United States Office of Research and EPA 600 3-90 076
Environmental Protection Development September 1990
Agency Washington, DC 20460
fa > ¦-
SEPA Red Algal
Sexual Reproduction
Supplemental Report for
Training Videotape
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Supplemental Report for the
Red Algal Sexual Reproduction Test
Training Videotape
U.S. Environmental Protection Agency
Center for Environmental Research Information
26 West Martin Luther King Drive
Cincinnati, Ohio 45268
U.S. Environmental Protection Agency
Environmental Research Laboratory
South Ferry Road
Narragansett, RI 02882
This report has been reviewed by the U.S. Environmental Protection Agency and approved for
publication. Mention of trade names, products, or services is not, and should not be interpreted as
conveying official EPA approval, endorsement, or recommendation.
PROPERTY OP
ENVIRONMENTAL PROTECTION AGENCY
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TABLE OF CONTENTS
Introduction
1
Culturing Champia pan'ula
2
Conducting the Test
4
Terminating the Test
5
Final Notes
6
References
8
Appendix A - Preparing Hypersaline Brine
Appendix B - Nutrients and Media
Appendix C - Materials Needed for Testing
Appendix D - Summary of Test Conditions
Appendix E - Sample Data Sheets
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Red Algal Sexual Reproduction Test
INTRODUCTION
The U.S. Environmental Protection Agency's
(EPA's) Environmental Research Laboratory in Duluth,
Minnesota, has developed a series of freshwater toxicity
tests to evaluate effluent toxicity. The tests use freshwater
fish, invertebrates, and plant species as indicators of toxic
effects to lakes, streams, and rivers. The EPA
Environmental Research Laboratory in Narragansett,
Rhode Island (ERL-N), has adapted the freshwater toxicity
testing methods and developed new methods to perform
similar testing in the marine or estuarine environments.
This report summarizes methods developed at the
Narragansett laboratory for estimating the chronic toxicity
of marine or estuarine effluents and receiving waters on
the sexual reproduction of the marine macroalga, Champia
parvula. Males and females are exposed in a static system
to effluents or receiving waters for 2 days, followed by a 5-
to 7-day recovery period for the female plants in a control
medium. Cystocarp production by the female, which
indicates sexual reproduction, is used as the endpoint.
The methods described in this report and demonstrated in
the accompanying tape are detailed in the EPA methods
manual, "Short-term Tests for Estimating the Chronic
Toxicity of Effluents and Receiving Waters to Marine and
Estuarine Organisms" (EPA/600/4-87/028).
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CULTURING CHAMPIA PARVULA
Although there are three macroscopic stages in the
life history of Champia, only the male and female plants
are used in toxicity testing. EPA's Environmental
Research Laboratory in Narragansett maintains unialgal
cultures of Champia year round, and plant material is
available from the lab for starter cultures.
To keep a constant supply of plant material
available, maintain several unialgal stock cultures of males
and females simultaneously. New cultures should be
started weekly from excised branches so that cultures are
available in different stages of development. To guard
against microalgal contamination, use sterile technique
when culturing the algae - autoclave all stock solutions,
and flame all tools before cutting or transferring plants.
Keep the cultures on a cycle of 16 hours of light
and 8 hours of darkness. The light level should not
exceed 500 ft-candles. The light may have to be adjusted
to that level, depending on the reflecting characteristics of
the incubators. The temperature should be maintained at
22 to 24° C and the salinity at 28 to 30 ppt. Natural
seawater or a 50-50 mixture of natural and artificial
seawaters are best for cultures. Gently aerate the
cultures. Change alternate cultures' medium every week,
so that if a stock solution should become contaminated,
the entire batch will not be lost. While replenishing the
medium, divide the growing algae in half with sharp
forceps or discard half of the biomass to prevent
overcrowding. New cultures can also be started at this
time, using 1-cm branch tips. Add nutrients using a pipet,
or, as Narragansett has found, a squeeze bottle is quick
and easy to use. Recipes for the culturing medium and
nutrient solutions are provided in Appendix B.
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sterile hairs
-¦ trichogynes
1 mm
Apical tip of the female branch
showing sterile hairs and
trichogynes (reproductive hairs)
(Aquatic Toxicity Testing Manual,
p. 16).
At the end of approximately 3 weeks, there should
be enough plant material to conduct the test. Examine
the stock cultures to determine their readiness for testing.
Place a few female branch tips in seawater in a petri
dish, and examine them under a compound microscope to
determine if trichogynes are present. An inverted scope
works best with the petri dishes, although standard slides
and microscopes also can be used. Trichogynes are the
short, fine reproductive hairs to which the spermatia
attach. They should be seen easily near the apex of the
branch tip. Although both sterile hairs and trichogynes
occur on the apex, sterile hairs occur over the entire plant
thallus. Sterile hairs are wider and generally much longer
than trichogynes, and appear hollow, except at their tip,
where they seem to be plugged.
spermotio
Spermatia attached to the sterile
and reproductive hairs on the
female branch (Aquatic Toxicity
Testing Manual, p. 17).
1 cm
^-spermatial
sorus
A portion of a male thallus
showing spermatial son (Aquatic
Toxicity Testing Manual P- 16).
Males should be visibly producing spermatia.
Sometimes, the presence of spermatial sori can be
determined by placing some male tissue in a petri dish and
holding it against a dark background. Mature son can also
be easily identified under a microscope along the edge of
the thallus. The sorus areas are generally thicker, and
lighter in color, than the rest of the plant body. At higher
magnification, the spermatia themselves can be seen. The
readiness of the male stock culture can also be assessed by
placing a portion of a female plant into some of the water
from the male culture for a few seconds. Under a
microscope, numerous spermatia should be seen attached
to the sterile hairs and trichogynes of the female plant.
Once readiness is established for both males and
females, the test can begin.
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CONDUCTING THE TEST
Prepare cuttings from the most healthy-looking
plants. Prepare the female cuttings first to minimize the
chances of contaminating them with water containing
spermatia from the male stock cultures. Place each plant
in a petri dish containing a little seawater. Using a fine-
point forceps or scalpel, prepare five cuttings from the
female plants for each treatment replicate, severing the
plant 7 to 10 mm from the ends of the branch. Try to be
consistent in the degree of branching in the cuttings, since
cystocarps form at the branch tips. For male plants, use
one cutting for each treatment replicate, severing the plant
about 2 cm from the end of the branch. If there are few
branches, or the spermatial sori appear sparse, larger male
cuttings may be needed. The cuttings can be kept at room
temperature for up to an hour.
The effluent sample should be kept at 4° C in an
incubator or refrigerator until use, but should not be
stored longer than 24 hours. Under a hood, prepare 5
dilutions using a 0.3 or 0.5 dilution factor, in 300 or 400-
ml replicates. At Narragansett, 125-ml Erlenmeyer flasks
are used as test chambers, but any clean container can be
used. Natural seawater is preferred for the test solutions,
although artificial seawater may be used during the
exposure part of the test. Maintain the salinity at 28 to 30
ppt, and use colored labels to indicate treatment and
replicates.
The 2-day exposure period starts when the algae are
added to the test chambers. Add 5 female branches and 1
male branch to each chamber. Pick up the branch at the
base or cut end to avoid injuring the tips. Cover the
chambers, then expose the cultures to 16 hours of cool
white light and 8 hours of darkness a day for the 2-day
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exposure and 5- to 7-day recovery periods. Maintain the
temperature between 22° and 24° C, and the salinity
between 28 and 30 ppt.
Check on the chambers twice a day, and gently
hand-swirl, or shake continuously at 100 rpm on a rotary
shaker. Spermatia are not motile, so some motion is
critical during the exposure period for reproduction to
occur. If desired, the media can be changed after 24
hours. Record the temperature daily from a thermometer
placed in a flask of water among the chambers.
After 48 hours, the algae are removed from the
effluent samples and allowed to recover for 5 to 7 days.
Label recovery bottles with the effluent concentrations
tested, and fill with 150 ml of natural seawater and
nutrients. Smaller volumes should be be used with a
shaker (about half the vessel volume), but make sure that
adequate growth will occur without having to change the
medium. With forceps, gently remove all of the females
from each test chamber, and place them into recovery
bottles. When all the replicates have been transferred,
place the vessels under cool white light and aerate or
shake for the 5- to 7-day recovery period. Aeration will
enhance the growth rate of plants in the recovery bottles,
although adequate growth will occur using a shaker.
Aerate using plastic tubes held in place by foam stoppers.
TERMINATING THE TEST
At the end of the recovery period, drain the
chambers and remove the females with forceps, starting
with the control plants and ending with those in the
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-ostiole
spores
A mature cystocarp. In controls
and lower effluent concentrations,
they often occur in clusters of 10
or 12 (Short-term Methods
manual, p. 286).
young branch
cells
immature
cystocarp
A young branch vj. an immature
cystocarp (Short-term Methods
manual, p. 286).
1 mm
An aborted cystocarp. A young
branch will eventually form at the
apex (Short-term Methods manual,
p. 286).
highest concentration. Place the female plants between
the inverted halves of a petri dish containing a small
amount of seawater, and count the cystocarps under a
stereomicroscope. Cystocarps are distinguished from young
branches by the darkly pigmented spores enclosed in the
nodule, and the apical opening for spore release. If there
is doubt about the identification of an immature cystocarp,
aerate the plants a little longer in the recovery bottles.
Within 24 to 48 hours, the suspected cystocarp will either
look more like a mature cystocarp or a young branch, or
will have changed very little, if at all, indicating an aborted
cystocarp. Occasionally cystocarps will abort, and these
should not be included in the counts. Aborted cystocarps
are easily identified by their dark pigmentation and, often,
by the formation of a new branch at the apex. Dead
plants lose their pigmentation and appear white.
FINAL NOTES
The algal sexual reproduction test requires that
several criteria be met before the test results are
considered acceptable.
¦ Control plants should average 10 or more
cystocarps and there should be no mortality in
the control group.
¦ Control and lowest-concentration exposed
algae should be in good physical condition -
for example, the branches should not be
fragmented. Broken or fragmented branches
could indicate that the plants were unhealthy
from the beginning of the test.
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¦ The results from the replicate control
chambers should be similar.
¦ All replicates from the affected concentration
chambers should show effect.
The methods manual, "Short-term Tests for
Estimating the Chronic Toxicity of Effluents and Receiving
Waters to Marine and Estuarine Organisms," describes the
statistical analysis performed on the test results. The red
algal sexual reproduction test is currently used to assess
the potential toxic effects of complex chemical mixtures on
marine and estuarine organisms. Used in conjunction with
chemical-specific methods, this test can provide a
comprehensive and effective approach to assessing the
impact of complex effluents discharged to the marine and
estuarine environments.
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REFERENCES
Aquatic Toxicity Testing Seminar Manual. 1985. U.S. EPA Environmental Research
Laboratoi^, Narragansett, RI. ERL-N Contribution No. 796.
(Provides detailed descriptions of the methods used at Narragansett to evaluate the
toxicity of discharges to marine and estuarine waters. Served as the basis of a series of
seminars conducted by ERL-N personnel.)
Biomonitoring for Control of Toxicity in Effluent Discharges to the Marine Environment. 1989.
U.S. EPA Center for Environmental Research Information, Cincinnati, OH; U.S. EPA
Environmental Research Laboratory, Narragansett, RI. EPA/625/8-89/015.
(Describes the use of biomonitoring as an effective, water quality-based approach to
controlling the toxicity of discharges to estuarine and marine waters. Covers regulatory
background, testing methods, and case studies.)
Short-term Methods for Estimating the Chronic Toxicity of Effluents and Receiving Waters to
Marine and Estuarine Organisms. 1987. Environmental Monitoring and Support Laboratory,
Cincinnati, OH. EPA/600/4-87/028.
(Describes methods, quality assurance, laboratory safety, facilities and equipment, data
analysis, report preparation, and organism culture and handling for six short-term tests to
estimate the chronic toxicity of effluents and receiving waters.)
Technical Support Document for Water Quality-based Toxics Control. 1985. U.S. EPA Office
of Water Enforcement and Permits, Washington, D.C.
(Provides guidance for each step in the water quality-based toxics control process, from
screening to compliance monitoring.)
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APPENDIX A
PREPARING HYPERSALINE BRINE
BACKGROUND
Salinity adjustments are a vital part of using marine and estuarine species for toxicity
testing. The majority of industrial and sewage treatment effluents entering marine and
estuarine waters contain little or no measurable salts. Therefore, the salinity of these effluents
must be adjusted before exposing estuarine or marine plants and animals to the solutions. In
addition, it is important to maintain constant salinity across all treatments throughout the test
for quality control. Finally, matching the test solutions' salinity to the expected receiving
water's salinity may require salinity adjustments. ERL-N uses hypersaline brine, prepared from
filtered natural seawater, to adjust exposure solution salinities. Note that commercially available
artificial sea salts have not been sufficiently tested, and therefore are not recommended for all
of the subchronic toxicity tests at this time.
Hypersaline brine has several advantages over artificial sea salts that make it more suitable
for use in toxicity testing. Concentrated brine derived from natural seawater contains the
necessary trace metals, biogenic colloids, and some of the microbial components necessary for
adequate growth, survival, and/or reproduction of test organisms. It may be held for prolonged
periods without any apparent degradation. Brine may be added directly to the effluent to
increase the salinity, or may be used as control water by diluting to the desired salinity with
deionized water. The brine can be made from any high quality, filtered seawater supply
through simple heating and aerating.
GENERATING THE BRINE
The ideal container for making brine from natural seawater has a high surface-to-volume
ratio, is made of a non-corrosive material, and is easily cleaned. Shallow fiberglass tanks are
ideal.
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Collect high quality (and preferably high salinity) seawater on an incoming tide to minimize
the possibility of contamination. Special care should be used to prevent any toxic materials
from coming in contact with the seawater. The water should be filtered to at least 10 um
before placing into the brine tank. Thoroughly clean the tank, aeration supply tube, heater,
and any other materials that will be in direct contact with the brine before adding seawater to
the tank. Use a good quality biodegradable detergent, followed by several thorough deionized-
water rinses. Fill the tank with seawater, and slowly increase the temperature to 40° C. If a
heater is immersed directly into the seawater, make sure that the heater components will not
corrode or leach any substances that would contaminate the brine. A thermostatically
controlled heat exchanger made from fiberglass works well.
Aeration prevents temperature stratification and increases the rate of evaporation. Use an
oil-free air compressor to prevent contamination. Evaporate the water for several days,
checking daily (or more or less often, depending on the volume being generated) to ensure that
the salinity does not exceed 100 o/oo and the temperature does not exceed 40° C. If these
changes are exceeded, irreversible changes in the brine's properties may occur. One such
change noted in original studies at ERL-N was a reduction in the alkalinity of seawater made
from brine with salinity greater than 100 o/oo, and a resulting reduction in the animals' general
health. Additional seawater may be added to the brine to produce the volume of brine desired.
When the desired volume and salinity of brine is prepared, filter the brine through a 10-
um Filter and pump or pour it directly into portable containers (5-gallon Cubitainers or
polycarbonate water cooler jugs are most suitable). Cap the containers, and record the
measured salinity and the date the brine was generated. Store the brine in the dark at room
temperature until used.
SALINITY ADJUSTMENTS USING HYPERSALINE BRINE
To calculate the volume of brine (Vb) to add to 0 o/oo sample to produce a solution at a
certain salinity (Sr), use this equation:
A-2
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vb * sb = Sf * V,
where Vb = volume of brine, ml
Sb = salinity of brine, o/oo
S, = final salinity, o/oo
Vf = final volume, ml (brine brought to this volume with 0 o/oo sample).
To calculate the volume of brine (Vb) required to raise the salinity of an effluent or
receiving water sample (S.) to a certain salinity (Sr), use this equation:
V„ = [(Vf * Sr) - (V. * S,)]/(Sb - Sf)
where Vb = volume of brine, ml
Sb = salinity of brine, o/oo
V, = volume of sample, ml
S. = salinity of sample, o/oo
Sf = final salinity , o/oo
V( = final volume, ml (final volume is combined brine and deionized water plus'the
sample volume; percent original sample in the final sample = VyVf * 100).
The table on the next page gives volumes needed to make 20 o/oo test solutions from effluent
(0 o/oo), deionized water, and 100 o/oo hypersaline brine.
A-3
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Quantities of effluent, deionized water and a hypersaline brine of 100 o/oo (only) needed for
conducting daily renewals of test solutions at 20 o/oo salinity.
Exposure
Deionized
Hypersaline
Concentration
Effluent
Water
Brine
(0 o/oo)
(100 o/oo)
(*>
(ml)
(ml)
(ml)
32
640
960
400
10
200
1,400
400
3.2
65
1,535
400
1.0
20
1,580
400
0.32
7
1,593
400
Control
1,600
400
A-4
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APPENDIX B
NUTRIENTS AND MEDIA
This table lists the additional nutrients to be added to natural seawater for stock cultures
and test medium. The concentrated stock solution is autoclaved at standard temperature and
pressure for 15 minutes. Adjust the solution to about pH 2 before autoclaving to minimize the
possibiity of precipitation.
AMOUNT/LITER CONCENTRATED STOCK SOLUTION
COMPOUND CULTURE MEDIUM TEST MEDIUM
Nutrient Stock Solution8
NaNOj
6.35 g
1.58 g
NaH2PO
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APPENDIX C
MATERIALS NEEDED FOR TESTING
Lab Supplies
Fine-point stainless steel forceps
100-ml polypropylene cups with covers (or 125-ml Erlenmeyer flasks) for exposure chambers
Polystyrene petri dishes for holding cuttings and treated plants for counting
100-ml graduated cylinder
1- and 10-ml disposable pipets
Digital micropipets (200 and 1000 ul maxima) if dilutions are made directly in test chambers
Disposable micropipet tips
Recovery bottles or flasks (one per treatment or control chamber)
Aquarium pump(s)
Air tubing (made of non-toxic material)
Plastic aeration tubes (1 ml disposable pipets work fine)
Foam plugs
Thermometer
Marking pens
Colored tape
Data Sheets (Appendix E)
Lab Equipment
Cool-white fluorescent lighting, sufficient to give 75 uE m'2 s'1 (about 500 ft-candles).
Rotary shaker (hand-swirling the exposure chambers twice a day can be substituted).
Stereo (dissecting) microscope
Refractometer (salinometer)
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APPENDIX D
SUMMARY OF TEST CONDITIONS
Test type:
Salinity:
Temperature:
Photoperiod:
Light source:
Irradiance:
Test solution volume:
Test chamber size:
Number of test organisms
per test chamber:
Number of replicate
chambers per treatment:
Aeration:
Dilution water:
Test duration:
Effect measured:
Static, non-renewal
30 o/oo
22 to 24 °C
16h light:8h dark
cool-white fluorescent
about 75 uE m"2 s"1
100 ml
110-ml polypropylene cups (with covers) or 125-ml
Erlenmeyer flasks
5 female and one male branch tip
3
None; chambers are either shaken at 100 rpm on a
rotary shaker or hand-swirled twice a day
30 o/oo natural or artificial seawater with additional
nutrients added
2-day exposure followed by a 5- to 7-day recovery
period for cystocarp development
Sexual reproduction (number of cystocarps per
female)
D-l
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APPENDIX E
DATA SHEETS
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CHAMP I A PAJtVULA
Cystocarp Data Sheet
Collection Date:
Exposure Began:
Recovery Began:
Counted:
Experiment No.
Treatments: (% Effluent, ug/L. or Receiving Water Sites)
Replicat Control
A1
2
3
4
Mean
B1
I i
t
1
l
2
— —
4
5
Mean
CI
2
3
4
i
5
i
Mean
1
Overall
Mean
St Dev.
% of Cont
Temperature:
Salinity:
Light:
Dilution Water:
Mean Sq.Error=
_ Degrees Freedom=
Dunnett's Tab Val=
n cont.=
n treat. =
Critical Valued'=
it l/.S. GOVERNMENT PR/NT I KG OFFICE: 1990 - 748-159/J0497
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