EPA 910/9-90-011
Puaet Sound	Estu
Protocol for
JUVENILE NEANTHES
SEDIMENT BIOASSAY
June 1990

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PTI Environmental Services
15375 SE 30th Place
Suite 250
Bellevue, Washington 98007
PROTOCOL FOR JUVENILE
NEANTHES SEDIMENT BIOASSAY
By
D. Michael Johns, Thomas C. Qinn, and Donald J. Reish'
)
For
U.S. Environmental Protection Agency
Region 10, Office of Puget Sound
1200 Sixth Avenue
Seattle, WA 98101
EPA Contract 68-D8-0085
PTI Contract C744-11
June 1990
* California State University, Long Beach, California

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CONTENTS
Eags
LIST OF FIGURES	iii
LIST OF TABLES	iii
NEANTHES SUBLETHAL BIOASSAY	1
OVERVIEW	1
BACKGROUND	1
INTRODUCTION	2
Species Sensitivity	3
Ecological Importance	3
USE AND LIMITATIONS	3
FIELD PROCEDURES	5
Collection	5
Processing	5
LABORATORY PROCEDURES	5
Test Animals	5
Control and Reference Sediments	7
Test Sediments	8
Bioassay Seawater	8
Bioassay Procedure	8
Experimental Design	12
DATA REPORTING REQUIREMENTS AND STATISTICAL ANALYSIS	13
REFERENCES	15

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UST OF FIGURES
Eage
Figure 1. Static exposure system used for the Neanthes sublethal bioassay	9
UST OF TABLES
Table 1. Sensitivity of Neanthes to various contaminants
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NEANTHES SUBLETHAL BIOASSAY
OVERVIEW
This protocol is for conducting a bioassay in which the survival and change in biomass
of juvenile Neanthes sp. are determined following a 20-day exposure to test sediments.
Parameters measured to determine the effects of exposure include mortality, total biomass,
and average individual biomass. Sediments can be either naturally occurring, field-collected
samples, or sediments that have been experimentally modified (e.g., sediment mixed with
other sediment to form a gradient of sediment types or sediment to which chemicals have
been added). The Neanthes bioassay is conducted as a static renewal exposure, and food
(i.e., TetraMarin®) is provided to the test organisms during the exposure period to promote
body tissue increases. Following the 20-day exposure period, all surviving worms are
collected, dried to a constant weight, and total and average individual biomass are determined.
BACKGROUND
The primary basis for this final protocol is a sublethal test demonstration study (Johns
1988) conducted for the Seattle District of the U.S. Army Corps of Engineers (Corps) and the
Puget Sound Dredged Disposal Analysis (PSDDA) study. The testing procedures also
incorporated information from the published literature and the testing approach developed
by the Los Angeles District Corps for evaluating the acute toxicity of sediments to Neanthes.
Following development of a draft protocol, the Washington Department of Ecology conducted
an experts workshop on the development of a Neanthes bioassay to be used as part of the
state's marine sediments management program. The general objectives of the workshop were
to evaluate the draft protocol for conducting lethal and sublethal sediment bioassays using
Neanthes, and to determine the information and research that may be needed for further test
development.
As part of the workshop, the experts were asked to categorize and rank the information
and research needs obtained during the review of the draft protocol in order to provide
guidance to the Washington Department of Ecology on suggested changes to the protocol.
Four categories of recommendations were made and each information need or research topic
identified during the workshop was assigned to one of the categories. One category included
recommended changes that could be incorporated into the draft protocol without the need
for further testing or research. Items fitting into this category include those for which sufficient
data have already been generated, or for which the experts felt a decision could be made
based on current knowledge and experience. An interim protocol was then developed that
was based on these workshop recommendations.
The other three categories of recommendations were concerned with research topics that
might be addressed to further develop the Neanthes sublethal bioassay. High priority research
topics identified at the workshop were addressed in work funded by the U.S. Environmental
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Protection Agency, Region 10 (Johns and Ginn 1990). Results from the research by Johns
and Ginn (1990) were incorporated into the interim protocol to further enhance and extend
the usefulness of the test and to produce this final protocol. This protocol is now ready for
widespread application and evaluation.
Research and other development documents relied upon in establishing this protocol
include:
Johns, D.M. 1988. Sublethal test demonstration. Prepared for U.S. Army
Corps of Engineers, Seattle District, for the Puget Sound Dredged Disposal
Analysis. Submitted to E.V.S. Consultants, Seattle, WA. PTI Environmental
Services, Bellevue, WA. 94 pp. + appendices.
Johns, D.M., T.C. Ginn, and J.E. Sexton. 1989. Evaluation of growth as an
indicator of toxicity in marine organisms. Prepared for the Washington
Department of Ecology, Olympia, WA. PTI Environmental Services,
Bellevue, WA. 28 pp. + appendices.
Johns, D.M., T.C. Ginn, and D.J. Reish. 1989. Interim protocol for juvenile
Neanthes bioassay. Prepared for the Washington Department of Ecology,
Olympia, WA. PTI Environmental Services, Bellevue, WA. 14 pp. +
appendices.
Johns, D.M., and T.C. Ginn. 1989. Test demonstration of a 10-day Neanthes
acute toxicity bioassay. Prepared for U.S. Army Corps of Engineers, Seattle
District. PTI Environmental Services, Bellevue, WA. 16 pp. + appendices.
Johns, D.M., and T.C. Ginn. 1990. Development of a Neanthes sediment bio-
assay for use in Puget Sound. Prepared for U.S. Environmental Protection
Agency Region 10, Office of Puget Sound, Seattle, WA. EPA 910/9-90-005.
PTI Environmental Services, Bellevue, WA. 51 pp. + appendices.
Johns, D.M., and T.C. Ginn. 1990. Neanthes long-term exposure experiment:
the relationship between juvenile growth and reproductive success. Pre-
pared for U.S. Environmental Protection Agency Region 10, Office of Puget
Sound, Seattle, WA. EPA 910/9-90-010. PTI Environmental Services,
Bellevue, WA. 15 pp.
Johns, D.M., R.A. Pastorok, and T.C. Ginn. 1990. A sublethal sediment toxicity
test using juvenile Neanthes sp. (Polychaeta: Nereidae). Submitted. In:
Aquatic Toxicity and Hazard Assessment: Fourteenth Symposium.
American Society for Testing and Materials, Philadelphia, PA.
INTRODUCTION
Neanthes sp., a marine nereid polychaete, is widely distributed throughout the world, and
has been collected in New England, Florida, California, Europe, and the central Pacific Ocean
(Reish 1980). Laboratory cultures of Neanthes have been successfully maintained since 1964.
Pesch et al. (1988) reported a difference in chromosome numbers from two populations
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(collected in Connecticut and California, respectively) of Neanthes. In addition to the
differences observed in chromosome numbers, differences were also noted in the morphology
indicating that these populations represent different species. Although specimens from both
populations have been used in testing, almost all the testing data are associated with
experiments conducted with specimens from the California population. The procedures
discussed in this protocol are for Neanthes originating from the California population.
Since 1966, various life stages of Neanthes have been used as bioassay organisms for
a wide variety of investigations including evaluating the effects of dissolved oxygen concentra-
tions, nutrients, salinity, temperature, metals, pesticides, hydrocarbons, and contaminated
sediments on survival, growth, and reproduction. In addition, Neanthes has also been used
to investigate the effects of mutagens (Pesch et al. 1981; Pesch and Pesch 1980) and
irradiation (Jones et al. 1983) on marine organisms; an interlaboratory comparison has been
conducted with a Neanthes 28-day flow-through seawater toxicity test (Pesch and Hoffman
1983); and a 96-hour acute sediment bioassay using Neanthes is currently being used for
dredged material testing by the U.S. Army Corps of Engineers, Los Angeles District (Reish
and Lemay 1988).
Species Sensitivity
Neanthes has been used to evaluate the toxicity of a wide variety of contaminants
including metals, hydrocarbons, and multicontaminated media (i.e., sediments). Examples
of the types of toxicity tests conducted with Neanthes and species sensitivity are presented
in Table 1.
Reish (1984) summarized data on the sensitivity of Neanthes to metals. In comparison
to other polychaetes, Neanthes appears to be moderately sensitive to most metals tested.
Studies indicate that mercury and copper are the most toxic to Neanthes, followed by
aluminum, cadmium, chromium, zinc, lead, and nickel.
Ecological Importance
Neanthes is distributed on the west coast from Mexico to southern California (Reish 1980).
Neanthes has not been collected from Puget Sound. The family Nereidae is widely distributed
and is a dominant taxa in intertidal and subtidal habitats. In Puget Sound, the nereid
Platynereis bicanaliculata is a dominant member of the polychaete fauna at many sites (Lie
1968; PTI and Tetra Tech 1988a,b). P. bicanaliculata is morphologically similar to Neanthes
in jaw structure and is also recorded to be an omnivore, feeding on algae and other detritus
(Fauchald and Jumars 1979). Both species also build similar tubes of organic material and
display similar aggressive behavior patterns (Gray 1974).
USE AND LIMITATIONS
The Neanthes sublethal bioassay is used to characterize the toxicity of marine sediments
based on worm survival and growth. Data reported by Johns and Ginn (1990) indicate that
the level of contamination affecting juvenile growth in Neanthes is similar to the level of
contamination that affects reproductive success. The bioassay may be used alone (e.g., as
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TABLE 1. SENSITIVITY OF NEANTHES TO VARIOUS CONTAMINANTS
Lowest
Concentration for
Contaminant
Observed Effect
(mg/L)
Endpoint
Test
Duration
Reference
Aluminum
>2.0
Mortality
4 days
Petrich and Reish (1979)
Cadmium (as CdClg)
3
1
Mortality
Reproduction
28 days
Life cycle
Reish (1980)
Reish and Gerlinger (1984)
Chromium (as CrOg)
0.6
Mortality
28 days
Reish (1980)
Hexavalent chromium
(as K^C^Oj)
0.0125
Reproduction
Life cycle
Oshida (1976)
Copper
0.1
Mortality
29 days
Pesch and Morgan (1978)
Lead [as Pb(CH3CO)2]
3.2
0.97
Mortality
Reproduction
28 days
Life cycle
Reish (1980)
Reish and Gerlinger (1984)
Mercury (as HgClg)
0.17
Mortality
28 days
Reish (1980)
Nickel
_ 49.0
Mortality
4 days
Petrich and Reish (1979)
Silver (as AgNO-j)
0.165
Mortality
28 days
Pesch and Hoffman (1983)
Zinc (as ZnSO^
1.4
0.32
Mortality
Reproduction
28 days
Life cycle
Reish (1980)
Reish and Gerlinger (1984)
DDT
0.1
Mortality
28 days
Reish (1985)
No. 2 fuel oil
2.7
Mortality
4 days
Rossi and Anderson (1976)
South Louisiana crude oil
12.5
Mortality
4 days
Rossi and Anderson (1976)
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a screening tool in broad-scale sediment surveys), in combination with sediment chemistry
and in situ biological indices, and in laboratory experiments to address various sediment and
water quality manipulations. The following constraints apply:
¦	The bioassay should be conducted with laboratory-cultured juvenile Neanthes
¦	Modification of the protocol may be required for tests conducted at salinities
(both interstitial and overlying water) less than 20 ppt.
FIELD PROCEDURES
Collection
Test Animals—Neamhes are not indigenous to Puget Sound and test organisms must
be obtained from laboratory cultures. (See Laboratory Procedures section for a discussion
on culturing and obtaining test organisms.)
Sediment—Control, reference and test sediments should be collected in solvent-cleaned
glass containers having teflon-lined lids. Each jar should be filled completely to exclude air.
A minimum sediment sample size of 0.25 liters for each bioassay chamber is recommended
for all sediment types.
Processing
Test Animals—Not applicable.
Sediment—All sediments should be stored at 4°C in the dark. Holding time should not
exceed 14 days.
LABORATORY PROCEDURES
Test Animals
Culturing—Almost all Neanthes used for laboratory tests come from laboratory cultures.
Culturing techniques have been described by Reish (1980) and Pesch and Schauer (1988).
Under laboratory conditions, the Neanthes life cycle is completed in 3-4 months at 20-22°C.
Cultures of adult Neanthes are maintained in glass aquaria under static (with monthly
renewal) or flow-through water conditions. After sexually mature males and females pair, the
pairs can be isolated in jars and maintained until juveniles are ready to be removed and used
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for testing. Eggs are laid within the worm tube, and the female dies within 2-3 days. The
zygotes are cared for by the surviving male. Larvae emerge from the worm tube in approxi-
mately 3 weeks (at 20-22°C) following fertilization. Hatched larvae feed on yolk reserves until
emergence from the adult worm tube. Following emergence, the juvenile worms are capable
of feeding and building independent tubes. Until testing, the juvenile worms are maintained
without sediment and are provided TetraMarin® (a food source) and powdered alga (either
Enteromorpha or Ulva sp.). Enough powdered alga (sieved to less than 0.3 mm) should be
provided to cover the bottom of the aquarium. The powdered alga provides material for tube
construction and increases survival (Pesch and Schauer 1988).
Shipping and Holding-Juvenile Neanthes are obtained from laboratory cultures. If test
organisms are obtained from an outside source, enough time should be allotted to allow the
worms to acclimate prior to starting a test. Neanthes can be shipped by overnight courier
without significant mortality. Worms are typically packed in plastic bags containing seawater
with 50 organisms per bag. Each bag should contain several fronds of dried Enteromorpha.
This alga can be collected, dried, and stored for extended periods. Prior to use, the alga
should be soaked in seawater. The bags are shipped in a hard-sided container (e.g.,
cardboard box). When the shipment arrives at the laboratory, the worms, still in the plastic
bags, are placed in a holding aquarium containing seawater at the proper test temperature.
The worms are released from the bags after temperature equilibration. The worms are
maintained in the holding aquarium for 1 -2 days prior to initiation of the bioassays. The
holding time will provide for acclimation between the culture temperature and the anticipated
testing temperature and for observation of the condition of the test organisms to ensure that
the bioassay is conducted with healthy individuals.
Neanthes juveniles should be held in all-glass aquaria containing clean seawater and
provided with gentle aeration [see Pesch and Schauer (1988) if flowing seawater is available].
Water temperature is maintained at 20 ± 1 °C, and salinity is maintained at the salinity at which
the bioassay will be conducted. Enough powdered green alga (Enteromorpha or Ulva sp.)
should be provided to cover the bottom of the holding tank.
During the holding period, organisms are provided with TetraMarin® on an every-other-day
basis. The amount of food provided should be calculated at approximately 8 mg (dry weight)
per juvenile, but the tank should be observed following feeding to determine if the food is
being consumed. If it is not being consumed, then the amount of food provided should be
reduced in order to avoid potential water fouling problems. If the entire amount of food
provided is being eaten, then an increase in the food ration might be appropriate.
No water changes in the holding tank are required if the worms are being maintained in
the aquaria for less than 1 week. If the worms are to be maintained for a longer period, then
the water should be replaced with fresh seawater once every 2 weeks. Rising salinity, due
to evaporative losses during the holding period, can be compensated for by adding sufficient
distilled water to lower the salinity to the desired level.
Test Animal Size—The size of juvenile worms used in the bioassays Is potentially a critical
factor to the eventual success of the bioassay. Worms should be 0.5-1.0 mg (dry weight) (i.e.,
2-3 weeks post-emergence) to ensure that they are in a rapid growth phase during the
exposure period. Worms of this age are large enough to be easily handled to avoid errors
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in placing the correct number of worms in each exposure chamber. For consistency in aging
test organisms, initiation of emergence should be considered as the point when feeding
juveniles emerge from the egg case. Commencement of feeding can be identified by the
presence of food particles in the digestive tract.
Feeding Requirements—Several different types of food have been used in culturing
Neanthes, including alfalfa flour, powdered alga {Enteromorpha or Ulva sp.), TetraMarin®, and
prawn flakes. Of these foods, prawn flakes and TetraMarin* appear to provide the best and
most consistent growth throughout the life cycle. Because of potential problems in obtaining
a consistent supply of prawn flakes, TetraMarin* should be used. TetraMarin* should be pro-
vided to juveniles maintained in holding tanks prior to testing and during the exposure period.
In both cases, the worms should be fed on an every-other-day basis. The amount of food
provided should be calculated at approximately 8 mg (dry weight) per juvenile Neanthes.
Control and Reference Sediments
In addition to exposure chambers containing test sediments, exposure chambers contain-
ing control and reference sediments are also prepared. Control sediment is typically collected
from the site at which the test organism is found or the substrate in which the organism is
cultured. The sediment provides a nontoxic sediment for evaluation of the condition of the
test organisms being used in the bioassay. For the Neanthes bioassay, sand should be used
as the control sediment.
Sand was initially chosen as an appropriate control sediment based on the work of Pesch
and Hoffman (1982), who used sand as a substrate in a series of experiments with Neanthes.
They reported no significant mortality associated with maintaining the worms in sand. For
the sublethal bioassay test demonstration study (Johns 1988) and subsequent testing (Johns
and Ginn 1990), sand collected from West Beach on Whidbey Island, Washington, was used
as the control sediment. Neanthes maintained in West Beach sand exhibited low mortality
and high percentage increases in biomass during the exposure period, indicating that West
Beach sand is a suitable material for a control sediment. In addition, West Beach sand was
selected because it was used as a control sediment for a number of the regulatory bioassays
conducted in Puget Sound and is known to be relatively free of contaminants.
Because control sediments may differ greatly from the test sediments with respect to
physical and chemical sediment characteristics (e.g., grain size and organic content), a
reference sediment is also included in the bioassay series. Data from the reference sediment
can be used to partition contaminant effects associated with a test sediment from those relat-
ing to the physical and chemical characteristics of the test sediment. Johns and Ginn (1990)
evaluated the influence of sediment grain size on Neanthes survival and growth following
exposure to sediments having differing granulometry (expressed as a percentage of the silt/
clay fraction in the sediment). The results of this experiment indicate that Neanthes are able
to survive and grow in a wide range of sediment types. Johns and Ginn (1990) also noted
that statistical differences in growth could occasionally be detected in Neanthes exposed to
widely differing sediment types, and cautioned that reference sediment used in Neanthes bio-
assays should have a similar grain size and organic content as the test sediments to avoid
potential differences in organism response related to the physical characteristics of the
sediment.
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Test Sediments
The natural geochemical properties of test sediment collected from the field must be within
the tolerance limits of the test species. Johns and Ginn (1990) determined the 96-hour LC^
for Neanthes exposed to seawater of different salinity to be 15 ppt. Caution should be used
when performing and interpreting the results of Neanthes bioassays conducted with sediments
with an interstitial salinity of less than 20 ppt. Modification to the test sediment (e.g., mixing
the sediment with high salinity water to raise interstitial salinity) or test protocol (e.g., use of
high salinity seawater in the exposure chamber) might be considered when testing sediment
collected from low salinity areas.
Bioassay Seawater
Seawater used in the bioassay should be maintained at a salinity of 28 ± 2 ppt and at
a temperature of 20 ± 1°C. If a series of experiments is planned, then the test temperature
and salinity should be the same throughout the series. The bioassay seawater must be
uncontaminated.
Bioassay chambers are 1-L glass containers with an internal diameter of approximately
10 cm. The chambers are covered with lids to reduce contamination of the contents and
evaporation of the seawater or loss of volatiles. The bioassay chambers are maintained at
20 ± 1°C in either a shallow waterbath or in a constant-temperature room. Exposure
chambers are gently aerated with air that is free of fumes, oil, and water. This air is delivered
to the exposure chamber by nontoxic tubing with a glass Pasteur pipette suspended 3-4 mm
below the water surface. The aeration rate should be between 150 and 300 mL/minute.
Prior to use, all glassware is thoroughly cleaned, rinsed in distilled water, soaked in a
10 percent nitric acid (HNOg) (or 10 percent hydrochloric acid) solution for 2 hours, and rinsed
with distilled water.
Bioassay Procedure
Overview—The bioassays are conducted using a static renewal exposure system. Each
exposure chamber consists of a 1-L jar containing 2 cm of sediment and seawater (Figure 1).
Prior to testing, all exposure chambers are cleaned and rinsed in turn with distilled water,
10 percent nitric acid (HNO,), and distilled water.
At the beginning of each test, five juvenile worms are randomly placed into each exposure
chamber. During setup, three subsamples of worms (five worms per subsample) are randomly
selected to provide an estimate of initial worm biomass.
During the exposure period, each exposure chamber is provided with 40 mg of food (i.e.,
8 mg per individual) on an every-other-day basis. Every third day, one-third of the seawater
in each exposure chamber is exchanged with fresh seawater. Water quality measurements
taken during the exposure period include dissolved oxygen concentration, salinity, and pH.
These measurements are made for each exposure chamber just prior to the seawater
exchange.
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Air Supply
Water
r V ;¦ V .*• V .*• V ;• V ;¦ V ;• V ;¦ V ;¦ V ;• V ;• V ;• V ;• V .*• V ;¦ V ;¦ V ;• V ;• V Vvv •• v ••'
• ' • : ¦ . •,. •.: •,: • : • .• • .• ¦ : • : • .• • : • .* • .• • ; ¦ .¦ ¦ : • : ¦. ,* • .• •
*. •• *. • •. • *. • \ ¦ *. ¦. •. • •.. •. •. • ' \ ' •. •. *.. •.. •. •: • ,* • .* ¦ ; •
r • v • v •'.* »v ¦ .* .*.• .*.~	.*.• .*.• ,
«v ••: • .¦ • ,* • ,* • ,• . ,• . ,~.; . .•. .•.•.•
¦Test Sediment
•V

Figure 1. Static exposure system used for the Neanthes sublethal bioassay

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Following the exposure period, the contents of each replicate chamber are sieved through
a 0.5-mm screen and the number of living worms is recorded. Surviving worms are then
placed in a vial containing clean seawater. After all chambers have been sieved, the surviving
worms in each vial are quickly rinsed with an ammonium formate solution, placed on a
preweighed aluminum pan, and dried at 50°C to a constant weight. Total weights are then
determined to the nearest 0.1 mg.
Initiation—Prior to initiation of a bioassay, all exposure chambers are cleaned as described
above, and test organisms are acclimated. On the day before test initiation, test sediments
are placed in each of the five replicate exposure chambers. Each chamber should be filled
so that a 2-cm sediment layer is formed in the bottom of the chamber. Sediment placed in
the chamber is smoothed by tapping the jar against the palm of the hand. Once the sediment
is smoothed, the chamber is filled with seawater by gently pouring the water down the side
of the chamber. Filled chambers are placed in a 20 ± 1°C waterbath and capped. An air
line is inserted through a hole in the cap. The exposure chambers are allowed to equilibrate
overnight to bioassay conditions. The photoperiod during testing should be continuous, using
ambient light of low to moderate intensity. Although the intensity does not have to be
measured, light levels should be similar to that obtained from fluorescent or incandescent
light sources that are not placed directly over the water bath.
On the day of test initiation, juvenile worms are collected from the holding tank for
distribution to the exposure chambers. The worms should be handled as little as possible.
Handling should be conducted quickly and carefully so that the worms are not unnecessarily
stressed. Any worms that are accidentally dropped onto hard surfaces or are injured during
handling should be discarded. To prevent possible damage to the worms during handling,
various handling procedures can be employed. One procedure is to use a small, fine-point
paint brush to remove the organisms from the holding tank. Because Neanthes produce
mucus over the body surface, individual worms are easily captured and transferred with this
procedure. Another handling procedure is to use a wide-bore pipette with an attached bulb.
Individual organisms can be collected in the pipette through suction and can be removed from
the pipette using a gentle flushing action.
Individual worms, in excess of the number needed to conduct the bioassay, are trans-
ferred from the holding tank to a shallow dish containing seawater maintained at the test
temperature and salinity. Worms placed in the shallow dish should be as similar in size as
possible, given the size range of worms available from the holding tank. Worms transferred
to the shallow dish should be observed to determine that they represent the best worms
available for testing (e.g., all appear healthy and represent the smallest range in size of test
organisms).
Individual worms are removed from the dish and randomly placed in a plastic cup (five
worms per cup) containing seawater. Enough cups are used to equal three more than the
number of exposure chambers that will be used during the bioassay. Once this procedure
has been completed, worms within a cup are randomly transferred to an exposure chamber
by pouring the contents into the chamber. A squirt bottle containing seawater maintained
at the test temperature and salinity can be used to free any worms adhering to the cup. During
the transfer process, three of the cups containing worms are randomly selected and set aside.
Worms from these cups are used to estimate initial total biomass. To determine initial total
biomass, worms from these three cups are quickly rinsed with an isotonic 0.9 percent (W:V)
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ammonium formate solution of distilled water, placed on a preweighed aluminum pan, dried
at 50°C to a constant weight, and weighed to the nearest 0.1 mg.
Once worms have been placed in all of the exposure chambers, each chamber is checked
to ensure that air is flowing to the chamber and that the worms have begun to burrow into
the sediment. Following setup, food (e.g., TetraMarin®) is provided to each chamber. To
ensure adequate distribution of the food within the exposure chamber, a small volume of
seawater (i.e., 5 mL) at test temperature and salinity is added to the cup containing the
preweighed food ration. Once wetted, the food is poured into the exposure chamber. Water
from a squirt bottle is used to rinse the cup of any remaining food.
Following placement of the worms in the exposure chamber, initial (i.e., 1 hour) observa-
tions of burrowing should be made. If a worm, or group of worms, do not appear to be
burrowing and the observer believes that the nonburrowing behavior results from factors other
than sediment toxicity (e.g., reduced viability or damage to test organisms), then those
organisms should be replaced.
Monitoring-During the 20-day exposure period, the test chambers are observed on a
daily basis to ensure that adequate aeration is provided and to note the general status of each
chamber (e.g., presence of accumulated food, burrowing activity of worms, and presence of
fouling on sediment surface). On an every-other-day basis, worms in each exposure chamber
are provided with food. As discussed earlier, 40 mg of food are provided to each exposure
chamber. This food ration is maintained throughout the exposure period, even though
mortality may occur during the test.
Every third day, one-third of the seawater in each exposure chamber is replaced. Water
replacement is achieved by removing the aeration line, then siphoning one-third of the volume
and carefully replacing it with fresh seawater that has been maintained at 20 ± 1°C and at
the appropriate test salinity. Steps should be taken during seawater replacement to ensure
that test sediments are not disturbed. One method of replacement is to add the fresh seawater
by allowing the water to slowly flow down the inside wall of the exposure chamber. When
the chamber is filled, the aeration line is placed back in the chamber and the air flow is
adjusted to the specified level (i.e., 150 to 300 mL/minute).
Prior to seawater replacement, dissolved oxygen, salinity, and pH are determined for each
exposure chamber. Dissolved oxygen is determined using a dissolved oxygen electrode.
Following determination of dissolved oxygen in each chamber, the electrode is thoroughly
rinsed with 20 ± 1°C seawater. Salinity is determined on a small sample of seawater using
a hand-held refractometer. The seawater sample for the salinity measurement is obtained
with a Pasteur pipette. The pipette should be thoroughly rinsed with seawater between
samples. The pH is determined with a portable pH meter and probe. As with the dissolved
oxygen electrode, the pH probe is rinsed between readings.
Termination—Following the exposure period, worms from each exposure chamber are
removed from the test sediment. Two methods can be used to collect worms from each
exposure chamber. In the first, surviving worms are collected by sieving the sediment through
a 0.5-mm screen. The sieve should be gently shaken in a water bath rather than sprayed with
water to remove the sediment. In the second method, the sediment is placed in a white
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enamel pan containing seawater and searched for surviving worms. Worms collected from
sediment often remain in their tubes. A worm can be removed from the tube by gently
prodding either end of the tube to force the worm to leave. Once out, the worms are removed
using either the tip of a small paint brush or a wide-bore pipette. Following collection, the
number of worms surviving is noted on data sheets.
To determine total biomass, surviving worms are quickly rinsed in isotonic 0.9 percent
(W:V) ammonium formate or distilled water, placed on a preweighed drying pan, and dried
at 50°C until a constant weight is attained. Total biomass is determined to the nearest 0.1
mg as the difference in weight of the aluminum pan with and without the worms. Prior to
rinsing the worms, observations should be made to determine if food or sediment is present
in the digestive tract. Such information may be useful in explaining changes in individual
biomass occurring during the exposure period.
During the sublethal test demonstration study, a constant dry weight was attained within
24 hours. To determine when a constant weight has been achieved, several aluminum pans
containing worm samples are removed from the drying oven, placed in a desiccator, and
allowed to reach room temperature. Following cooling, each aluminum pan is placed on the
balance and the weight is determined. Following dry weight determinations, all samples are
placed back in the drying oven. After additional drying (i.e., at least 1 hour), the same samples
are again removed from the drying oven, allowed to cool In the desiccator, and reweighed.
When the dry weights for the samples are the same for consecutive readings (i.e., within 0.1
mg of each other), a constant weight has been attained.
Experimental Design
Logistics—A typical Neanthes bioassay for testing 10 sediment samples involves about
50 to 60 exposure chambers. Collection and preparation of test organisms, sediment, and
seawater requires at least four people for 2 days. Three or four people are required on the
days tests are initiated and terminated. One or two people can monitor a test in progress.
Typically, five replicate exposure chambers should be included for each sediment tested
including all control and reference sediments. Five replicates provide sufficient observations
per treatment to allow for statistical differentiation between treatments. However, the number
of replicates used in any experiment should be based on the objectives of the study rather
than on the need to meet the statistical testing requirements recommended in this protocol.
For example, if the bioassay is to be used during a reconnaissance survey to identify
potentially contaminated sediments, the replicates may not be needed to meet study
objectives.
Controls—A control sediment and a reference sediment should be included as part of
every test. The control sediment provides a nontoxic sediment to evaluate the condition of
the test organisms being used in the bioassay. The reference sediment provides a test
reference to partition contaminant effects associated with the treatment sediment from those
relating to noncontaminant characteristics (e.g., grain size and total organic carbon).
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A positive (toxic) control is also required for all testing. This involves determining 96-hour
LCjo values for Neanthes juveniles exposed in clean, filtered seawater without sediment to
reference toxicants (following standard bioassay procedures and under the same general test
conditions as the sediment bioassays). Such data are necessary to determine the relative
sensitivity of the animals (e.g., seasonal difference in sensitivity) for each test series to ensure
comparability of the data. The commonly used reference toxicant is reagent-grade cadmium
chloride. Reported 96-hour LC^ values for Neanthes exposed to cadmium chloride range
between 12.0 and 22.0 mg/L (Reish 1984; Johns and Ginn 1990).
The positive control should be conducted with 10 juveniles per exposure chamber. The
worms should not be fed during the 96-hour LCso exposure.
The acute lethality results must be reported along with the sediment bioassay results.
Bioassays to establish an LC^ involve four or five logarithmic concentration series and a
control. At least one treatment should give a partial response below the LC^ and one above
the LCsq. Statistical procedures for the LC^ estimate are given in American Public Health
Association (1985).
Response Criteria—Survival, total biomass (dry weight), and average individual biomass
(i.e., total biomass divided by the number of surviving worms) are the three response criteria
that can be determined for the Neanthes bioassay.
One of the three endpoints, data collected to date indicate that the survival endpoint is
the least sensitive to changes in level of contamination. Although survival rates of worms in
each replicate have generally been similar, it should be noted that variability in percent survival
within replicates could be high since each worm in a replicate represents 20 percent of the
replicate survival. The total biomass endpoint is an estimate of the biomass produced by the
group of worms in the exposure container. Total biomass represents an integrated measure-
ment of lethal and sublethal effects. Thus, a reduction in total biomass could indicate that
one or more worms had died during the exposure or that the growth of all worms had been
reduced. Average individual biomass is an estimate of the biomass of each surviving worm.
Unlike the survival and total biomass endpoints, worm survival is not integrated into the
determination of individual biomass. Worm survival is an important ancillary measurement
and should always be considered in the interpretation of either biomass endpoint. Each of
these response criteria should be monitored in a "blind" fashion; that is, the observer must
have no knowledge of the treatment of the sediment in the beakers.
DATA REPORTING REQUIREMENTS AND STATISTICAL ANALYSIS
The following data should be reported by all laboratories performing this bioassay:
¦	Water quality measurements during testing (i.e., dissolved oxygen, temperature,
salinity, pH)
¦	20-day survival in each exposure chamber and the mean and standard deviation
for each treatment
¦	Initial total biomass (dry weight) for three groups of five worms
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¦	20-day total biomass (dry weight) in each exposure chamber and the mean and
standard deviation for each treatment
¦	20-day average individual biomass (dry weight) in each exposure chamber and
the mean and standard deviation for each treatment
¦	Interstitial salinity values of control, reference, and test sediments (both initial
and final)
¦	96-hour LCgo values with reference toxicant
¦	Any problems that may have influenced data quality.
Data resulting from the Neanthes bioassay can be statistically analyzed using a number
of procedures depending upon study objectives and test design (i.e., number of replicates
used). Statistical procedures that might be used include the f-test and an analysis of variance.
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