett's multiple-comparisons t-test; * = p<0.05. N=l for 10-d exposures and N=3 for 28-d
exposures, except for 28-d Performance Control (N=6) and size in the 147 mg/dry kg
treatment (N=2). Size at T0= 1.75 (0.16 SD) mm. Animals were not measured in some of the
acute-exposure treatments because of time constraints.
-------
2-69
Sediment Variable
Median Diameter (urn)
% Gravel
% Sand
% Silt
% Clay
% Water
%TOC
0
4.2
0.0
2.6
49.8
47.6
67.7
1.82
6.25
6.0
1.8
4.9
49.9
43.5
67.7
2.39
% Curtis Cre
12.5
4.6
0.6
7.0
46.3
46.1
66.8
2.66
ek Sediment
25
5.0
2.1
10.0
44.1
43.9
66.6
2.54
50
8.1
1.6
17.0
37.1
44.3
68.0
3.58
100
21.1
6.1
33.2
22.6
38.1
67.6
4.23
Table 2-15. Grain size analysis, water content, and TOC content for six substrates from a dilution series of
sediment from Curtis Cr. (Baltimore Hbr., MD) and Corsica R., MD.
-------
2-70
Compound
(ug/dry kg)
Naphthalene
2-Methyl naphthalene
1-Methyl naphthalene
Biphenyl
2,6 Methyl naphthalene
2,3,5 Dimethyl naphthalene
Acenaphthylene
Acenaphthene
Fluorene
Phenanthrene
Anthracene
Methyl phenanthrene
Fluoranthene
Pyrene
Benz(a)anthracene
Chrysene
Benzo(b)fluoranthene
Benzo(k)fluoranthene
Benzo(e)pyrene
Benzo(a)pyrene
Perylene
Benzo(ghi)perylene
0
28
89
bdl
bdl
bdl
bdl
bdl
bdl
bdl
77
bdl
bdl
104
105
85
110
93
bdl
bdl
58
1L
bdl
6.25
137
271
311
108
170
90
94
1080
892
1950
398
256
3620
2420
1240
1380
570
bdl
444
599
136
257
% Curtis Cre
12.5
149
213
348
139
162
119
bdl
1480
1480
2710
589
245
4830
3070
1130
978
569
bdl
377
454
97
228
ek Sediment
25
196
375
394
220
300
255
214
2440
2940
9220
2200
751
14300
9540
4270
4500
2740
2000
1330
1740
377
704
50
935
1120
1230
737
822
799
523
8500
10300
50000
6010
2640
30700
22400
10800
12200
7690
5100
3880
4940
1360
2290
100
1170
1980
2580
1640
1590
1450
1000
21300
14100
71800
14300
5830
71800
46600
23800
28500
20200
12200
9410
12800
4360
5640
Table 2-16. Total-sediment concentrations of selected PAHs measured in a six substrates from a dilution series
of sediment from Curtis Cr. (Baltimore Hbr., MD) and Corsica R., MD. bdl = below detection limits.
-------
2-71
Compound
AVS (umole/dry g)
Cd (fig/dry g)
Cu (ng/dry g)
Ni (ug/dry g)
Pb (ug/dry g)
Zn (ug/dry g)
Total SEM (pmole/dry g)
Total SEM/AVS
0
35.8
0.18
25.0
46.4
77
176
4.25
0.12
6.25
20.5
1.15
26.9
29.4
63
109
2.91
0.14
% Curtis Cre
12.5
42.2
0.23
38.6
25.1
114
153
3.92
0.09
ek Sediment
25
72
0.30
87.6
56.8
196
263
7.31
0.11
50
86.3
0.97
148
30.1
243
419
10.43
0.12
100
491.3
2.95
368.9
89.2
817
1264
30.62
0.06
Table 2-17. Acid-volatile sulfides (AVS) and simultaneously extracted metals (SEM) measured in six substrates
from a dilution series of sediment from Curtis Cr. (Baltimore Hbr., MD) and Corsica R., MD.
-------
2-72
Exposure
Duration
10
10
10
10
10
10
10
Treatment
(% Curtis Cr.)
Performance
Control
0%
6.25%
12.5%
25%
50%
100%
%Mortality
0.0
(0.0)
1.7
(2.9)
15.0
(5.0)
0.0
(0.0)
21.7
(10.4)
40.0*
(18.0)
65.0*
(15.0)
Size
(mm)
3.33
(0.11)
3.09
(0.11)
3.28
(0.07)
3.26
(0.13)
3.21
(0.20)
3.23
(0.13)
3.14
(0.29)
Fertility
_
_
.
_
_
_
_
Sex Ratio
_
_
_
_
_
.
.
28
28
28
28
28
28
28
Performance
Control
0%
6.25%
12.5%
25%
50%
100%
12.0
(12.5)
9.2
(3.8)
10.0
(5.0)
11.7
(7.6)
16.7
(5.8)
25.0
(21.8)
86.7*
(2.9)
6.16
(0.11)
6.53
(0.67)
6.90
(0.10)
6.23
(0.15)
6.30
(0.17)
5.60*
(0.26)
5.17*
(0.40)
10.32
(1-71)
5.20
(1.57)
5.53
(3.52)
6.97
(2.08)
4.70
(0.60)
3.13
(1.99)
0.10*
(0.71)
1.87
(0.76)
0.91
(0.42)
1.73
(0.94)
1.72
(1.13)
1.15
(0.46)
2.38
(1.47)
1.50
(0.71)
Table 2-18. Responses of L. plumulosus to a Chesapeake Bay sediment dilution series after
10-d and 28-d exposure periods. Statistical significance of responses among the treatments,
relative to the O% treatment, was tested with anova and Dunnett's multiple-comparisons t-
test; * = p<0.05. N=3 for all treatments except Performance Controls (N=5). Size at T0= 2.94
(0.19 SD) mm for 10-d exposures, and 1.83 (0.08 SD) for 28-d exposures.
-------
2-73
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-------
CHAPTER III
DEVELOPMENT OF A CHRONIC SEDIMENT BIOASSAY WITH
AMPELISCA ABDITA
3.1 INTRODUCTION
Research with Ampelisca abdita sought to develop culture methods and a chronic
bioassay for this species. Bioassay development built on the research of Scott and Redmond
(1989), who showed that A., abdita could be used to test chronic and population endpoints.
Culturing methods and results are described in Chapter I above. The approach to
chronic test development was to 1) establish cultures, 2) estimate optimum temperature and
salinity regimes, 3) outline a proposed chronic test design, 4) evaluate the chronic test design
with uncontaminated sediment, and 5) evaluate the chronic test design with contaminated
sediment. The experiments we conducted addressed points 2-4. Both cultures and the
controlled experiments described in this section utilized amphipods from a Narragansett, RI,
source population; details of collection and handling were presented in Chapter 1.
3-1
-------
3-2
3.2 MATERIALS AND METHODS
General Methods
An overview of the six experiments conducted with A. abdita is shown in Table 3.1.
With the exception of #1 (Temperature and Salinity Effects), all experiments were conducted
at approximately 30%e salinity. In all experiments, the photoperiod was maintained at 16 hr
light: 8 hr dark. All experiments used uncontaminated sediment collected from Yaquina Bay
(OR) tide flats sieved through a 0.5mm or 0.25mm screen. Grain size analyses were
conducted on the processed sediments. In Experiment #5 (Sediment- and Animal-Source
Effects), an additional uncontaminated sediment was tested. In all but Experiment #6
(Container, Aeration, and Nutrition Effects), testing was conducted in temperature-controlled
water baths.
Unless otherwise noted, the following daily observations were made: temperature;
salinity; numbers, sex, and life stage of amphipods caught on the water surface tension;
number of amphipods on the sediment surface; number and sex of dead amphipods. Dead
amphipods were removed daily except where noted.
At the conclusion of each experiment, the contents of each exposure container were
sieved through a 0.25mm screen and the recovered amphipods counted and sexed. Two types
of survival calculations were made: 1) percent survival and 2) percent survival corrected for
senescence mortality. A. abdita males die shortly after mating, and females die at some
indeterminate time after completing their reproductive cycle. Males and spent females were
easily recognized when dead individuals were examined under a dissecting microscope.
-------
3-3
Treatment effects may be more easily detected by counting these individuals as live in the
survival calculations, thus correcting for mortality due to senescence.
In Experiments #2, 3, and 5 (Life Cycle at 25°C, Temperature and Nutrition Effects,
and Sediment and Animal Source Effects, respectively), recovered amphipods were preserved
in 70% ethanol with glycerin for later length measurements. Amphipods from Experiments
#2 and 3 were measured with an ocular micrometer and those from Experiment #5 with an
image analysis system. Length was measured along the dorsal surface, from the base of the
first antennae to the base of the telson.
Experiment #1: Temperature and Salinity Effects
The goal of this experiment was to estimate optimum temperature and salinity for A.
abdita culture and testing. Thirty adult amphipods from the initial field collection (Jan. 1990
- see Chapter I) were placed in sediment in each of 12 one-gallon jars. There were three jars
in each of the following treatments: 20°C, 20%0; 20°C, 30%0; 25°C, 20%0; and 25°C, 30%c. The
amphipods were acclimated gradually to the test temperatures and salinities. Three times
per week, approximately 75% of the overlying water in each jar was siphoned out for renewal
and replaced with 500-1000 ml of a salinity-adjusted suspension of the alga Pseudoisochrysis
paradoxa and seawater of the appropriate salinity and temperature. Algal density was not
measured, but undiluted algal cultures ranged between 106-107 cells/ml.
During the eighth week (days 52-56), the contents of the first jar in each treatment
were sieved and the-material remaining on the sieve examined, with the amphipods still
-------
3-4
alive. Contents of the second jar in each treatment were sieved on day 66 and preserved for
later examination, and contents of the third were examined live during week 10 (70-73 days).
The results were compared non-statistically, since there were no true replicates (jars of the
same temperature/salinity treatment sampled at the same time). Those amphipods not
sacrificed were returned to the cultures.
Experiment #2: Life Cycle at 25°C
A draft chronic sediment toxicity test design for A. abdita (Appendix E) was tested in
this experiment. Another goal of this experiment was to define the life cycle of A. abdita at
25°C. Methods essentially followed the chronic test design except: 1) all exposure chambers
were under control conditions; treatments were predefined sampling times, 2) there were only
3 replicates per sampling time, and 3) the last replicates were sieved after 49 days. The 20
juveniles (8-10 days old) used to initiate the experiment were obtained from cultured females.
A flowing seawater delivery system was set up to supply one volume replacement of an algae-
seawater mixture per day per jar. The mixture was prepared so as to supply 100 ml algal
culture plus 500 ml filtered seawater in each jar's volume replacement. The proportion of
algae to seawater in the overlying water was the same for these exposure chambers as it was
in the chambers used in the previous experiment. The algal density was not measured, but
the undiluted culture density ranged between 106-107 cells/ml. Results were examined
graphically.
-------
3-5
Experiment #3: Temperature and Nutrition Effects
This experiment examined the effects of temperature and food type on reproduction.
Two temperatures were compared, 20°C and 25°C. Feeding treatments were: no food
controls; Pseudoisochrysis paradoxa; an algal mixture of P. paradoxa. Phaeodactylum
tricornutum. and Chaetoceros calcitrans (1:1:1 by volume; se Table 3-2 for algal density); and
the algal mixture plus approximately 4 mg of finely ground (<125um) Neo-Novum® pellets
sprinkled onto the water surface daily. Static exposure with daily renewals was used because
of the variety of feeding and temperature regimes necessary. No water was siphoned out;
seawater-food mixtures (100 ml algal culture mixture and 500 ml seawater per jar) were
added with a funnel system that delivered water into jars approximately 3/4 of the way down
in the water column and the displaced water exited the chamber through an screened
overflow port. Disturbance of the sediment surface was prevented by the use of a plastic "T"
at the end of the delivery tube.
The test was initiated with newly-released juveniles, to examine the feasibility of
using that life stage to start a test. The juveniles were released in seawater only from
brooding females that had been collected in Narragansett, RI, shipped to Newport, Oregon,
and held in the gallon jar culture system with feeding until their broods were near release.
A staggered start, adding juveniles to replicate jars, 10/replicate, as they were released, was
used due to the limited number of young produced on any given day. Cell counts and
volumes of algal cultures used, and weight of dry food material added were measured daily.
Twenty replicates were sieved after 14 days to count, preserve, and measure length of
survivors, and the remaining 18 were sieved, preserved, and examined after 41 days. Five
samples of amp bipods taken on day zero (<1 day old) were also preserved and measured.
-------
3-6
Lengths of 15-day-old amphipods were compared with those of initial animals using
t-tests. T-tests were also used to compare the lengths of amphipods from identical food
treatments at different temperatures. Within each temperature treatment, an analysis of
variance followed by Tukey's Studentized Range Test was used to determine differences
between feeding treatments.
Experiment #4: Density Effects
This experiment examined the potential effects of amphipod density on reproduction
at 20°C. There were three replicates each at 10, 20, and 40 amphipods per jar. The
experiment was initiated with juveniles 8-10 days old, as in Experiment #2 (Life Cycle at
25°C), and was terminated after 38 days. Newly released juveniles were collected in
seawater only from females which were obtained from the field in RI, shipped to Oregon, and
held under culture conditions until they carried broods. All treatments were fed the 1:1:1
algal mixture with the quart jar renewal system as described for Experiment #3
(Temperature and Nutrition Effects), with a mixture of 100 ml algal culture and 500 ml
seawater added daily per jar, plus approximately 5 mg ground Neo-Novum® daily. Fifty
milligrams of ground Neo-Novum® were stirred with 10 ml of seawater, and 1 ml of the slurry
was added to each replicate. No statistical analyses were performed.
-------
3-7
Experiment #5: Sediment and Animal Source Effects
This experiment examined the hypotheses that 1) Yaquina Bay sediment might be
sublethally toxic to A. abdita, and 2) offspring of cultured animals might show poorer survival
and reproduction than those of field-collected animals. Two sediments were tested: the
Yaquina Bay culture sediment, and sediment from central Long Island Sound. The latter,
which has been and continues to be used as a reference sediment in A. abdita tests at the
EPA-Narragansett, RI, laboratory, was collected on Dec. 5, 1989, from the South Reference
site described in Scott and Redmond (1989), and pressed through a 2 mm sieve. Both
sediments were held at 4°C until used.
There were 4 treatments, each with 6 replicates, all conducted at 20°C: offspring of
cultured animals tested in Yaquina Bay sediment, offspring of cultured animals tested in
Long Island Sound sediment, offspring of field amphipods tested in Yaquina Bay sediment,
and offspring of field amphipods tested in Long Island sediment. Because of an abundance
of amphipods, an additional treatment with 3 replicates was added with the offspring of field
amphipods in Yaquina Bay sediment at 15°C.
This experiment was conducted using the quart jar renewal system as described for
Experiment #3 (Temperature and Nutrition Effects), with a mixture of 100 ml algae and 500
ml seawater added daily per jar. All treatments were fed a 1:1 mixture by volume of P.
paradoxa and P. tricornutum, plus 1 ml/jar of a brine shrimp (Artemia salina) nauplii
suspension (density not measured) at regular intervals when amphipods became large enough
to capture the nauplii and were approaching sexual maturity. Preliminary observations
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3-8
indicated that A. abdita would capture and eat A. salina nauplii if they came within capture
range. Cell counts of algal cultures were taken daily.
To maximize the hypothesized sediment effect, brooding females and their offspring
were held in the same sediment in which they were to be tested, and fed the algal mixture
plus brine shrimp nauplii. Also, females were allowed to release their young in sediment
rather than in seawater, in case the juveniles had been stressed by the water- only releases
in previous experiments.
The experiment was initiated with amphipods which were 1-6 (field offspring) or 1-7
(cultured offspring) days old, with 20 amphipods per test container. Treatments testing
cultured offspring were initiated 9 days after treatments testing field offspring, since females
from the two sources released their young at different times. Fifteen replicate jars were
sampled after 10 days to examine survival and growth endpoints, and the remainder ended
after 42 days. Amphipods from the 10 day sampling and initial samples were preserved and
measured to determine growth in a 10-day period. T-tests were conducted to determine
significant differences between 10-day treatments.
Experiment #6: Container, Aeration, and Nutrition Effects
This experiment examined the potential effects of container type, aeration, and the
amount of food on reproduction. The experiment was initiated with cultured juvenile
amphipods of indeterminate age, at 20°C, and was terminated after 56 days. There were four
treatments with three replicates of 30 amphipods each: 1) aerated quart jar exposure
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3-9
containers which were fed a 1:1 by volume algal mixture of P. tricornutum: P. paradoxa; 2)
nonaerated quart jars fed the algal mixture; 3) aerated quart jars fed the algal mixture plus,
twice per week, blended A. salina nauplii (<48 hours post-hatch, blended 10-30 sec. in
seawater); and 4) aerated plastic bins (12 cm high x 17 cm diameter) fed the algal mixture.
The quart jars were renewed daily using the renewal system described for Experiment #3
(Temperature and Nutrition Effects) above. Bins were renewed by pouring off about 2/3 of
the overlying water and replacing it with the algal mixture. Each jar received approximately
250 ml algal mixture and 350 ml seawater daily; each bin received 300 ml algal mixture and
400 ml seawater. The amount of algae provided was greatly increased to determine whether
the amounts (i.e., density) used in previous experiments had be sufficient. Algal density was
not measured, but typically ranged from 106-107 cells/ml in the undiluted stock culture.
Minimal daily biological observations were made, and dead amphipods were not removed
daily, due to limited visibility in the containers. No statistical analyses were performed.
3.3 RESULTS AND DISCUSSION
Table 3.2 summarizes the physical and feeding data for all six experiments.
Temperature and salinity variation was minimal. Where cell counts were taken, the
estimated number of cells per exposure container was approximately the same in all
experiments, 3 to 4 x 108 cells/replicate/day, and exceeded values used in previous successful
long-term experiments with A. abdita. Scott and Redmond (1989) and Gentile et al (1987)
reported delivering 108 - 109 cells P. tricornutum per day to each of their gallon jar exposure
containers. The gallon jars had approximately a 3000 ml water volume, yielding 108/3000 to
109/3000 = 3.3 x 104 to 3.3 x 105 cells/ml. The quart jars in our study have approximately a
-------
3-10
600 ml water volume, and the algal cell density in each exposure chamber was projected to
be approximately 107-108 cells/replicate/d [i.e., 3.3x10" - 3.3xl05 cells/ml x 600 ml/replicate «
2xl07 - 2xl08 cells/replicate], which was comparable to cell densities used by Scott and
Redmond (1989).
Experiment #1: Temperature and Salinity Effects
Live recovered amphipods appeared healthy, since they exhibited normal pink
coloration and were active. The population in each jar increased from 4 to 17 times its
original value of 30. Table 3.3 shows the number and life stage of animals recovered for
each sampling period. Terms for the various life stages were taken from Scott and
Redmond (1989): females with oostegites just developing were called developing females
or FdV, females with eggs in the oviduct FE, females with eggs or developing young in
the brood pouch FOV or ovigerous (brooding) females, females which have released their
young spent females or FS, males M, and undifferentiated, including juveniles and
subadult males and females, UD.
Higher temperature accelerated the timing of life cycle events. Jars at 25°C
produced the first Fx juveniles (F0 designates the adults initially added to the jars), the
first observed sexually mature Fj individuals, and at least some F2 juveniles by week 8
(earliest sampling time). There were no F2 individuals in the 20°C treatments at 8
weeks.
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3-11
The day 66 and day 70-73 data suggested that the combination of 25°C and 20%o
was not a good long term condition for amphipods from this population. At both
sampling times the lowest recovery was found in that treatment (Table 3.3). Subsequent
experiments were conducted with ambient salinity seawater (28 - 35%o).
Previous experiments conducted at EPA's Narragansett laboratory, however
(Redmond and Scott, unpublished data), suggested 20%o might be an acceptable short-
term salinity for amphipods from this field source. In one experiment, amphipods
collected at 18°C and 30%o were immediately tested for 96 hours without acclimation,
under static daily renewal conditions in jars with no sediment or aeration. There was
0% survival at 5 and 10%e, 48% at 15%o, and greater than 98% survival at 20,25, 30, and
35%o. A second experiment utilized amphipods collected at -1°C and 27%o, acclimated
to 20°C at a salinity of 30-3 l%o, then exposed for 96 hours to a range of salinities.
Exposure jars contained uncontaminated sediment in which the pore water salinity had
been adjusted with deionized water to that of the salinity treatment. The overlying
water in each exposure jar was renewed daily. Results resembled those of the first
experiment: 0% survival at 10%c, 60% at 15%o, and greater than 97% at 20 and 30%c.
Experiment #2: Life Cycle at 25°C
Survival was >90% in replicates sieved at day 14, and <10% of the individuals
were unaccounted for in any replicate (Table 3.4). The 14-day data would thus meet the
standard criteria for acceptable control survival in a 10-day acute test with this species
(ASTM 1990). Additionally, nonsenescent survival for all sampling times was £90% in
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3-12
all but one replicate. Amphipods were observed feeding during the experiment, and
survivors were active and had normal healthy coloration. The survival curve (Figure
3.1) resembles the one hypothesized for control survival in the chronic test design
(Appendix E): high survival in the early part of the life cycle, followed by senescence and
death in the later weeks.
A preliminary outline of the A. abdita life cycle was obtained from this
experiment. Amphipods grew (Figure 3.2), became sexually mature, and produced eggs.
A female with eggs in the oviduct was first observed when amphipods were 18-20 days
old, a male at 20-22 days old, and ovigerous females at 23-25 days old. (The daily
observation data were qualitative in the sense that not all life cycle events were
necessarily observed). Young were released in only one jar, when initial animals were
34-36 days old.
Ovigerous females observed to be brooding eggs in an early stage (dark brown,
no gut or eyes formed) did not necessarily produce young. For instance, no young or
brooding females were recovered from the first replicate sampled at 21 days (Table 3.4).
However, at least one brooding female was observed in this jar on day 15 of the
experiment, and one spent female was recovered. The brooding female's eggs were
apparently not fertilized and probably disintegrated.
When young were produced or eggs were obviously fertilized (advanced
development), mean number of young per female was 9.8 - 19.5 ([13+26]/4 - [13+26]/2)
(Table 3.5). Scott and Redmond (1989) reported means of 13.6 and 15.8 eggs/female in
laboratory-produced control females from their chronic tests, amphipods from the same
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3-13
source locality as in our study. Their ovigerous females were larger than ours, and since
number of eggs per female was related to female size (Mills 1967), our fecundity data
from successfully reproducing amphipods were not unreasonable. However, successful
reproduction was only observed in 2-4 females in the whole test (Table 3.5).
Newly released amphipods could easily be obtained by holding brooding females
in an aerated beaker of seawater and harvesting juveniles within 24 hours of their
release, as described in Appendix E. Juveniles collected in this manner were held in
sediment for 8-10 days and survival at the end of that time was 90.5%. A test could
therefore be started with newly released juveniles, although 8-10 day old juveniles were
easier to work with. The most time-consuming step in obtaining newly-released
juveniles was the isolation of ovigerous females, and if healthy producing cultures could
be established, that effort could be reduced.
It was possible to accurately determine the number of females in each replicate
despite starting the experiment was with juvenile animals and variability in the
experience of laboratory personnel with life stages and sexes of A. abdita. Only 6 out
of 181 amphipods initially added to the experiment were not accounted for, or 3.3%
(Table 3.4). Undifferentiated (UD) amphipods recovered on sampling days 14, 21, and
28, introduced some error into the sex ratios. However, in an actual chronic test
sampling would not take place until test day 35, and the numbers of UD recovered
would be reduced. UD's were likely immature males, since developing females were
identified even at the earliest sampling time. The ratio of males:females in the exposure
chambers varied from 4:6 to 8:2 if UD's were assumed to be immature males.
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3-14
Experiment #3: Temperature and Nutrition Effects
Two factors potentially responsible for low reproduction in Experiment #2 (Life
Cycle at 25°C) were examined in this experiment. Either the diet provided (food source
and ration) or the high temperature (25°C), or both together, could have produced
detrimental effects which might not have appeared until after several generations. Some
nutritional factor might have been lacking, or a long period of time at a high
temperature could have increased nutritional needs.
Unfortunately, survival in this experiment was poor, and survival differences
could not be definitively related to temperature or feeding treatments (Tables 3.6 and
3.7, Figure 3.3). The algal mix with Neo-Novum® seemed to produce higher survival
after 14 days than the other food source treatments (Table 3.6), but that pattern did not
continue in the 41-day nonsenescent survival data (Table 3.7).
Survival of newly-released juveniles in Experiment #2 (Life Cycle at 25°C) after
8-10 days was 90.5%; after 10 days in this experiment, survival was 13-77% (Figure 3.3).
Similarly, nonsenescent survival for amphipods 43-59 days old in Experiment #2 was 90-
100%, but only 0-70% for 41-day-old amphipods in this experiment. The 41-day survival
data showed the expected senescent mortality pattern (i.e., a dead male was first
observed on day 19), but much poorer survival in the early portion of the life cycle than
in Experiment #2 (Life Cycle at 25°C; Figure 3.1, Table 3.4). The percentage of missing
amphipods ranged from 10-90%. Missing amphipods were assumed to have died in the
first portion of the experiment, when they were very small and not easily observed.
-------
3-15
Initial shipping and handling stresses on test amphipods might account for the
survival difference between the two experiments. Experiment #2 (Life Cycle at 25°C)
used laboratory-produced brooding females, whereas this experiment utilized field-
collected females. The field-collected females had been shipped in seawater-only, under
poor shipping conditions (ice packs omitted); most arrived dead. Those which carried
early stage broods and were still active on arrival were used to isolate juveniles for the
experiment, after the broods matured for 2 weeks. Even though females survived and
broods matured, the initial shipping stress on the developing broods may have been
significant.
Tested juveniles may have been further stressed in this experiment because at
least some of them had been in the brood pouch under no-sediment conditions for 3 days.
In Experiment #2 (Life Cycle at 25°C), females were sieved from holding jars and placed
into beakers with seawater over a 3-d period. During this time, juveniles were
harvested daily from the beakers and placed in holding jars until the test began.
Frequently, females bearing late-stage broods released their young soon after being
removed from the sediment. Thus, most of the test animals were probably without
sediment for <24 h. In this experiment, females were sieved from the sediment and
placed in seawater beakers on the first day; juveniles were harvested from the beakers
over the next 3 d and placed directly into the experimental chambers. Thus, juveniles
harvested on the second or third day had been in the brood pouch under no-sediment
conditions for ca. 48-72 h. The 14-day treatments receiving the algal mixture plus dry
food were the only 14-day treatments started with animals isolated during the first 24
hours, and had the highest survival. Those 41-d treatments with the lowest
nonsenescent survival (25°C, P. paradoxa; 25°C, mix+dry) had been started with animals
-------
3-16
isolated during the second day; other treatments had been started with amphipods
isolated on the first day.
Possibly the water-only isolation affected survival because shipped broods were
already stressed. The double stress could partially explain reasonable production in
culture jars started with late stage brooding females that had survived shipment; late
stage broods may also have been less stressed by shipping. However, it does not explain
why "extra" newly-released test juveniles, collected after the experiment had been set
up, grew and produced a new generation in a culture jar. In that case, sufficient
numbers of juveniles may have been added (about 100) for reproduction to occur even
if there was poor survival.
Differences in lengths of amphipods recovered were detected after the 14-day test
period (Table 3.6, Figure 3.4). Newly released amphipods were uniform in size.
Amphipods in all fed treatments were significantly larger than initial animals and unfed
controls. At 20°C, amphipods were significantly larger when fed the algal mixture or the
mixture with Neo-Novum® than when fed R paradoxa alone. Amphipods in the fed 25°C
treatments were significantly larger than those in the corresponding 20°C treatments.
(In all cases, p<0.05). Increased growth in the algal mix and algal mix with Neo-
Novum® treatments was most likely due to increased nutritional diversity. Because food
was supplied in excess, differential treatment survival presumably did not affect growth.
These data should, however, be considered preliminary in light of the survival problem
in this experiment.
-------
3-17
The factor(s) responsible for low reproduction were not identified with this
experiment. No young were produced, although males, females with eggs in the oviduct,
and brooding females were observed.
Experiment #4: Density Effects
Amphipod survival in this experiment was poor (Table 3.8). Only 28% of the
juvenile amphipods isolated survived the 8-10 day pretest holding period. Up to 35% of
the tested amphipods were unaccounted for in some replicates, suggesting high mortality
initially when dead amphipods were small and difficult to see. Nonsenescent survival
was still only 40-70%. Recovered amphipods had healthy coloration and were active, and
both males carrying sperm and females carrying eggs in the brood pouch were observed.
However, reproduction did not occur in any of the replicates, so the question of how
amphipod density in an exposure container might affect reproduction was not resolved.
Experiment #5: Sediment and Animal Source Effects
Broods of the field-collected amphipods were evidently stressed during the
processes of shipping, acclimation, and holding. There was no apparent survival
difference at 10 days between the two sediment treatments (Yaquina Bay and Long
Island Sound), but survival of the cultured amphipods (>95%) was better than that of
the offspring of field-collected animals (65-85%) (Table 3.9). After 43 days, survival
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3-18
varied from 20-85% (Table 3.10). Nonsenescent survival of field offspring was 35-70%,
whereas that for cultured amphipods was 85-100%. There were also more animals
missing among the field amphipods, likely small animals which died unobserved early
in the test.
A treatment-related length difference was detected in this test after only 10 days,
even though the ages of initial amphipods varied by 5 days. Graphical examination of
the length data (Figure 3.5, Table 3.9) indicated that the only potential growth difference
was between the 15°C and 20°C treatments with offspring of field animals in Yaquina
Bay sediment. A T-test showed that those animals tested at 20°C were significantly
larger.
None of the amphipods in this experiment reproduced, so the potential effects of
sediment and animal source on reproduction were not defined. Also, although all
juveniles tested were released from females held in sediment rather than in seawater
only, elimination of the hypothesized water-only stress didn't result in successful
reproduction, or in control-level survival (>90%) across all treatments.
Experiment #6: Container. Aeration, and Nutrition Effects
Survival data from this experiment are shown in Table 3.11. Since dead
amphipods were not removed daily, no conclusions could be drawn regarding treatment
vs. natural senescent mortality. However, since no amphipods in the nonaerated jars
survived, it was clear that aeration was required in tests with this species, even if
-------
3-19
overlying water and food material were renewed daily. Tested animals did not
reproduce, so effects of the tested parameters on reproduction could not be determined.
General Discussion
The expected long-term control survival pattern for A. abdita was demonstrated
in Experiment #2 (Life Cycle at 25°C, Figure 3.1), which stands in contrast to those of
later experiments (e.g., Experiment #3, Temperature and Nutrition Effects, Figure 3.3).
Unacceptably high numbers of amphipods died early in the life cycle, before senescence,
in Experiments #3 and #4 (Temperature and Nutrition Effects and Density Effects) and
in the portion of Experiment #5 (Sediment and Animal Source Effects) which tested
offspring of field-collected animals. Early deaths were indicated by poor recovery after
pre-experimental holding periods (#4 and #5), by poor survival in 10-day and 14-day
treatments (#5 and #3, respectively), and in all three experiments by the relatively large
number of missing amphipods (probably small animals which died unobserved), and the
nonsenescent survival values.
Clearly some factor or factors involved in the process of shipping, acclimation, and
holding could affect survival of this species in laboratory experiments. This process was
a suspect stress in Experiment #3 (Temperature and Nutrition Effects), and results of
Experiment #5 (Sediment and Animal Source Effects) showed that offspring of cultured
animals survived better than offspring of recently shipped animals. The results
particularly suggest that brooding A. abdita should not be shipped, unless the stress(es)
could be identified and eliminated, and that individuals developing in the brood pouch
-------
3-20
may be a very sensitive stage in the life cycle. Other factors besides shipping and
acclimation probably also affected survival, since success of cultures was variable.
Growth of recovered amphipods in Experiments #2,3, and 5 (Life Cycle at 25°C,
Temperature and Nutrition Effects, and Sediment and Animal Source Effects,
respectively) was within the range of values seen in the literature (Figure 3.6). Mills
(1967) developed length-age curves for summer and winter generations of A. abdita from
Barnstable Harbor, Massachusetts, over two years; he derived his curves from length-
frequency distributions of field samples. Lengths of amphipods in our study, and those
from the control treatments in the chronic tests conducted by Scott and Redmond (1989),
fall within the range of the summer curves. As the growth data from Experiment #3
(Temperature and Nutrition Effects) showed, these data were affected by factors,
including nutrition and temperature. Differences in the methods of age estimation and
measurement also introduce some error into the figure.
The life cycle appears to be shorter at 25°C than at 20°C (Figure 3.7, Tables 3.3
and 3.4). We have no definitive data to show whether 25°C (or a salinity of 20%e) was
harmful over the long term.
The question of why this species reproduced so little in this series of experiments
still has not been resolved. Recovered amphipods generally appeared healthy in
coloration and were active. Amphipods grew, became sexually mature, and produced
eggs and sperm, but young were produced only in the first two experiments. We tested
temperature, type and amount of food, type of sediment, type of container, amount of
aeration, density of amphipods in the exposure container, collection of juveniles from
-------
3-21
females held in sediment rather than in seawater only, and offspring of cultured vs.
field-collected ovigerous females. Reproduction was not improved by changing any of the
listed variables, although any or all of them might prove to be important once the critical
factor or factors for reproduction were identified.
Chronic tests have been run previously with this species (Scott and Redmond
1989; Gentile et al. 1985, 1987) yet our success with culture and chronic testing of this
species has been inconsistent. Previous tests utilized F, or F2 laboratory generations of
field-collected organisms, which needed no shipment and sometimes no acclimation, and
thus may have been healthier than our test organisms. As discussed in Chapter I above,
production might be improved by manipulation of photoperiod and temperature to more
closely simulate natural conditions. There could be some natural, noncontaminant factor
in the water from our laboratory area to which this Atlantic population was sensitive or
susceptible, e.g., a bacterial disease, although we have no evidence that this was the
case.
Experiment #2 (Life Cycle at 25°C) showed results very close to what would be
hypothesized with a healthy control population of A. abdita: survival and growth were
good, and there was some reproduction at the proper time. Therefore it seems that the
best approach to further research with this species would be to repeat that experiment,
taking care to correct the problems which were identified in this study. A mixture of
algae should be fed daily, rather than the single species (P. paradoxa) used in
Experiment #2, since results of Experiment #3 (Temperature and Nutrition Effects)
showed that a mixed food source produced better growth. Based on the number of
cells/ml/day provided, as well as the general health, growth, and production of eggs and
-------
3-22
sperm in tested animals, it would appear that the mixed food source we provided was
adequate. If a laboratory conducting a life cycle experiment was not located close to a
source of field animals, offspring of cultured animals rather than those of shipped
animals could be used. Otherwise the experimental design should be as specified in
Appendix E. It might be advisable to try using a flow-through system which delivered
more than 1 volume replacement/day, e.g., 5 volume replacements/day.
It would also be advisable to conduct a series of experiments examining factors
involved in the shipping, acclimation, and laboratory handling processes in more detail.
Once further research identifies the factors involved with poor laboratory
reproduction with A. abdita, and successful tests have been conducted under control
conditions, the next step would be to test the chronic test design and the 10-day growth
tests using a contaminated material. Interlaboratory testing should be initiated to verify
that this species could be tested in seawater from various regions of the country.
-------
3-23
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3-34
Table 3.5. Fecundity of brooding female Ampelisca abdita recovered in Experiment
#2, Life Cycle at 25°C. "Egg" indicates a very early stage of potential young in the
brood pouch, and "gut" indicates that the gut has formed in the developing
amphipods.
Test Amphipod Repli- #eggs/ egg
day age cate# female stage #voung
21 29-31
28 36-38
35 43-45
49 57-59
2
3
3
1
2
3
2
2
1
1
7
2
3
1
1
1
13
1
1
egg
egg
egg
egg
egg
egg
gut
egg
26 *
egg
* 3 spent females were recovered.
-------
3-35
Table 3.6. Ampelisca abdita recovered from various feeding treatments in Experiment #3
(Temperature and Nutrition Effects) after 14 days at the indicated temperatures. P.p. =
Pseudoisochrysis paradoxa at approximately the same concentration as in Experiment #2
(Life Cycle at 25°C). "Mix"=P. paradoxa, Phaeodactylum tricornutum. and Chaetoceros
calcitrans in a 1:1:1 mixture. "Mix+dry" = the algal mix plus about 4mg/day of ground Neo-
Novum®. Mix+dry treatments were started two days before other fed treatments. Ten
amphipods were tested in each replicate.
Live %sur- Mean% Length in
Treatment recovered Missing vival survival mm. x±SD(n)
20°C,nofood 6 4 60 40.0 1.09±0.07(3)
3 7 30
3 7 30
20°C,RE. 5 5 50 50.0 2.15±0.20(3)
3 7 30
7 3 70
20°C, mix 6 4 60 46.7 2.91±0.21(3)
3 7 30
5 5 50
20°C,mix+dry 10 0 100 96.7 3.02±0.13(3)
10 0 100
9 1 90
25°C,no food 4 6 40 50.0 1.06±0.01(2)
6 4 60
25°C, mix 2 8 20 36.7 3.69±0.15(3)
4 6 40
5 5 50
25°C,mix+dry 10 0 100 96.7 3.84±0.08(3)
9 1 90
11 0 100
initial samples 1.08±0.03(5)
-------
3-36
Table 3.7. Ampelisca abdita recovered from various feeding treatments in Experiment
#3 (Temperature and Nutrition Effects) after 41 days at the indicated temperatures.
P.p. = Pseudoisochrysis paradoxa at approximately the same concentration as in
Experiment #2 (Life Cycle at 25CC). "Mix"=P. paradoxa, Phaeodactylum tricornutum,
and Chaetoceros calcitrans in a 1:1:1 mixture. "Mix+dry" = the algal mix plus about
4mg/day of ground Neo-Novum®. Ten amphipods were tested in each replicate.
Treatment
20°C, P.p.
20°C, mix
20°C,mix+dry
25°C, P.p.
25°C, mix
25°C,mix+dry
Live
recovered
5
3
1
1
0
4
2
5
3
0
0
0
1
0
0
0
0
0
Missing
4
6
5
2
5
3
5
2
4
8
9
8
3
3
1
7
7
6
%
survival
50
30
10
10
0
40
20
50
30
0
0
0
10
0
0
0
0
0
% survival w/o
senescence3
60
30
40
40
50
70
50
60
50
20
0
10
70
60
70
30
0
30
a Those individuals which were assumed to have died as a result of senescence (i.e.,
males and spent females) were not counted as dead for the purposes of this
calculation.
-------
3-37
Table 3.8. Percent survival of amphipods, Ampelisca abdita, in Experiment #4,
Density Effects Experiment, after 38 days at 20°C, with varying densities in exposure
containers. Amphipods were 46-58 days old when the experiment was terminated.
Treatment
10 per
replicate
20 per
replicate
40 per
replicate
Live
recovered
4
4
7
8
5
8
14
22
21
Missing
3
3
2
0
4
7
11
6
11
%
survival
40
40
70
40
25
40
35
55
52.5
% survival w/o
senescence3
50
40
70
70
65
50
40
62.5
65
a Those individuals which were assumed to have died as a result of senescence (i.e.,
males and spent females) were not counted as dead for the purposes of this
calculation.
-------
3-38
Table 3.9. Experiment #5, Sediment and Animal Source Effects. Percent survival and length
of recovered amphipods, Ampelisca abdita, from two sources after 10 days in the indicated
sediments. "Field" animals = offspring of ovigerous females from the field; "cultured" animals
= offspring of cultured ovigerous females. At the end of the 10 day exposure period, field
animals were 11-17 days old, cultured animals 11-18 days old. Twenty amphipods were
tested in each replicate.
Animal Sediment Live re- Miss- %sur- Length in mm
source treatment covered ing vival mean% (mean±sd;n=3)
field
cultures
Long Island
Yaquina Bay
Yaquina Bay
15°C
Long Island
Yaquina Bay
13
13
13
14
16
17
14
15
16
20
19
19
20
19
20
6 65 65.0 3.35 ± 0.22
5 65
5 65
6 70 78.3 3.44 ± 0.09
4 80
2 85
6 70 75.0 2.90 ±0.11
5 75
3 80
0 100 96.7 2.81 ± 0.06
0 95
1 95
0 100 98.3 3.00 ± 0.16
1 95
0 100
-------
3-39
Table 3.10. Experiment #5, Sediment and Animal Source Effects. Percent survival
of recovered amphipods, Ampelisca abdita, from two sources after 43 days in the
indicated sediments. "Field" animals = offspring of ovigerous females from the field;
"cultured" animals = offspring of cultured ovigerous females. At the end of the 43-day
exposure period, field animals were 44-49 days old, cultured animals 44-50 days old.
Twenty amphipods were tested in each replicate.
Animal
source
Sediment
treatment
Live re-
covered
Miss-
ing
% sur-
vival
% survival
w/o senescence3
field Long Island 7 4 35 60
9 5 45 70
4 8 20 35
YaquinaBay 8 4 40 70
8 8 40 55
9 5 45 45
cultured Long Island 15 3 75 85
13 3 65 85
14 1 70 95
YaquinaBay 12 1 60 100
17 0 85 100
8 3 40 85
a Those individuals which were assumed to have died as a result of senescence (i.e.,
males and spent females) were not counted as dead for the purposes of this
calculation.
-------
3-40
Table 3.11. Amphipods, Ampelisca abdita. recovered after 56 days at 20°C in
(Container, Aeration, and Nutrition Effects Expe
e exposed in each of three replicates per treatment.
Treatment
aerated
bins
aerated
jars
nonaerated
jars
aerated jars
fed Artemia
Live
recovered
18
4
15
8
3
10
0
0
0
13
7
16
% survival
60.0
13.3
50.0
26.7
10.0
33.3
0
0
0
43.3
23.3
53.3
Mean
%survival
41.1
23.3
0
40.0
-------
APPENDIX A
LITERATURE REVIEW OF SELECTED CHESAPEAKE BAY AMPHIPODS
INTRODUCTION
The ecological and life history characteristics of over 60 amphipod species which have
been reported from the Chesapeake Bay are summarized in Table A-l. Further detail of
these characteristics for five amphipod species (Lepidactylus dytiscus, Ampelisca abdita,
Leptocheirus plumulosus, Monoculodes edwardsi and Neohaustorius schmitzi) are presented
in the text below and summarized in Table A-2. These species were selected as possible
candidates for development as sediment toxicity test organisms for Chesapeake Bay region.
A sixth species (Hyalella azteca) is listed in Table A-2, but its ecological and life history
characteristics are not further discussed in the text because this is an established sediment
toxicity species, and information concerning H. azteca may be found in ASTM (1990a).
A-l
-------
A-2
Characteristics of Selected Species of Interest
Lepidactylus dytiscus
(Figures A-l,A-6,A-7)
Habitat. Distribution, and Ecology
L.dytiscus is found from the upper Chesapeake Bay to the Florida Atlantic coast, typically
in the intertidal zone, but also subtidally to 3m, often sympatric with other haustoriids such
as Neohaustorius schmitzi, and others of the genera Acanthohaustorius, Parahaustorius,
Protohaustorius, Haustorius, and Pseudohaustorius (Grant and Lazo-Wasem 1982, Bousfield
1970, Dexter 1967, Croker 1967a).
This species burrows freely in clean to muddy sand, typically sand with a high silt or
organic content (Mountford et al 1977, Dexter 1967, Grant and Lazo-Wasem 1982), but not
excessive silt-clay (Grant and Lazo-Wasem 1982). Deaver and Adolphson (1990) found that
in short-term experiments, survival was slightly better in 95% sand (90% survival) than in
50%sand/50% mud (79% survival) or > 85% silt/clay (77% survival). The burrowing pattern
of L. dytiscus is almost identical to that of Neohaustorius schmitzi (Howard and Elders 1970).
L_. dytiscus may be found ranging from estuarine sands to exposed beaches, to a depth of 9
cm, but is most common in sheltered sand habitats (Grant and Lazo-Wasem 1982, Croker
1967a, Fox and Bynum 1975), in approximately the upper 5 cm (Croker 1967a). The amount
of light available may have an effect on depth of burrowing (Howard and Elders 1970). In
North Carolina, it has been reported in densities of up to 1500/m2, concentrated at the mid-
-------
A-3
tide level (Dexter 1967), and may be commonly taken in estuarine plankton (Dexter 1967, Fox
and Bynum 1975).
Feeding and Nutrition
L,. dytiscus is reported to be a suspension-feeder (Croker 1967a, Bousfield 1970). Gut
contents of field-collected specimens included diatoms, detritus and algae (Croker 1967a).
In the laboratory, L. dytiscus fed on materials in field-collected beach sand, and on a slurry
composed of beach sand detritus, diatoms, and crushed fecal pellets of the ghost shrimp
Callianassa major (Croker 1967a).
Reproduction
Reproduction in this species may take place year-round, with maximum activity in the
spring and summer (Grant and Lazo-Wasem 1982, Croker 1967a, Dexter 1967). In Georgia,
females were usually dominant in the population, and an annual life cycle was reported
(Croker 1967a). Dexter (1967) reported the mean length of gravid females to be 5.42 mm,
with a mean egg number of 11.0, and young released at 1.36 - 1.52 mm.
-------
A-4
Physical Tolerances
This amphipod tolerates salinity conditions from fresh water to fully marine, and may
occur in brackish or virtually fresh water (Grant and Lazo-Wasem 1982). It has been
reported from study areas of 5 - 30%c in North Carolina (Dexter 1967), and 7 - 18%o in the
Chesapeake Bay (Mountford et al 1977). Temperature at the latter site ranged from -0.3 to
27.5°C. Deaver and Adolphson (1990) reported >90% survival when L. dytiscus were tested
for 14 days in salinities ranging from 5 to 40%o, and held and tested them in the laboratory
at 20°C. Ij. dytiscus is reported to be fairly tolerant of desiccation and high temperature,
and has a negative response to light (Croker 1967a).
Distribution and Abundance in the Chesapeake Bay
In the Chesapeake Bay, Ij. dytiscus was reported to be dominant in 3 m sand communities
at Calvert Cliffs, with a mean summer density of 151/m2 (Mountford et al 1977), and at sand-
bottom stations in the James River (Jordan and Sutton 1984, Diaz 1989). Other researchers
have also reported it in the Bay (Loi and Wilson 1979, Feeley and Wass 1971).
Other Notes
Ecotypic plasticity has been noted in size and morphology of L,. dytiscus, and its name
means "scaly-fingered diver." (Grant and Lazo-Wasem 1982). This species may be abundant
-------
A-5
in the plankton at times of the new moon (Williams and Bynum 1972). Ray Alden (Old
Dominion University, Norfolk, VA) has attempted to culture this species and has used it in
several sediment toxicity test exposures (pers. comm.); Deaver and Adolphson (1990) reported
successfully testing this species in 96-hour seawater-only acute toxicity tests with cadmium
and fluoranthene.
Marcia Nelson (U.S. Fish and Wildlife, Columbia, MO) and Scott Carr (U.S. Fish and
Wildlife, Corpus Cristi, TX) have also worked with a related species (_L. triarticulatus) (pers.
comm.); M. Nelson cultured this amphipods by feeding Cerophyll or rabbit chow about once
a week, and reports their life cycle to be 3-4 months at 20°C.
Ampelisca abdita
(Figures A-2, A-6, A-7)
Habitat, Distribution, and Ecology
Ampelisca abdita is a tube-dwelling amphipod belonging to the family Ampeliscidae,
found mainly in protected areas from the low intertidal zone to depths of 60m. It ranges
from central Maine to south-central Florida and the eastern Gulf of Mexico (Mills 1964,
Bousfield 1973), and has also been introduced into San Francisco Bay (Nichols and Thompson
1985). Where A. abdita are present, they are often dominant members of the benthic
community with densities up to 110,000 m'2 (Nichols and Thompson 1985, Stickney and
Stringer 1957, Santos and Simon 1980). This species generally inhabits sediments from fine
-------
A-6
sand to mud and silt without shell, although it may also be found in relatively coarser
sediments with a sizable fine component (Mills 1967). A.abdita is often abundant in
sediments with a high organic content (Stickney and Stringer 1957).
This amphipod is a common food source for fish. "Ampelisca sp.," probably A.abdita, was
reported in gut contents of silversides, Menidia menidia. and juvenile flounder
Pseudopleuronectes americanus in the lower Pettaquamscutt River, Rhode Island (Mulkana
1966), and A. abdita was reported to be food for spot Leiostomus xanthurus and star drum
Stellifer lanceolatus (Stickney et al 1975).
Feeding and Nutrition
Ampelisca abdita is a particle feeder, feeding both on particles in suspension and on those
from the surface of the sediment surrounding their tubes. Gut contents of field-collected
specimens have been found to include algal material, sediment grains, and organic detritus
(Mills 1967, Stickney and Stringer 1957).
Reproduction
In the colder waters of its range, A. abdita produces two generations per year, an
overwintering generation which breeds in the spring and a second which reproduces in mid
to late summer (Mills 1967, Nichols and Thompson 1985). In New England, breeding of the
-------
A-7
overwintering generation begins when the water temperature is about 8°C, but in warmer
waters south of Cape Hatteras, breeding may be continuous throughout the year. Adults
mate in the water column, and intense breeding activity is correlated with the full moon and
spring tides. Females in a population from Barnstable Harbor, Mass., were found to carry
a mean of 26 eggs. Juveniles are released after approximately two weeks in the brood pouch,
at about 1.5 mm in length. It then takes 40-80 days for newly released juveniles to become
breeding adults (Mills 1967).
Physical Tolerances
A. abdita has been collected in waters of -2 to 27°C (Redmond and Scott, unpublished
data). It is euryhaline, and has been reported in waters which range from fully marine to
10%c salinity (Bousfield 1973). This species is photonegative, and has a strong mortality
response when exposed to sunlight (Redmond and Scott, unpublished data).
Distribution and Abundance in the Chesapeake Bay
A. abdita has been reported to be present in several areas of the lower Chesapeake Bay
(Reinharz and O'Connell 1983, Boesch 1977, Marsh 1973, Orth 1973, Schaffner et al 1987),
and in some cases it is abundant or dominant (Dauer et al 1984, Boesch 1973, Holland et al
1988). Lippson et al (1979) reported Ampelisca spp. from the Potomac, and Lynch and
Harrison (1969) reported A. abdita from the York River, Virginia.
-------
A-8
Other Notes
An acute test procedure with this species is well-established, as is its sensitivity to a
variety of contaminated materials (ASTM 1990b, Redmond et al in prep., Scott et al in prep.,
DiToro et al in press, Breteler et al L989, Yevich et al 1986, Rogerson et al 1985, Botton
1979). Chronic tests have also been conducted, and this amphipod can be maintained in the
laboratory with an algal diet (Scott and Redmond 1989, Gentile et al 1985).
Leptocheirus plumulosus
(Figures A-3, A-6, A-7)
Habitat, Distribution, and Ecology
Leptocheirus plumulosus ranges from Massachusetts to Florida, from the intertidal zone
to water approximately 5 m in depth. It builds an unlined, U-shaped burrow of sand grains
and debris in the upper 5-7 cm of sediment, and is typically found in mud to sandy mud and
detritus, especially in areas with a current (Bousfield 1973, Sanders et al 1965, Holland et
al 1977, Jordan and Sutton 1984, Reinharz and O'Connell 1983, Holland et al 1987).
Shoemaker (1932) reports it at depths between 3 and 12 m in Chesapeake Bay.
-------
A-9
L^. plumulosus was reported in gut contents of the American eel Anguilla rostrata and the
blue crab (Callinectes sapidus) in the James, York, and Rappahannock Rivers (Wenner and
Musick 1975). Marsh (1988) found that in the Patuxent River, there was close timing
between the late spring to early summer increase in production of _L. plumulosus and an
increase in abundance of predators, which consisted primarily of juvenile fish. He
determined that this was an opportunistic species with "boom and bust" population dynamics.
In winter, population growth was limited by low temperatures, and in early spring by
nitrogen availability. In late spring to early summer, most reproduction and population
growth occurred, correlated with warmer temperatures and increased nutrient availability.
By late summer, juvenile survival, growth, and reproduction appeared to be limited by lack
of essential micronutrients.
-------
A-10
Feeding and Nutrition
_L. plumulosus is reported to be a surface deposit feeder (Holland et al 1988, Marsh 1988),
and may leave its burrow to forage on the sediment surface (Sanders et al 1965). McGee et
al. (1990) maintained this species in laboratory experiments for 28 days on 6mg Tetramin fed
3 times per week.
Reproduction
This species has an annual reproductive cycle, with ovigerous females most abundant
from May to September, and two broods per female (Bousfield 1973). Marsh (1988) reported
that in the Patuxent River, Maryland, during 1984 - 1986, this species reproduced primarily
in May and June. Toward the end of this period, growth of juveniles, mean body size of
gravid females, and fecundity decreased. Fecundity decreased even when normalized for
female body size and it was concluded that the population dynamics were controlled by the
food supply at that time. C. Schlekat and B. McGee collected gravid L. plumulosus as late
as December and as early as February from subestuaries in northern Chesapeake Bay (pers.
comm.).
Physical Tolerances
-------
A-ll
Although able to tolerate salinities from 3 - 31%c (Sanders et al 1965), this species is
generally found in low to mid salinity areas (Boesch et al 1976, Reinharz and O'Connell
1983), and may be termed an estuarine endemic species (Holland et al 1987). Holland et al
(1988) classify this amphipod as one of a group of organisms tolerant of a wide range of
salinities and sediment types, but note that it had high production values in 5-10%o habitats
in Baltimore Harbor and the Chester River. Similarly, Marsh (1988) reported L,. plumulosus
as a dominant species at a Patuxent River site where salinity varied from 10 to 14%c.
Schlekat et al (1992) reported no significant differences in adult survival in salinities ranging
from 2 to 32%e in 10 to 28 day laboratory experiments. It has been collected in water
temperatures of 0-30°C (Holland et al 1977, Marsh 1988, Jordan and Sutton 1984). Marsh
(1988) reported that this species was inactive when temperatures were less than 5°C, and
suggested that its optimum temperature for reproduction was probably between 10 and 20°C.
Feeley and Wass (1971) collected _L. plumulosus from a variety of substrates, and Schlekat
et al. (1992) reported good adult survival in the laboratory in sediments with a large
variation in particle size distributions and organic content. The latter authors also
successfully tested these amphipods using artificial seawater.
Distribution and Abundance in Chesapeake Bay
L. plumulosus is widely distributed in the upper Bay and tidal tributaries (Jordan and
Sutton 1984, Holland et al 1987, Mountford et al 1977, Holland 1985, Shoemaker 1932) and
is frequently abundant or dominant (Reinharz and O'Connell 1983, Hines and Comtois 1985,
-------
A-12
Dauer et al 1987, Schaffner et al 1987, Diaz 1989, Holland et al 1988, Holland et al 1977,
Marsh 1988).
Holland et al (1988) observed that the abundance of this species in Baltimore Harbor has
increased over the past decade, to a maximum of approximately 15,000/m2 in 1987 from
essentially zero in 1970, apparently due to a lower contaminant load in that area. They also
reviewed power plant impact studies from the Chesapeake, which indicated that L.
nlumulosus had higher abundance in thermal impact areas of Chalk Point and Wagner plant
discharges, which they conclude is due to organic enrichment from entrainment mortalities.
This agrees with the findings of Diaz (1989), who reported that L. plumulosus was dominant
in a soft-bottom community in the immediate area of a large sewage outfall. In the thermal
impact area of the Morgantown plant, reproduction in this species started and ended earlier
than normal (Holland et al 1988).
Other Notes
L. plumulosus may be more abundant in the plankton at times of the new moon (Williams
and Bynum 1972). Schlekat et al (1992) successfully tested this species in 4-day seawater-
only tests and 10- to 28-day sediment tests.
-------
A-13
Monoculodes edwardsi
(Figures A-4, A-6, A-7)
Habitat, Distribution, and Ecology
Monoculodes edwardsi is widely distributed, ranging from the Gulf of St. Lawrence to
Cape Cod and the mid-Atlantic to north Florida and the Gulf of Mexico (Bousfield 1973). It
burrows freely in fine and silty sands from the low intertidal to 75 m (Bousfield 1973,
Mountford et al 1977, Bousfield 1970, Watling and Maurer 1972, Fox and Bynum 1975).
Feeley and Wass (1971) reported taking it at all depths, from the upper layers and surface
of the bottom, and in both sand and mud with considerable detritus. Van Dolah and Bird
(1980) suggest that it probably burrows partially exposed at the sediment-water interface.
M. edwardsi has been reported in gut contents of young-of-the-year striped bass, Morone
saxatilis in the James, York, and Rappahannock Rivers (Markle and Grant 1970); of the
American eel Anguilla rostrata in the York River (Wenner and Musick 1975); and of the
Atlantic croaker Micropogon undulatus, the spot Leiostomus xanthurus and the star drum
Stellifer lanceolatus in estuaries from South Carolina to Georgia (Stickney et al 1975).
-------
A-14
Feeding and Nutrition
This species is reported to be a suspension feeder (Dexter 1969), although Bousfield (1970)
notes that the mouthparts in this genus are of a non-specialized, omnivorous feeding type
rather than being specialized for filter-feeding.
Reproduction
The reproductive cycle for M. edwardsi is annual, with ovigerous females most abundant
May to September and several broods produced per female (Bousfield 1973, Holland 1985).
M. edwardsi may sometimes be abundant in plankton tows, and this may be related to its
breeding cycle (Fish 1925), to temperature changes (Whitely 1948), or to the presence of a
new moon (Williams and Bynum 1972). Whitely (1948) collected egg-bearing females in the
plankton on Georges Bank in June, and concluded that primary reproduction took place in
the summer. Van Dolah and Bird (1980) examined specimens of this species from the
collections of the National Museum of Natural History (location not stated), and reported that
females of 5-7 mm long carried a large number of small eggs relative to other amphipod
species examined (Y = 3.69 e034L, r = 0.81 where Y = egg number and L - female length).
Physical Tolerances
-------
A-15
This species is found in waters which range from fully marine to oligohaline and brackish
(Bousfield 1973). Mountford et al (1977) collected it in abundance at a study site in
Chesapeake Bay where the salinity ranged from 7 to 18%o, and temperature from 1 to 30°C
(Holland et al 1977).
Distribution and Abundance in Chesapeake Bay
M. edwardsi has been reported from several areas in the Chesapeake (Boesch 1977, Ewing
et al 1982, Holland et al 1987, Loi and Wilson 1979), and Mountford et al (1977) reported it
to be dominant in some 3m sand communities. Holland et al (1988) noted that Monoculodes
sp. had consistently high abundances in habitats approximately 5-18%o, and reported it at a
maximum of 3200/m2 at Calvert Cliffs, with year-to-year fluctuations in abundance.
Neohaustorius schmitzi
(Figures A-5, A-6)
Habitat. Distribution, and Ecology
This species is found from Cape Cod to Georgia and southern Florida It coexists with
other haustoriid species, such as Lepidactylus dytiscus, in the intertidal zone (see
aboveXBousfield 1973, Bousfield 1970, Croker 1967a, 1967b, Dexter 1967, 1971).
-------
A-16
N. schmitzi burrows freely in clean to muddy sands, fine to medium in particle size
(Bousfield 1973, Dexter 1967, Croker 1967a). It prefers cleaner sand to that with more silt,
debris, and shell material (Croker 1967a). Although this species can burrow up to 10 cm, in
North Carolina 97% of the population was found in the upper 5 cm, and 83% in the top 2.5
cm (Dexter 1971). Croker (1967a) similarly found it to be most abundant in the upper 2.5 cm
in Georgia. The amount of light may have an effect on depth of burrowing (Howard and
Elders 1970).
In North Carolina, N. schmitzi was found to be most abundant on sheltered beaches in
sounds (Fox and Bynum 1975) or on sandy beaches of inlets (Dexter 1967, 1971). At some
sites this amphipod was present at an average density of 800/m2 (Dexter 1967). N. schmitzi
was reported most abundant in the upper intertidal zone in Georgia (Croker 1967a) and
North Carolina (Fox and Bynum 1975, Dexter 1967), middle and lower intertidal in South
Carolina (Knott et al 1983), and mean low tide (0-2 hrs exposure/ tidal cycle) to high tide zone
(8-10 hrs exposure) in North Carolina (Dexter 1971).
The population may show an aggregated distribution. The amphipods are also
concentrated lower in the tidal zone in winter than in summer, and this may be an avoidance
response to increased wave action and lower temperatures (Dexter 1971). The sexes may be
separated in the tidal zone; females (Bousfield 1970, 1973), particularly gravid females
(Dexter 1971) have been generally reported higher, but Croker (1967a) reported males higher
in the zone. The juveniles located lower in the intertidal zone may be a dispersal stage
(Bousfield 1970).
-------
A-17
Feeding and Nutrition
N. schmitzi is reported to be a suspension-feeder (Bousfield 1970; Croker 1967a, 1967b,
Dexter 1969) which feeds for short periods at frequent intervals in the laboratory and during
periods of tidal immersion in nature (Croker 1967b). Croker (1967b) reported gut contents
of this species to consist of detrital masses which include flagellates, ciliates, diatoms,
unicellular chlorophytes, and bacteria, as well as small amounts of fine sand. Diatoms of the
genera Navicula, Hantzschia, Nitzschia. Coscinodiscus, Grammatophora, and Cocconeis were
ingested. The size range of ingested items was 0.5 - 78.5 um. Ivester and Coull (1975)
reported similar results of gut content analysis, but a larger size range of material of 0.5 -
300um, and noted that zooplankters such as copepods were occasionally found. These
authors also indicate Rhodes (personal communication) has observed N. schmitzi feeding on
polychaetes and other amphipods.
In the laboratory, N. schmitzi was reported to feed on materials in field-collected beach
sand, and on a slurry composed of beach sand detritus, diatoms, and crushed fecal pellets of
the ghost shrimp Callianassa major (Croker 1967a), or on ghost shrimp fecal pellets alone
(Frankenberg 1967).
Reproduction
N. schmitzi is reproductively active for most of the year (Dexter 1969), with the
reproductive period variously reported to extend from May to September (Bousfield 1973), or
-------
A-18
February to October in North Carolina, with peaks in spring (April) and summer (August)
(Dexter 1967, 1971). Knott et al (1983) reported peak densities of animals in February and
May in South Carolina. In North Carolina, there are two generations per year: a winter
generation which lives about 8 months and reproduces in the spring, and a summer
generation which lives about 4 months and produces the overwintering individuals. The
mature individuals in the summer generation are smaller than those in the winter generation
(Dexter 1971). In Georgia, maximum reproductive activity occurred in April, with fecundity
decreasing as autumn approached and the summer generation began to age (Croker 1967a,
1968b).
The maximum range in brood siz,e reported is 2 - 14 eggs per female, with egg number
depending on size of the female and the generation to which she belongs (Dexter 1971). Van
Dolah and Bird (1980) reported the relationship between female length (L) and egg number
(Y) as Y = 1.63L - 2.52, from Georgia specimens in the National Museum of Natural History.
Dexter (1967) reported the mean length of gravid females in North Carolina to be 3.6 mm,
with a mean egg number of 5.69; juveniles were 1.4mm at release. Females appear to produce
only one brood in their lifetime (Dexter 1971), and females have been reported to be dominant
in the population (Dexter 1971, Croker 1967a). Juveniles (Croker 1976a) and some adults
(Williams and Bynum 1972) have been taken in plankton tows, but generally haustoriids in
this subfamily are scarce in the plankton, so although little is known of the mating behavior,
it is not likely that they mate in the water column (Dexter 1971). The young may crawl back
into the brood pouch one or more times before their final release (Croker 1968a).
-------
A-19
Physical Tolerances
Living high in the intertidal zone, this species appears to be relatively tolerant of high
temperature and desiccation. Croker (1967a) measured temperatures of > 39°C in the upper
2.5 cm of Georgia sands with N. schmitzi present, and reported that most animals exposed
to 40°C for 2 hours in the laboratory lived for several months afterward at approximately
27°C. Dexter (1971) collected this species in substrate temperatures from 9 - 33 °C. Animals
exposed to air for 20 min lived for several months once returned to seawater (Croker 1967a).
Gravid females have been found to be more resistant to desiccation than non-gravid females,
males, and juveniles, which may be associated with their concentration in higher tide zones
(Dexter 1971).
This species is reported from 6%c to fully marine conditions in the Chesapeake Bay
(Bousfield 1973), and was collected in 16.5 to 24.4%0 in North Carolina (Dexter 1971).
N. schmitzi is strongly photonegative and very active, and sympatric species may be
separated from collections partially by the speed of their reaction to light (Croker 1967a).
Abundance and Distribution in Chesapeake Bay
This species is reportedly found throughout the Bay (Lippson and Lippson 1984, Bousfield
1973), but thus far no community studies have been located which mention its occurrence.
-------
A-20
Holland (unpublished data cited in Lippson et al 1979) reported this species from the high
mesohaline to near polyhaline areas in the Potomac River.
-------
A-21
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APPENDIX B
PROCEDURES TO MINIMIZE THE RISK OF RELEASING NON-INDIGENOUS
AMPHIPODS, PATHOGENS, WATERS, OR SEDIMENTS
INTO LOCAL WATERS OR WATERSHEDS
General Principles:
1) No non-indigenous animals or sediment will be released into the environment;
2) All water coming into contact with non-indigenous amphipods or sediment in the
laboratories will be sterilized prior to disposal;
3) All equipment or materials (i.e., glassware, paper, plastic, etcj contacting non-
indigenous amphipods or sediment will be contained for proper sterilization;
4) All non-disposable materials will be sterilized or confined to the culture room.
Containment Protocol:
Culture, holding, handling, and experimentation of non-indigenous species is restricted
to "non-indigenous laboratory rooms" separate from those used to hold or culture native
species. Any material (living or dead) or equipment used in "non-indigenous laboratories" are
considered as potentially infected and are treated accordingly. Non-indigenous sediments are
kept in clearly marked, clean sealed containers in a refrigerator and opened only in a "non-
B-l
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B-2
indigenous laboratory". Access to the "non-indigenous laboratory" is limited to trained and
authorized personnel. All drains from "non-indigenous laboratories" are either sealed off or
directed to separate designated holding tanks in which the liquid waste can be sterilized prior
to disposal. The amphipods are cultured in a static-renewal manner to minimize the amount
of water that must be treated. As fresh seawater is added to each culture bin, the displaced
seawater is directed to storage barrels or tanks and treated with chlorine bleach (i.e, 0.5%
chlorine) for sterilization.
Materials for sterilization of animals, sediment or equipment are kept in the "non-
indigenous laboratory". These materials include disinfectant soap (for cleaning hands),
chlorine bleach (for sterilizing all non-human materials and surfaces), mops, sponges, and
buckets (for cleaning floors and surfaces), a bleach dip bath (for sterilizing glassware), and
a labelled trash can (for disposal of contaminated paper, gloves, etc.). All spills are cleaned
immediately.
Personnel wear lab coats while in the "non-indigenous laboratory" and these coats
remain in the laboratory. Hands must be washed with disinfectant soap prior to leaving the
"non-indigenous laboratory".
No equipment leaves the "non-indigenous laboratory" without first being sterilized by
dipping or wiping with bleach. Glass- or plasticware that has been in direct contact with
non-indigenous species, sediment or associated water are held overnight in the chlorine
bleach dip. Water, sediment, and non-indigenous materials are sterilized by either
autoclaving or soaking overnight in a chlorine bleach solution. This material is later
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B-3
neutralized with sodium thiosulfate and then disposed down a sanitary drain to the
municipal sewage system, unless it is also chemically contaminated. Paper and plastic
discarded in the labelled trash can is autoclaved prior to disposal to a municipal landfill.
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APPENDIX C
METHODOLOGY TO ASSESS THE ACUTE TOXICITY
OF MARINE AND ESTUARINE SEDIMENTS
WITH THE BENTHIC AMPHIPOD. LEPTOCHEIRUS PLUMULOSUS
(Leptocheirus plumulosus annex to the ASTM E1367-90 Document, Draft no. 3, May 1992.
Contacts: Beth L. McGee, Christian E. Schlekat, Maryland Department of the Environment,
Ecological Assessment Division, 2500 Broening Highway, Baltimore, Maryland, 21224. phone:
(410) 631-3782, Fax: (410) 631-4105).
A5.1 Ecological Requirements - Leptocheirus plumulosus (family Aoridae) is an
infaunal amphipod distributed subtidally along the east coast of the United States from Cape
Cod, Massachusetts to northern Florida (Bousfield, 1973). In Chesapeake Bay, Ij. plumulosus
is indigenous to oligohaline and mesohaline regions (Feeley and Waas, 1971; Jordan and
Sutton, 1984; Holland et al., 1988), though it can tolerate an even broader salinity range,
from near 0 to 33%0 (Feeley and Waas, 1971; Jordan and Sutton, 1984; Schlekat et al, 1992).
This species constructs U-shaped burrows in sediments ranging from fine sand to silty clay
(Jordan and Sutton, 1984; Holland et al., 1988; Schlekat et al, 1992). Due to its broad
salinity and sediment tolerances, it is a desirable test species for east coast estuarine
sediments and has been used successfully in the assessment of contaminated sediments in
Chesapeake Bay (MDE 1991a,b; Pinkney et al., 1991; see Chapter 2).
C-l
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C-2
A5.2 Collecting and Handling Techniques - Leptocheirus plumulosus is most abundant
in the upper 2 cm of sediment, rarely penetrating to depths below 5 cm (Reinharz, 1981).
Amphipods can be collected with benthic grab samplers (e.g., Peterson, Ponar) from various
tributaries of Chesapeake Bay. The contents of each grab are sieved through a 0.5-mm mesh
screen and the retained material is gently rinsed into polyethylene buckets, containing
collection site sediment and water. These containers are transported to the laboratory where
they are aerated. It is desirable to sort amphipods from collection site debris within 12
hours. A 0.5-mm mesh sieve can be used to separate amphipods from transport sediment.
The material retained on the screen can be rinsed into sorting trays containing collection site
water. Healthy, active amphipods can be removed from detritus by using a bulb pipette of
a suitable size (e.g., one with a 5-mm diameter bulb).
A5.2.1 For acclimation, L plumulosus can be placed in an aquarium (e.g., 40-L)
containing a 1-2 cm deep layer of 0.5-mm sieved collection site sediment at a density of
approximately 200 to 300 per aquarium. Aeration should be vigorous. Two to three days are
sufficient for acclimation to the test environment. A gradual change from collection site
water to test water is desirable. This can be accomplished by gradually increasing the
proportion of test water in the tanks over 2 to 3 days.
A5.2.2 Culture techniques have been developed (see Chapter 1). Presently, laboratory
populations can be maintained through several generations in shallow plastic tubs or glass
aquaria containing a 1-2 cm layer of fine grained sediment from the amphipod collection site
or a texturally similar sediment (Pfitzenmeyer, 1975; see Chapter 1). Water exchange is
static-renewal, with 30-100% of water volume in each container replaced 2 to 4 times per
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C-3
week. Culture containers are aerated, maintained at a temperature of approximately 20°C,
a salinity of 20 g/kg and a photoperiod of 16h light:8h dark. Cultures receive a mixture of
micro-algae (e.g., Pseudoisochrysis paradoxa. Phaeodactylum tricornutum, Tetraselmis
suecica) and approximately 0.1 g of amphipod "gorp" (a mixture offish food flakes, yeast,
alfalfa powder, ground cereal leaves and shrimp maturation feed) 2-3 times per week (see
Chapter 1). Amphipods can be separated from acclimation or culture sediments using a 0.5
mm sieve immediately prior to initiating the toxicity test.
A5.3 Toxicity Test Specifications - The effects of different physical conditions on the
sensitivity of L. plumulosus to toxic materials are currently under investigation. This species
is routinely tested at 20°C or 25°C1. Salinity of overlying water will depend on the objectives
of the study. Toxicity test seawater can be diluted to the same salinity as the interstitial
water of the test sediment, the ambient bottom salinity at the test site or a selected test
salinity in the range of 2 to 32%e. Laboratory investigations indicate Leptocheirus is tolerant
of a range of sediment types (Schlekat et al., 1992); however, a grain size reference should
be included for coarse sediments since these may be somewhat stressful. Fine grained
sediments from the amphipod collection site or laboratory cultures are desirable as the
negative control. The exposure chamber routinely used to test L_. plumulosus is a 1-L glass
beaker. The exposure chamber should be covered with a watch glass to reduce contamination
of the contents and evaporation of the water and test materials. Aeration can be provided
to each test chamber through a 1-mL glass pipette positioned not closer than 2 cm from the
sediment surface. Each test chamber should contain a 2-cm deep layer of sediment and
TA test temperature of 25°C is recommended in this report for
consistency with test conditions in the chronic sediment toxicity
test.
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C-4
enough overlying water to create approximately a 4:1 (v/v) water to sediment ratio. Sediment
and water should be added to the test chambers the day before the amphipods are added to
allow suspended sediment particles to settle, and to allow time for equilibration of
temperature and the sediment-water interface.
A5.3.1 After overnight equilibration of the test chambers, amphipods can be randomly
distributed to each of the containers. It is desirable to sacrifice a random sample of at least
20 animals from those being sorted on day 0 to provide an initial size range estimate of test
animals. Twenty amphipods should be tested per replicate. Animals caught on the water's
surface can be gently pushed under using a glass rod. Amphipods should be allowed 5 to 10
min to burrow into the test sediments. Amphipods that have not burrowed within that time
should be replaced with healthy animals, unless the amphipods are repeatedly burrowing into
the sediment and immediately emerging in an apparent avoidance response. In that case,
the amphipods are not replaced. Amphipods are not removed from the surface of test
sediments during the course of the toxicity test even if they appear dead, since some
amphipods that seem dead might actually be alive and might later rebury into test substrate.
A5.3.2 The toxicity test can be terminated after 10 days by sieving amphipods from
test sediments using a 0.5-mm mesh screen. Mortality is the endpoint for this short-term test.
Burrows generally disintegrate during sieving and animals can be transferred to a sorting
tray for enumeration. The ability of surviving amphipods to rebury into clean sediments can
be used as a sublethal test endpoint.
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C-5
A5.3.3 Other Testing - Partial life cycle tests (28 - 30 days) initiated with juveniles are
being conducted with this species, with amphipod length, reproduction, and survivorship as
viable endpoints. Research is currently underway to determine the optimum conditions for
these tests.
A5.4 Life Cycle and Age Classes - Leptocheirus plumulosus is an annual species
capable of producing a least two broods, with peak periods of reproduction in early to mid
spring and in the fall (Schlekat et al., 1992; Ray, 1982). Gravid females have been observed
in Chesapeake Bay as late as December and as early as February, indicating that timing of
reproduction varies yearly depending on climatic conditions. In cultured populations, females
produce multiple broods and gravid females are available year round (Sewall et al., 1991; see
also Chapter 1). Size range of field-collected test organisms might depend on the size
structure of the field population, as the mean size of amphipods collected in early spring is
generally greater than those collected in the summer or fall. Size range of cultured
amphipods is less variable seasonally. Immature and adult amphipods, approximately 3 to
5 mm as measured from the base of the first antenna to the end of the third pleon segment
along the dorsal surface, should be used in toxicity tests because they are easy to handle and
count. The potential effects of age, size, sex, and seasonal variation of field collected
organisms on the sensitivity of L/. plumulosus to contaminants is currently being examined.
Evidence to date indicates mixed-sex populations within the recommended size range show
consistent responses to field-collected contaminated sediments and 96 h water only exposures
to cadmium (Schlekat et al., 1992; MDE 1991a,b; Pinkney et al., 1991).
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C-6
A5.5 Control Survival - Mean control survival using Leptocheirus must be at least
90% for the toxicity test to be considered valid.
A5.6 Sensitivity - Leptocheirus plumulosus is tolerant of handling and a range of
sediment types and salinities. The sensitivity of this species is comparable to Hyalella azteca
in 96 h water only exposures to cadmium (Schlekat et al., 1992; Pinkney et al., 1991). A
review of benthic surveys and sediment contamination in Chesapeake Bay indicates a
negative correlation between the presence of L_. plumulosus and the degree of contamination
(Reinharz, 1981; Pfitzenmeyer, 1975).
A5.7 Interpretation - When interpreting the results of acute toxicity tests, it should
be kept in mind that the early life stage, the reproductive ability, or the long-term survival
of L_. plumulosus might be affected by contaminants at concentrations lower than those that
produce a lethal response. Partial life cycle sediment toxicity test procedures are under
development for L_. plumulosus and should resolve these questions.
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APPENDIX D
RESEARCH METHODOLOGY TO ASSESS THE CHRONIC TOXICITY
OF MARINE AND ESTUARINE SEDIMENTS
WITH THE BENTHIC AMPHIPOD. LEPTOCHEIRUS PLUMULOSUS
Abstract
A generic chronic sediment toxicity test with the amphipod Leptocheirus
plumulosus is described. This is a draft design which has not been fully tested. This is a
static test conducted at 25°C, 20%o, and with a 16:8 h light:dark photoperiod. Food is
provided three times per week. The experiment begins with 20 juveniles (i.e., <1 day old)
per replicate, 5 replicates per treatment, and is terminated after 28 days. At termination,
the contents of each exposure chamber are sieved through two sieves to collect adults and
offspring. Endpoints are mortality of the initial cohort, body length of the survivors (i.e.,
size), and number of female offspring produced per female survivor (i.e., fertility).
Exposure Conditions
Exposure chambers are 1-L glass beakers. Each chamber receives either test
sediment, performance control sediment (i.e., culture sediment), or reference toxicant in
water. Test sediments may be contaminant-spiked sediment, field sediment, or dilutions
of field sediments. The position of the test and control beakers are randomized within the
D-l
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D-2
water table or room in which the toxicity test is conducted. If there are sufficient
personnel available, the test can run blind, with test and control beakers coded so that
personnel monitoring the test have no knowledge of the identity of treatments in the
exposure chambers.
The recommended temperature is 25°C, which is an acceptable culture and testing
temperature for this species. The lifs cycle of L. plumulosus is also shorter at 25°C than
at 20°C. This amphipod has been tested at salinities within the range of 1.5 - 35%c
(Schlekat et al., 1992), and the salinity of the overlying water should be adjusted to match
that of the sediment pore water (and not vice versa). However, if this test is to be used
with spiked or reconstituted sediments, 20%o is recommended for comparability with other
tests. This is also the recommended culture salinity.
A photoperiod of 16 h light and 8 h dark was selected to approximate conditions
existing during summer when reproduction in the field is expected to be high. The same
photoperiod is maintained for the L,. plumulosus cultures. This photoperiod has been
shown to consistently maintain reproductive activity in Hyalella azteca (de March 1977,
cited in Arthur 1980).
In nature, L/. plumulosus is found in a wide range of sediment types, from mud or
detritus to sand. The performance control uses culture sediment, which is very fine
grained.
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D-3
The exposure container receives 175 ml of test sediment and 725 ml of overlying
water which is aerated constantly. The sediment is added to each beaker the day prior to
starting the exposure. This allows the settling of sediment suspended by the addition of
the overlying water and equilibration of the test sediment and water to the exposure
temperature. Each exposure chamber is covered with a glass plate or evaporating dish to
reduce evaporation of the water overlying the sediment. The exposure chambers are
placed in a constant temperature water bath throughout the exposure.
Food is provided three times per week (Monday, Wednesday, and Friday) by using
a screened siphon tube to remove approximately 400ml/beaker of overlying seawater, and
replacing this with a salinity-adjusted, algal-seawater mixture and lOmg of a dry food
mixture ("gorp"). A glass disk attached to a glass rod is used to prevent disturbance of
the sediment while the algae is added. The algal is prepared as a 1:1 v/v mixture of
Pseudoisochrysis paradoxa and Phaeodactylum tricornutum to a final density of 106
cells/ml. The gorp is a finely ground, dry mixture of 48.5% TetraMin®, 24% dried alfalfa,
24% dried wheat leaves and 4.5% Neo-Novum® (a maturation feed for shrimp mariculture;
Argent Chemical Laboratories, Redmond, WA) suspended in 20%o seawater at a
concentration of lOmg/ml; 1ml of the gorp suspension is pipetted into each exposure
chamber at the time of water renewal and feeding.
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D-4
Controls
Three types of control treatments may be used the sediment toxicity test; two are
mandatory (i.e., the QA/QC performance and reference toxicant controls), and the third
(i.e., the experimental) is optional, but highly desirable. The performance control
measures the responses of JL. plumulosus in the absence of contaminant stress and under
the best possible conditions for the amphipods. The performance control uses culture
sediment as the test substrate and is conducted at 20%o and 25°C. The exposure periods
is 28-d and is conducted in all ways the same as test sediments. Performance controls are
used for QA/QC, to assure that the test organisms are healthy. The performance control
is replicated five times.
The reference toxicant control tests the sensitivity of the animals to a single
toxicant under repeatable exposure conditions. The reference toxicant control consists of
96-h, water-only exposure to cadmium chloride at 20%« and 25°C. The reference toxicant
control for the chronic test is initiated 1 wk after the start of the sediment toxicity test
because the newborn amphipods cannot survive 96-h without sediment or food, having
been released from their mothers' marsupium for less than 1 d. A subset of the newborns
used for the test sediment treatments are placed in culture sediment and fed in the same
manner as amphipods in culture a 1 wk period. After this growing-out period, they are
sieved from the sediment and randomly allocated to the different cadmium concentrations.
The cadmium concentrations for the this control typically range from 0.19-6 mg/L. The
reference toxicant control is also employed for QA/QC, to determine whether the
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D-5
sensitivity of the test animals is consistent among experiments. Only one replicate of
each concentration is used.
The third control is an experimental control in which one non-contaminant
environmental parameter (such as grain size, TOG, temperature, a carrier solvent, etc.) is
allowed to vary from the standard environmental condition to the same extent that this
parameter varies within one or more of the test sediments. This control thus allows for
an estimation of the effect of this non-contaminant parameter on the response of K
plumulosus. This control is treated the same as the other test sediments in all other
ways. These controls are included as the uncontaminated treatment against which the
toxicity of the other test treatments are compared statistically.
Exposure condition modifications which may affect the test results
1. Temperature. Growth and fertility will decrease as temperature is lowered. Higher
temperatures may increase the magnitude of these responses, but also may stress the
amphipods and so introduce variability. Newborn L/. plumulosus may be very sensitive to
temperature change, and every effort should be made to maintain them at constant
temperature (i.e., 25°C) during all phases of bioassay setup (i.e., release from maternal
brood pouch, isolation from adults, sorting, and transfer to test beakers). Generally, the
test temperature should be 25°C. If another temperature is to be used, one or more
temperature control treatments would be recommended.
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D-6
2. Salinity. Choice of test salinity will generally depend on the necessity to match the
salinity of overlying water to that of the test sediments. Experiments conducted under
control conditions showed no significant difference in survival over 10-28 days in salinities
ranging from 1.5-25%o, and no apparent differences in number of young per female after
28 days when exposed to salinities from 5-32%o (Schlekat et al., 1992). However, the
response of the organism to a toxic substance might vary depending on salinity selected,
particularly metals for which bioavailability changes with salinity. In general, the
salinity of the performance and reference toxicant controls should remain constant (i.e.,
20%c), and this salinity should be used unless there are compelling reasons to do
otherwise. If other salinities are used, a salinity control may be advisable.
3. Physical characteristics of test sedi.ment. In 20-day exposures, Schlekat et al. (1992)
reported no significant differences in survival or number of young per female in
uncontaminated sediments ranging from 98.1% sand to 96.5% silt/clay. However,
Maryland Dept. of the Environment (1991) indicated coarse sediment texture might have
been a factor contributing to mortality in acute sediment tests with this organism.
DeWitt et al (in prep.) reported mortality or growth of sub-adult and newborn L.
plumulosus were not correlated with sediment grain size, total organic carbon content,
sediment water content, or Eh. If there is reason to believe sediment characteristics
might affect the amphipod's response, an experimental control for these factors should be
included.
4. Type of exposure. Some amphipod species have shown increased sensitivity to
toxicants in a static system compared to flow-through or static daily renewal (eg. Word et
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D-7
al 1989). No research has been conducted to compare the sensitivities of K plumulosus
under different water-renewal-rate exposure systems.
5. Amount and type of food provided. 10-d growth is reduced if food is withheld, and
mortality, growth, and reproduction probably would be significantly altered if L,.
plumulosus was not fed during a 28-d exposure. Comparisons of feeding regimes
employed by Schlekat et al. (1992) and DeWitt et al. (in prep.) suggest that omission of
live algae from the diet may lead to substantially reduced production of offspring. It
seems likely that nutrition may have a significant impact on toxicological sensitivity, as is
suggested in recent work by McGee et al (in prep.). At this time, changing the diet from
that described above may substantially affect the repeatability, variability, and magnitude
of the toxicological responses.
6. Age of the amphipods. Newborn juveniles (i.e.,
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D-8
Logistics for Conducting the Test
The test is initiated with juveniles which are <24 hours old. Juveniles are
harvested within one day of their release from the brood pouch; their exact age is known,
and their size is essentially uniform. Measurement of an initial sample of juveniles added
to the test will confirm the size range.
A large number of ovigerous females are isolated 5 days before the test is due to
start (i.e., day -5), and only a small percentage of these females need to release their
young on day zero to produce an acceptable supply of test organisms. Experience with
some cultured populations has shown that when ovigerous females are isolated without
regard to developmental stage of the brood, a peak release of young will occur after about
5 days. Cultured populations are sieved through a 1.0mm mesh screen to isolate adults.
Ovigerous females are then selected by examining the adults in a culture dish containing
20%e seawater at 25°C. Approximately 1 gravid female should be isolated for each
juvenile needed for the toxicity test. Presence of a brood can easily be detected with the
naked eye. The isolated females are transferred to a holding container with culture
sediment and seawater 25°C and 20%c salinity. The fe'males are fed in the same manner
as amphipods in other culture containers. After 3 days (day -2), the amphipods are sieved
from the culture sediment with a 1.0mm screen using water at the same temperature and
salinity, the females are transferred to a large glass dish at a density of about 300
amphipods/dish with seawater at the test temperature and salinity, and algal food is
added as in culture containers; however, no sediment is added to the dish. Females
placed in the dish should be carefully inspected to insure that no young have been
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D-9
transferred with them. Juveniles released overnight (i.e., by day -1) are collected and set
aside in a culture tub containing culture sediment pre-sieved to 0.25mm; these juveniles
have not been used in the 28-d test to, but might be available as a back-up if necessary or
to establish new cultures. Care must be taken to be sure that all juveniles released are
separated from the females. The females are returned to the sediment-free glass dishes
until the next day. Juveniles released over the next 24 h (i.e., between day -1 and day 0)
are used to start the test; these juveniles will be <24 hours from brood release. There
should be more young produced than gravid females initially isolated. If insufficient
numbers of newborns are produced for an experiment, it is advisable to delay starting the
exposure of some beakers for 1-d until more
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D-10
offspring (i.e., Fl generation) may be produced in each beaker. After 28 d, the contents of
each exposure chamber are sieved through 1.0mm and 0.25mm sieves to recover the
surviving adults (i.e., F0 generation) and their offspring (i.e., Fl generation), respectively.
Some adults may pass through the 1.0mm screen (especially if the treatment retards
growth) and must be separated from the young. The surviving adults are counted, and
then measured, alive or preserved. If preserved, it is recommended that the adults be
relaxed with magnesium chloride, CO2, or other means before preservation to minimize
curling of the dead animal; curling can stretch the animals and lead to larger apparent
size than for animals measured alive. Body length, from the rostrum to the junction
between the abdomen and urosome, may be measured with an optical micrometer or
computer-assisted digital image analyzer. Each surviving adult should be sexed in order
to determine the number of females which is required for estimating fertility. Males may
be identified by a notched palm on the distal segment of the gnathopod or by the presence
of penile papillae ventrally on the abdomen (only visible in preserved animals). Females
are identified by the presence of eggs in the ovaries or brood pouch. Females lack the
notch on the palm of the last segment of the gnathopod and by the presence of oostigites
(brood plates).
Juveniles may be counted at the termination of the exposure, but it is more
convenient to stain, preserve, and count them at a later time. The material retained on
the 0.25mm screen is transferred to a sample jar, stained overnight with a few ml of
concentrated Rose Bengal in seawater, and then preserved for several weeks in 70%
ethanol until resources permit enumeration. Care should be taken not to dilute the 70%
-------
D-ll
ethanol with seawater in the vials by pipetting off all of the stain solution prior to adding
the alcohol.
Endpoints
Mortality is measured as the percent of the P0 L. plumulosus that were not
recovered alive at the end of 28 d. Most of the survivors of the F0 cohort are collected on
the 1.0mm screen on day 28, although some may pass through to be collected on the
0.25mm sieve if growth rate was severely reduced. Mortality should not exceed 10-20%,
as senescence for this species at 25°C is not observed until after 6 wk of age (DeWitt et al.
in prep.). Dead amphipods are not removed from the exposure chambers on a daily basis,
but they are noted as on the daily observation record.
Fertility is measured as the number of female offspring produced per surviving
female in the exposure chamber. The sex ratio of the offspring must be estimated as 1.0
(females to males) since it is not possible, at this time, to sex newborn L. plumulosus.
Thus, the number of female offspring is 0.5 x the number of juveniles collected on the
0.25mm screen. The number of surviving females can be determined directly by sexing
each animal using the morphological characteristics described above.
Size of surviving F0-generation L. plumulosus is measured as the length of the
body (in mm) from the tip of the rostrum to the base of the urosome (posterior end of the
third abdominal segment; a major point of articulation between the abdomen and the
-------
D-12
urosome). Rate of growth may be related to timing of reproduction and thus to population
parameters (Scott and Redmond 1989). All recovered individuals are measured and sexed.
In some treatments, there may not have been sufficient growth and development to allow
for production of young during the test. In these cases, the state of sexual maturity of the
recovered adults may be important.
Acceptability of the Test
There should be no more than 20% mean mortality among the performance-control
replicates during the first 28 days of the test. This criterion is only a suggestion as there
are insufficient data to establish definitive criteria for any of the responses.
Statistics
The final statistical endpoint is the MATC (Maximum Acceptable Toxicant
Concentration) range for the most sensitive endpoint. The upper bound of the MATC
range is the lowest concentration that shows a statistically significant effect (=Lowest
Observed Effect Concentration = LOEC), and the lower boundary is the highest
concentration that shows no statistically significant effect (=Highest No Observed Effect
Concentration = NOEC). If an analysis of variance (or its nonparametric equivalent)
shows no significant results, then the NOEC is the highest concentration included in the
analysis. (References: Gelber et al. (1985), Weber et al. (1988; appendices), Capizzi et al.
(1985), Daniel (1978), Sokal and Rohlf (1981))
-------
D-13
If initial examination of the data indicates that there may be significant lethal or
sub-lethal effects within treatments, the following analyses are conducted:
1) Tests (Shapiro-Wilks, Bartlett's) are conducted to establish if the assumptions of
normality and homogeneity of variances are met.
2) Arc sin square root transformation of the proportional mortality data is
conducted to stabilize the variance and more closely approximate a normal
distribution if the data are found to be non-normal or the variances non-
homogeneous. Square root or log transformation of the size or fertility data may
be required for the same reasons. Re-test for normality and homogeneity of
variance.
3) If parametric assumptions are met, an analysis of variance (ANOVA) is
conducted, followed by Dunnett's procedure if the ANOVA shows a significant
result. If parametric assumptions are not met, Steel's Many-One Rank Test is
used to compare treatments with the control. This test does require equal
variances, but is fairly insensitive to such deviations from homogeneity. Another
non-parametric analysis that may be conducted is the Kruskal-Wallis test which is
analogous to ANOVA, and is used to determine whether significant differences
exist among treatments, followed by a non-parametric multiple-comparisons
procedure, such as Dunn's test. Remember to test treatment means against the
experimental control (i.e., carrier control, reference sediment, site control, salinity
or temperature control, etc.), not the performance control (i.e., culture sediment).
-------
D-14
CULTURE METHOD SUMMARY
Physical requirements
Tubs: 12" x 14" x 6" plastic dishpans
Salinity: 20%c
Temperature: 20°C
Photoperiod: 16hr light : 8hr dark (fluorescent)
Lights: Fluorescent, ceiling mounted; Natural, N. facing skylight
Sediment: <1 cm layer; mud or muddy-sand sieved <0.5mm; 1-3% TOC
Water: Seawater diluted with deionized water; 10-12 cm layer
Water Change: Static-renewal: 50% vol. water change 3x/wk
Aeration: Constant bubbling
Density: 300-400 adults/tub (= 0.3-0.4 adults/cm2)
Feeding
3x/wk, at time of water change
Algal mixture: -7-L per culture tub
Pseudoisochrysis paradoxa (chrysophyte)
Phaeodactylum tricornutum (diatom)
1:1 v/v mixture, final cone. 106cells/ml, 20%e
Gorp: dry food mixture; fine powder; 0.5 g per tub
48.5% TetraMin®
24% dried alfalfa
24% dried wheat leaves
4.5% Neo-Novum® (shrimp maturation feed; Argent Chem. Lab.)
Culture Renewal
Start culture with ca. 100 adults and 200 juveniles
Thin cultures every 6-8 wk
Inspect culture sediment for worms, copepods; discard if present
Replace sediment at least every 6 mo.
Special Considerations
Non-indigenous Species Laws and Practices
Chlorination or sterilization (autoclave) all materials used
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D-15
Leptocheirus plumulosus Chronic Sediment Toxicity Test Design: SUMMARY
Exposure Conditions
Chamber:
Sediment Vol.:
Overlying Water Vol.
Water Source:
Salinity:
Aeration:
Temperature:
Photoperiod:
Duration:
Life stage:
No./replicate:
Feeding:
Performance Control:
Ref. Tox. Control:
1-L glass beakers
175ml
725ml
Seawater diluted with deionized water
20% or match test sediment IW salinity
Constant
25°C
16hr light : 8hr dark
28 days
<24-hr old newborn (F0-generation)
20
3x/wk, 400ml algae @ 106cells/ml; 10 mg gorp
28-d, culture sediment, 20%o, 25°C; <24-hr old
96-h, Cd in water; 20V, 25°C; 1-wk old amphipods (F0)
Handling & Recovery
Obtaining Test Animals
Isolate newborn (F0) from gravid females in dishes w/out sediment
Isolate gravid females from cultures 5d before T0
Maintain constant 25°C temperature while handling
Seeding Exposure Chambers
Transfer by pipette; newborns very fragile
Double or triple count at T0
Preserve >1 subsets of 20 newborns
Size at Tn
Recovery at Test Termination
1.0mm sieve - F0-generation (adults)
Count, measure, sex survivors
0.25mm sieve - Fj-generation (offspring)
Stain & preserve, count under magnification
-------
APPENDIX E
AMPELISCA ABDITA: GENERIC LIFE CYCLE TEST DESIGN
Abstract
A generic life cycle sediment toxicity test with the amphipod Ampelisca abdita is
described. This is a draft design which has not been fully tested, and the procedures
outlined have not always produced high survival or reproduction under control conditions.
This is a flow-through test conducted at 20 - 25°C, 30%o, and 16 hours light and 8 hours
dark. Test sediment is mud to sandy mud, and algal food is delivered daily with
seawater. The experiment begins with 10 juveniles, 8 to 10 days old, per replicate, 5
replicates per treatment, and is terminated after 35 days when the contents of each
container are sieved and examined. Endpoints are initial mortality, time to first observed
juvenile tubes (brood release), number of surviving young produced per female, number
and size of animals recovered, and life stage of animals recovered.
Exposure Conditions
Test treatments, typically 5 concentrations plus negative (uncontaminated
sediment) and positive (reference toxicant) controls, are based on acute toxicity test data.
Treatments can be "spiked" sediment, field sediment, or dilutions of field sediments. If
E-l
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E-2
there are sufficient personnel available, the test can be set up "blind," i.e., so that
personnel monitoring the test have no knowledge of the treatments in the test chambers.
The recommended temperature is 20°C, since it has been shown to be an
acceptable culture and testing temperature for this species. The life cycle is slightly
shorter at 25°C vs. 20°C, but it has not been definitively established that the higher
temperature is acceptable for A. abdita. This organism has been maintained and tested
at salinities within the range of 28 - 35%o.
A light cycle of 16 hours light and 8 hours dark was selected to approximate
conditions existing during summer breeding. This photoperiod has been shown to
consistently maintain reproductive activity in Hyalella azteca (de March 1977, cited in
Arthur 1980). Feeding (Mills 1967) appears not to be affected by the laboratory light cycle.
In nature, this species is found in fine sand to silt without shell, is abundant where
the major proportion of sediment is >0.05 mm and <0.25mm, and may also be found in
coarse sediments with a considerable fine fraction (Mills 1967). Test sediments should
approximate these characteristics, i.e., mud or sandy mud.
The exposure container used is an aerated 900 ml glass "canning" jar, which was
selected as a smaller version of the gallon "pickle jar" used for maintaining these
amphipods in the laboratory and for previous chronic tests (Scott and Redmond 1989); it
is inexpensive and easily drilled. A 250 um screened overflow hole prevents escape of
juvenile amphipods during a flow-through test. This exposure container has a reasonably
-------
E-3
high water column, allowing the animals to swim during their mating activities, and
sloping "shoulders" to allow for efficient circulation of suspended food material. Mills
(1967) indicates that turbidity and currents are likely stimulants for feeding in A. abdita.
Sediment depth in the exposure container is 3.5-4 cm, which is the maximum tube length
of A. abdita.
A flow-through system is used as a closer approximation to a natural situation
than a static system. The number of volume replacements per day needed has not been
determined. Algal food is delivered with the seawater so as to ensure an even distribution
to all exposure containers. The amount added is measured by cells/ml in the culture used,
and the delivery rate to each exposure container.
Exposure condition modifications which may affect the test results
1. Temperature. If a temperature other than 20°C is used, the duration of the test will
change (e.g., at 25°C the duration would be <35 d) and the response of A. abdita to
chemicals might change.
2. Salinity. Choice of test salinity will generally depend on the seawater supply available
to a laboratory, and whether or not it is necessary to match the salinity to that of the test
sediments. It may be possible to acclimate A. abdita to lower salinities, since it is reported
in the literature down to 10%c (Bousfield 1973), but successful acclimation to low salinities
-------
E-4
has not been demonstrated experimentally. The response of the organism might vary
depending on salinity selected.
3. Particle size of test sediment. This species can be tested in coarser materials for acute
tests, but some preliminary information indicates that sandy sediment could result in
adverse effects when exposure is for longer periods of time (Redmond and Scott
unpublished). If sediment characteristics might affect the amphipod's response, a grain
size control should be included.
4. Type of exposure. This species has sometimes shown increased sensitivity to toxicants
in a static system compared to flow-through or static daily renewal (e.g., Word et al 1989).
5. Amount and type of food provided: The quality and quantity could affect growth rate,
fecundity, and exposure to bedded contaminants.
Biological Design
A schedule for conducting a typical A. abdita life cycle sediment toxicity test is
shown in Table E-l. The test is initiated with juveniles which are 8 to 10 days old.
Newly-released juveniles are collected in containers with no sediment from ovigerous
females carrying broods in a late stage of development. Late-stage broods can easily be
identified under a dissecting scope. If juveniles are harvested immediately after release,
their exact age and size are known, since newly-released juveniles are essentially uniform
-------
E-5
in size at about 1.5 mm in length (Mills 1967, also this report). A large number of
ovigerous females are isolated before the test is due to start. Only a small percentage of
these females need to release their young overnight to produce an acceptable supply of
test animals. Juveniles collected with this "no-sediment" procedure are transferred to
aerated containers with a small amount of sediment and held, with feeding, for 8 to 10
days. With this procedure, questions of natural mortality (which has not been
quantified) and initial mortality due to release of the young in no sediment are
eliminated, but age of the organisms is still known. Also, older juveniles are easier to
work with than newly-released individuals, e.g., daily mortalities are more easily
observed. If it is not possible to obtain enough newly-produced young overnight, young
may be collected over a period of 1 to 3 days, with only a small variation in test size
range. In either case, measurement of an initial sample of juveniles added to the test will
confirm the size range.
During the course of the test, juveniles added to experimental chambers grow,
molt, reproduce, and die (especially males, i.e., following reproduction). Exposure to
chemical contaminants might alter these responses and their timing relative to the
amphipods' life history patterns in the control beakers. Thus, exposure and control
containers must be checked daily. Sex ratio in Ampelisca abdita populations is
approximately 1:1 at breeding times (Mills 1967). The test begins with 10 juvenile
amphipods. If 50% of the original amphipods are females, and each female produces 10 -
20 young, the approximate number of young produced in the Fj generation in each jar will
be 50 - 100. After 35 days the contents of each container are sieved and the amphipods
-------
E-6
preserved in 70% ethanol with 5% glycerin1 for later examination. Scott and Redmond
(1989) started replicates at 20°C with ovigerous females and observed differences in the
abundance of F2 generation animals after 56 d. This test design starts with week-old
juveniles, thus eliminating 2 wk of development in the brood pouch and 1 wk of growth,
and results in a test with a 5 wk duration.
Endpoints
Several endpoints may be measured in this test (Table E-2). Initial mortality is
measured as the number of juveniles initially added minus the number of the initial
generation recovered. Mortality in the initial generation is expected, since males die after
mating and females die at some point after releasing a brood. Dead amphipods are
removed every day and examined immediately to determine sex and reproductive
condition. By removing dead individuals daily, the timing of their death can be precisely
determined, and important demographic data can be recorded before the bodies
decompose. These demographic data are necessary in order to estimate the sex ratio
within each beaker. Also, mortality of prereproductive animals due to toxicants can be
separated from what may be natural or background mortality. Dead females are
preserved in 70% alcohol and glycerin, and later are measured in order to relate size of
l. Alcohol is less hazardous than formalin but may not be an adequate
preservative for storage of specimens longer than several weeks; buffered formalin
is better. Glycerin is added to prevent specimens from becoming brittle. Seawater
transferred with the amphipods should be removed because it will dilute the
alcohol and salt crystals may precipitate.
-------
E-7
females to the number of young produced. Time to brood release is determined by the
first appearance of juvenile tubes in each exposure chamber.
After final sieving and preservation, the young in each replicate are counted. The
number of young is divided by the number of females that have produced young
(determined by removal of dead animals as described above, and by examination of sieved
and recovered individuals), to produce a mean number of surviving young per female per
replicate. The number of juvenile tubes observed in each exposure jar are also counted.
The utility of counting the number of tubes observed is that high mortality in the F,
juveniles may be observed. For example, if 20 juvenile tubes are observed during the
course of the test, but at sieving only 5 young are recovered, then there has been an acute
effect on the juveniles in that container. The number that died cannot be quantified
accurately since the small tubes are difficult to count and juveniles may construct >1
tube, but the presence of a large effect can be observed.
Size of recovered individuals is determined as the length in mm, from the base of
the first antenna to the base of the telson. All recovered individuals are measured and
sexed. In some treatments, there may not be sufficient growth and development to allow
for production of young during the test. In these cases, the state of sexual maturity of the
recovered adults is particularly important. When ovigerous females are recovered, they
are preserved individually. Females often release their eggs in preservative, and
individual preservation allows an accurate determination of the number of eggs carried by
a particular female. The number of eggs can then be directly related to the length of that
-------
E-
individual. Growth rate is related to timing of reproduction and thus to population
parameters (Scott and Redmond 1989).
Acceptability of the Test
There must be no more than 10% mean mortality in prereproductive control
individuals during the first 10 days of the test. (There is insufficient data to support
control mortality criteria for longer time periods.) Also, the controls should develop,
mature, and reproduce normally, which at this point in the test development is a
qualitative judgement made by the researcher.
Statistics
The final statistical endpoint is the MATC (Maximum Acceptable Toxicant
Concentration) range for the most sensitive endpoint. The upper bound of the MATC
range is the lowest concentration that shows a statistically significant effect (=Lowest
Observed Effect Concentration = LOEC), and the lower boundary is the highest
concentration that shows no statistically significant effect (=Highest No Observed Effect
Concentration = NOEC). If an analysis of variance (or its nonparametric equivalent)
shows no significant results, then the NOEC is the highest concentration included in the
analysis. (References: Gelber et al. (1985), Weber et al. (1988; appendices), Capizzi et al.
(1985), Daniel (1978), Sokal and Rohlf (1981)).
-------
E-9
If initial examination of the data indicates that there may be significant lethal or
sub-lethal effects within treatments, the following analyses are conducted:
1) Tests (Shapiro-Wilks, Bartlett's) are conducted to establish if the assumptions of
normality and homogeneity of variances are met.
2) Arc sin square root transformation may be applied to the proportional mortality
data. This transformation may be required in some applications as a general
practice, but in other cases, it might only be used (e.g., to stabilize variances and
more closely approximate a normal distribution) when the data are found to be
non-normal or the variances non-homogeneous. Square root or log transformation
of the size or fertility data may be required for the same reasons. Re-test for
normality and homogeneity of variance.
3) If parametric assumptions are met, an analysis of variance (ANOVA) is
conducted, followed by Dunnett's procedure if the ANOVA shows a significant
result. If parametric assumptions are not met, Steel's Many-One Rank Test is
used to compare treatments with the control. This test does require equal
variances, but is fairly insensitive to such deviations from homogeneity. Another
non-parametric analysis that may be conducted is the Kruskal-Wallis test which is
analogous to ANOVA, and is used to determine whether significant differences
exist among treatments, followed by a non-parametric multiple-comparisons
procedure, such as Dunn's test. Remember to test treatment means against the
-------
E-10
experimental control (i.e., carrier control, reference sediment, site control, salinity
or temperature control, etc.), not the performance control (i.e., culture sediment).
Survival
Initial evaluation of data involves determination of effects on survival. Survival in
each treatment can be graphically represented. Figure E-l depicts the survival curve of
A. abdita under control conditions: low mortality over the first 2-wk (i.e., ca. <10% after
10-d), followed by increasing mortality due to the death of post-reproductive amphipods.
If there is significant mortality of i ndividuals in some treatments which is not due to
natural (post reproductive) mortality, these treatments should be removed from further
analysis of "sublethal" effects.
Other parameters
Time to first appearance of juvenile tubes in test-days, and length of young
produced are analyzed as described above. The number of young produced is divided by
the number of females present which have released a brood (determined by the number of
amphipods initially added and the daily observations), and analyzed as above except that
an analysis of covariance is used, with female length the covariate.
-------
E-ll
SUN MON TUES WED THURS FRI SAT
isolate collect
WEEK 1 ovigerous juveniles
females
sediments amphipods
into test are 8-10
WEEK 2 containers days old;
time-zero
Table E-l. A. abdita: example of organism collection schedule for life cycle test.
-------
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Figure E-l. Mortality of A. abdita under typical control-exposure conditions.
-------
R-l
REFERENCES
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American Society for Testing and Materials. 1990b. Standard guide for conducting solid-
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Boesch, D.F. 1973. Classification and community structure of macrobenthos in the Hampton
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Boesch, D.F. 1977. A new look at the distribution of benthos along the estuarine gradient. In:
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Botton, M.L. 1979. Effects of sewage sludge on the benthic invertebrate community of the
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Capizzi, T., L. Oppenheimer, H. Mehta, H. Naimie, and J.L. Fair. 1985. Statistical
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