EPA-600/3-76-114
December 1976
Ecological Research Series
TOXICITY OF DDT FOOD AND WATER
EXPOSURE TO FATHEAD MINNOWS
Environmental Research Laboratory
Office of Research and Development
U.S. Environmental Protection Agency
Duluth, Minnesota 55804
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RESEARCH REPORTING SERIES
Research reports of the Office of Research and Development, U.S. Environmental
Protection Agency, have been grouped into five series. These five broad
categories were established to facilitate further development and application of
environmental technology. Elimination of traditional grouping was consciously
planned to foster technology transfer and a maximum interface in related fields.
The five series are:
1. Environmental Health Effects Research
2. Environmental Protection Technology
3. Ecological Research
4. Environmental Monitoring
5. Socioeconomic Environmental Studies
This report has been assigned to the ECOLOGICAL RESEARCH series. This series
describes research on the effects of pollution on humans, plant and animal
species, and materials. Problems are assessed for their long- and short-term
influences. Investigations include formation, transport, and pathway studies to
determine the fate of pollutants and their effects. This work provides the technical
basis for setting standards to minimize undesirable changes in living organisms
in the aquatic, terrestrial, and atmospheric environments.
This document is available to the public through the National Technical Informa-
tion Service, Springfield, Virginia 22161.
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EPA-600/3-76-114
December 1976
TOXICITY OF DDT FOOD AND WATER
EXPOSURE TO FATHEAD MINNOWS
by
Alfred W. Jarvinen
Molly J. Hoffman
Todd W. Thorslund
Environmental Research Laboratory-Duluth
Duluth, Minnesota 55804
ENVIRONMENTAL RESEARCH LABORATORY-DULUTH
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
DULUTH, MINNESOTA 55804
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DISCLAIMER
This report has been reviewed by the Environmental Research Laboratory-Duluth,
U.S. Environmental Protection Agency, and approved for publication. Mention
of trade names or commercial products does not constitute endorsement or
recommendation for use.
ii
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FOREWORD
Our nation's freshwaters are vital for ail animals and plants, yet our
diverse uses of water for recreation, food, energy, transportation, and
industry physically and chemically alter lakes, rivers, and streams. Such
alterations threaten terrestrial organisms, as well as those living in water.
The Environmental Research Laboratory in Duluth, Minnesota develops methods,
conducts laboratory and field studies, and extrapolates research findings
—to determine how physical and chemical pollution affects aquatic life
—to assess the effects of pollutants
—to predict effects of pollutants on large lakes through use of models
—to measure bioaccumulation of pollutants in aquatic organisms that are
consumed by other animals, including man
This report determines the effects of DDT on fathead minnows when they
are exposed to it in the food and/or water.
Donald I. Mount
Director
Environmental Research Laboratory
Duluth, Minnesota
iii
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ABSTRACT
Fathead minnows (Pimephales promelas) were exposed during a partial chronic
toxicity test to two DDT concentrations in the water, one in the diet, and
combinations of water and diet for 266 days through a reproductive period
of their life cycle. Tissue-residue analyses were performed on test fish at
preset intervals throughout the exposure and also on embryos, larvae at hatch,
and 30- and 60-day progeny. The contribution of DDT from each source was
monitored with gas-chromatography and liquid-scintillation techniques". The
diet was clams that had accumulated liiC-DDT when exposed at a DDT water
concentration similar to that in the high fish exposure.
Higher total DDT tissue residues were accumulated .from the water than from the
diet. Residues contributed by dietary DDT were additive to those from the
water. Mean concentration factors were 1.2 times from the diet and 100,000
times from the water. Mortality was higher in fish exposed to DDT in both
water and diet than in fish exposed to only one or the other of these sources.
DDT in the diet significantly reduced the probability of survival of the test
fish (P=0.025). Estimated maximum acceptable toxicant concentrations for DDT
are 0.9 Wg/1 for fish exposed to DDT in the water only or O.U yg/1 for fish
exposed to DDT in both water and diet. Embryo DDT residues and larval
mortality were about twice as great for embryos and larvae from parent fish
that had been exposed to DDT in both water and diet as for those from parent
fish exposed to DDT in the water only.
About 60% of the mean total micrograms of combined DDT analogs in fish that
had been exposed to DDT at 0.5 yg/1 in the water and in the diet was
eliminated within 56 days. Almost all of the eliminated DDT was dietary
DDT. Elimination in fish that had been exposed to DDT in the water only
was negligible.
iv
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CONTENTS
Foreword iii
Abstract iv
List of Figures vi
List of Tables vii
Acknowledgments „ viii
I Introduction 1
II Conclusions 2
m Recommendations 3
IV Materials and Methods 4
Physical Conditions 4
Fathead Minnov Exposure 4
Clam Exposure 5
Biological Conditions 5
Fish 5
Clams 7
Chemical Conditions 8
Fish 8
Clams 8
Residue Analysis ..... 9
Gas Chromatography 9
Liquid Scintillation 9
Statistics 10
V Results 11
Adult Fish 11
Embryos 20
Larvae at Hatch 24
Progeny at 30 and 60 Days 27
Elimination Study 30
VI Discussion 38
References • • 49
Appendix
Recommended Bioassay Procedure for Fathead Minnow
Pimephales promelas Rafinesque Chronic Tests 53
v
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LIST OF FIGURES
Number Page
1 Total DDT residues in the controls and fish exposed to DDT in
the food 13
2 Total DDT residues in fish exposed to DDT in the water (0.5 Vg/l)
or in combination of food and vater 13
3 Total DDT residues in fish exposed to DDT in the water (2.0 Vg/l)
or in combination of food and water 14
U Accumulative probability of survival for fish that had been exposed
to DDT „ 21
5 Elimination of total DDT from fish 32
6 Elimination of DDT from fish 35
7 Elimination of DDE from fish 36
8 Elimination of TDE from fish 37
9 Estimated probability of death for fish exposed to DDT at various
water concentrations and fed clean or DDT-contaminated food ... 45
VI
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LIST OF TABLES
Number Page
1 DDT Water Concentrations in Exposure Chambers 12
2 DDT, DDE, and TDE Residues in Fish Exposed to Various Test
Conditions 16
3 DDT, DDE, and TDE Residues in the Fish Food 17
U Percentage of Total DDT Caused by the ll*C-DDT Food Source . . „ . . 19
5 Estimated Probability of Survival for Fish Exposed to Various Test
Conditions for 266 Days 19
6 Hatchability of Embryos at Various Test Exposures 22
1 Total DDT Residues in Embryos from Fish Exposed to Various Test
Conditions, and Percentage Total DDT Contributed by the
Contaminated Food 23
8 DDT, DDE, and TDE Residues in Embryos from Fish Exposed to Various
Test Conditions 25
9 Total DDT Residues in Larvae at Hatch from Fish Exposed to Various
Test Conditions, and Percentage Total DDT Contributed by the
Contaminated Food 25
10 DDT, DDE, and TDE Residues in Larvae at Hatch from Fish Exposed to
Various Test Conditions 26
11 Total DDT in 30- and 60-Day-Old Progeny of Fish Exposed to Various
Test Conditions, and Percentage of Total DDT Contributed by the
Contaminated Food 28
12 DDT, DDE, and TDE Residues in 30- and 60-Day-Old Progeny of Fish
Exposed to Various Test Conditions 29
13 Percentage Survival of 30- and 60-Day-Old Progeny from Parent Fish
Exposed to Various Test Conditions 31
li* Mean Percentage of Total DDT Residues Remaining in Fish that was
Attributed to the Food or to the Water 34
15 Estimated P Values, Average Measured DDT Water Concentrations, and
Values for Estimated Parameters from Non-Linear Least Square
Estimation for Fish Exposed to Various Test Conditions 44
vii
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ACKNOWLEDGMENTS
We wish to thank Ms. H. E. Herrmann for daily assistance, clam shucking,
and routine chemical analysis; Messrs. L. H. Mueller, K. D. Kempfert,
D. Seeger, R. M. Pieper, and Ms. S. Kubicek for DDT water and tissue-residue
analysis; Mr. D. T. Allison for assistance in development of liquid-scintillation
techniques; and all other members of the pesticide research team for assistance
and advice. We also wish to thank Mr. J. G. Eaton and members of the Environmental
Research Laboratory-Duluth committee for advice and review of the manuscript.
viii
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SECTION I
INTRODUCTION
There are many opinions among aquatic researchers about the importance of water
concentration and food chain as sources for biological magnification of pesticides
in the aquatic environment. Some authors suggest the food chain as the major
source (Macek and Korn, 1970; Harrison et_ aJ^. , 1970; Johnson et_ al_., 1971;
Eberhardt et_ al., 1971), whereas others (Reinert, 1967, 1970; Chadwick and
Brocksen, 1969; Grzenda et_ ad., 1970; Murphy, 1971; Hamelink _et_ al_., 1971;
Epifanio, 1973) stress the water concentration. No data are available indicating
the relationship of either of these views to those situations where both sources
are involved at threshold levels of chronic toxicity.
Current pesticide standards are based upon water concentration alone. We must
also know what effect the presence of pesticide-laden food-chain organisms has
on aquatic life so that accurate pesticide standards can be developed. The
following study was initiated in response to this problem. The objectives were
to determine whether DDT accumulation in fathead minnows (Pimephales promelas)
is more affected by a food or water source, to determine whether persistent
pesticide exposure through both food and water is more toxic (or creates higher
residues) than exposure through only one or the other of these routes, to estimate
a partial chronic maximum acceptable toxicant concentration for DDT, and to
determine DDT concentration factors for fathead minnows and freshwater clams used
as the food.
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SECTION II
CONCLUSIONS
Fathead minnow DDT concentration factors are about 1.2 times from the diet and
100,000 times from the water. Clams used as the DDT food source had a
magnification factor of about 25,000 times when exposed to a similar water
concentration (2.0 yg/l). Tissue residues in fish exposed to dietary DDT only
were about one-fourth as high as residues in fish exposed to a water concentration
equal to that at which the food had been exposed. Dietary DDT tissue residues
were additive to those resulting from DDT water concentrations.
Fathead minnow mortality was greater in fish exposed to DDT through both food
and water than in those exposed through only one or the other of these routes.
Water exposure alone, however, was more toxic than food exposure alone. Mortality
results agree closely with those for residue accumulation, indicating that higher
mortality occurs with higher mean tissue residues.
An estimated maximum acceptable toxicant concentration for this test is 0.9 yg/1
for water exposure alone or O.k yg/1 when DDT is present both in water and in
diet (U5.6 yg/g).
Elimination of DDT from fathead minnows that had been exposed to it in the water
at 0.5 yg/1 or at the same water concentration and in the diet also indicated
that about 60% of the accumulated mean total micrograms of total DDT in fish
that were exposed to it in diet and in the water was eliminated within 56 days.
Elimination from fish exposed to DDT in the water only was negligible. It appears
that there is a selective mechanism for elimination of dietary DDT.
Exposure of fish to DDT in diet or in water are both important and should be
considered together in future studies. Presence of dietary DDT can reduce the
maximum acceptable toxicant concentration.
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SECTION III
RECOMMENDATIONS
It is recommended that chronic toxicity studies on persistent toxicants be
performed with additional consideration for possible accumulation through the
food chain. More such studies are needed to evaluate the combined food and
water toxicant effect on fish and other aquatic life. Future studies should be
designed to provide food-effect data needed to derive more refined criteria
necessary to determine the survival requirements for aquatic life.
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SECTION IV
MATERIALS AND METHODS
The basic design of the partial chronic exposure followed the recommended
procedures set forth by the National Water Quality Laboratory Committee
on Aquatic Bioassays (Appendix).
PHYSICAL CONDITIONS
Fathead Minnow Exposure
A proportional diluter (Mount and Brungs, 19&7) was modified to deliver
two test concentrations and a control with the low test concentration
one-quarter that of the high concentration. The toxicant was introduced
by a 50-ml injector syringe with a TefloiWneedle from an acetone stock
solution. The syringe was calibrated to inject 8.7 pi of stock solution
per cycle. The highest level of acetone ever reached, within any 2U-hr
period, in the high concentration test chamber was 5 mg/1. Nominal DDT
test concentrations, selected on the basis of acute and preliminary partial
chronic data, were 2.0 and 0.5 pg/l» respectively.
The test water was sand filtered Lake Superior w,ater sterilized with ultra-
violet light and warmed to approximately 25° C by a coiled stainless steel heat
exchanger located in a stainless steel headbox. A thermoregulator relay
system (Syrett and Dawson, 1972) activated tandem solenoid valves that
controlled the flow of hot water through the heat exchanger.
The test chambers used for adult exposures measured 91 x 30 x 30 cm and
held a water volume of.55 1. Approximately 3 months after the start
of the test the adult tank was separated into two sections by stainless
steel screen. Two 30.5 x 13.5 x 31.5 cm larval chambers were placed in the
back section of each adult chamber, and flow rates were adjusted to provide
250 ml of test water to each larval chamber and 500 ml to the adult
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section. The flow of water to the test chambers was adjusted to maintain
dissolved oxygen levels at greater than 65$ saturation and to provide a
99% replacement time of the test water in each test chamber within 10 hr
as determined from Sprague (1969).
Chambers were siphoned daily 1-2 hr after feeding to remove leftover food
and were brushed and siphoned weekly.
The photoperiod followed the normal daylight hours of Evansville, Indiana,
except that it was necessary to extend the peak photoperiod of 15 hr ^5
min for approximately 6 weeks to insure enough larvae for gas-chromatography
and liquid-scintillation analysis of tissue residues. Daytime light
intensity varied from 25 to Ul ft-c in the adult chambers.
Clam Exposure
The clam exposures were conducted in a flow-through system. The system
consisted of a stainless steel headbox with coiled stainless steel heat
exchanger and thermoregulator relay system. The test water flowed from
the headbox through a solenoid valve to a 2^.5 x 13.5 x l6.5 cm water cell.
When a predetermined volume was reached, the water siphoned through an
inverted u-tube into a 19«5 x 17-5 x 13.0 cm toxicant chamber. Action of
the siphoning water flowing into a cup, mounted on an arm, activated a
microswitch to shut off the water flow from the headbox and also activated
a 50-ml injector syringe with a Teflon-^needle to inject 8.7 pi of
l^C-DDT-acetone stock solution into the toxicant chamber for a nominal DDT
concentration of 2.0 yg/1. The test water then passed through a standpipe
siphon into a common 28.0 x lU.7 x 31.0 cm glass chamber where the toxicant
and test water were mixed. Water from this chamber flowed through two
standpipe siphons to duplicate 152.5 x 30.5 x 28.0 cm stainless steel exposure
chambers. Water volume in each chamber was regulated by a standpipe at 7U 1.
The flow-through apparatus delivered 3.2 1 per cycle or 1.6 1 per chamber with a
99% replacement of the test water in about 6 hr as determined from Sprague (1969),
BIOLOGICAL CONDITIONS
Fish
On September 13, 1972, one hundred ^5(±30-day-old fathead minnows were randomly
assigned to each of 12 test chambers. The 12 chambers were used to expose the
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fish through a reproductive period of their life cycle to duplicates of (l) a
control; (2) DDT-exposed food, but no DDT in the water; (3) unexposed food, but
0.5 yg/1 DDT vater exposure; (k) DDT-exposed food and 0.5 yg/1 DDT water exposure;
(5) unexposed food, but 2.0 yg/1 DDT water exposure; and (6) DDT-exposed food
and 2.0 yg/1 DDT water exposure. The fathead minnows were fed chopped and
ovendried clam tissue which was either clean or contaminated by litC-DDT.
The fish were sampled to determine total body tissue residues at 7> 1^» 28, 56,
112, 22k, and terminally at 266 days exposure. The fish were not fed for 24 hr
before samples were taken. Samples consisted of 10 fish per duplicate test
chamber at 7 days of exposure and 5 fish per duplicate thereafter, until terminally,
when all remaining fish except those used for an elimination study were sampled.
Samples were placed in preweighed glass vials, reweighed, and frozen at -8° C until
analyzed. Results were determined both on a whole body wet weight and lipid basis.
At the termination of the 266-day partial chronic exposure 20 adult fathead
minnows (10 from each duplicate chamber) were transferred from the 0.5-Mg/l DDT
water exposure and the 0.5-yg/l DDT water and DDT-contaminated food exposure to
separate control chambers for use in an elimination study. Five fish were removed
for tissue-residue analysis from each of these chambers at 75 1^» 28, and terminally
at 56 days. To prevent bias caused by weight changes, results were determined
on a total microgram basis.
During the spawning period embryos in excess of the 50 required for hatchability
studies were placed in preweighed glass vials, reweighed, preserved with petroleum
ether to prevent dehydration, and stored in a freezer at -8° C. Individual test-
chamber samples were composited to provide a minimum of four 0.3-g (about 300
embryos) samples for residue analysis.
Larvae at hatch were transferred in groups of kO each to larval chambers
for 30- and 60-day growth, mortality, and residue studies. The fish were
photographed at 30 and 60 days by the method of Martin (1967) as modified
by McKim and Benoit (l9Tl) for growth determination. Tissue residues
were analyzed for two 30- and one 60-day samples for each of the
12 test chambers.
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Larvae at hatch not saved for 30- and 60-day studies were weighed and grouped by
the same method used for embryo-residue samples to provide a minimum of two 0.3-g
(about 600 larvae) samples per test chamber for residue analysis. Residue analysis
on larvae at hatch and embryos was performed only on a whole-body wet-weight basis.
Clams
Chopped, ovendried clam tissue was used as the food source. Clams were chosen
because they have a nearly average accumulation factor when compared to other
invertebrates (Johnson et_ aJ^., 1971; Eberhardt et_ al_. , 1971), were readily
available, are well suited to laboratory conditions, provide a large bulk of
storable tissue, and attain equilibrium with DDT in a relatively short time as
was indicated in preliminary tests. Five species of clams were collected from
the Eau Claire River in Wisconsin: Lampsilis siliquoidea, Lampsilis ventricosa,
Lasmigona costata, Fuscoraia flava, and Liqumia recta. The clams were held before
DDT exposure in a fiberglass tank through which lake water flowed. Four separate
8-week clam exposures to l^C-DDT were conducted, as preliminary studies indicated
that an 8-week exposure period was necessary for the clams to achieve an
equilibrium with an exposure concentration of 2.0 yg/1 DDT (the same DDT
concentration as in the higher fish water exposure).
Clams were placed in the exposure chambers and slowly acclimated to 20° C. Fifty
clams were used in each duplicate chamber per exposure, and 100 were held in a
fiberglass control chamber through which lake water flowed from the same headbox
as the exposure system. The control chamber had a volume of 170 1 and a water
flow rate to provide 99% replacement in 6 hr. Dissolved oxygen concentrations
never dropped below 80$ saturation.
The clams were fed daily with a commercial fish fry food and plankton. The
flow-through apparatus was monitored daily, the chambers were siphoned every
other day, and the sides of the chambers were scrubbed whenever algal or fungal
growth became excessive.
After completion of an 8-week exposure the soft parts of the clams were removed,
chopped in a blender, and ovendried for 1 1/2-2 hr at 110° C. The dried clam
meat was supplemented with a vitamin and mineral mix. A list of ingredients
for the mix was obtained from the Fish-Pesticide Research Laboratory, Columbia
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Missouri (Mehrle, personal communication). The prepared food supply was kept
frozen, and a small portion was removed daily for feeding.
CHEMICAL CONDITIONS
Fish
Water temperatures were maintained between 2h.O and 25-5° C and were checked
daily in all test chambers. Routine water chemistries were determined weekly
by the methods described by the American Public Health Association et_ al_. (l9Tl).
Dissolved oxygen levels were never lower than 5-^- mg/1 nor higher than
8.2 mg/1. Mean total hardness, acidity, and alkalinity were ^3.9» 2.8, and
U2.5 mg/1, respectively, and were similar to those mentioned by Hermanutz
et_ al_. (1973); pH was between 7-2 and 7-8. DDT stock solutions were prepared
*
with DDT, Technical grade (p,pr isomer 77$) and DDT concentrations in the
water were measured once a week. In each sample set, analyses were made
on a duplicate and a spiked sample of control water. DDT was extracted from
the test water with petroleum ether and analyzed by gas chromatography.
Percentage recovery from the spiked control water samples ranged from 38 to
115/2; mean recovery was 86.9+_3.H/5 (n=39).
Clams
P,p' DDT ring-UL-lUc in benzene with a specific activity of 3.85 yc/mM
was procured in 50-yc lots."1" The benzene was evaporated under a stream
of nitrogen, and the p,p' DDT ring-UL-l^C was redissolved in 50 ml of acetone.
To prepare the lUC-DDT stock solution a calculated gram weight of DDT (Tech.)
was dissolved in acetone to which U5 ml of the l^C-labeled DDT in acetone was
added. Total volume was brought to 250 ml with acetone and thoroughly mixed.
The clams were exposed to 2.0 yg/1 l^C-DDT nominal concentration and 1.35 PC
of lUC per day, or about 76 pc of lUC per 8-week exposure. The lUC-DDT
water concentrations ranged from 1.05 to 2.60 yg/1, with a mean concentration
of 1.81+0.13 (n=lH). Percentage recovery of spiked control water samples ranged
from 80 to 121%; mean recovery was 102.6+_8.5% (n=5). Water analysis for DDT was
performed by the same method as for the fathead minnow exposure.
3f
DDT (Tech. ) was obtained from the Nutritional Biochemicals Corporation,
Cleveland, Ohio.
P,p' DDT ring-UL-l^C was purchased from Mallinckrodt/Nuclear, St. Louis,
Missouri.
8
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RESIDUE ANALYSIS
Gas Chromatography
Extraction of adult fish and 30- and 60-day progeny was accomplished by transfer
of the samples to an Eberbach Semi-micro or Micro Container explosion-proof
blender (depending upon sample weight). Anhydrous Na2S(\ was added to dry the
tissues and insure homogenization. Samples were blended with petroleum ether
for 2 min at high speed. The solvent extract was decanted onto an anhydrous
Na2S04 column and collected in a tared beaker for lipid determination. Each
sample was extracted three times. The solvents were evaporated, and lipid
weight was determined. The lipids were redissolved in petroleum ether and then
cleaned up on a 20-g florisil column. Samples were eluted with 200 ml of 6%
ethyl ether/petroleum ether as described by the U.S. Department of Health,
Education, and Welfare (l9Tl). Samples were concentrated to a volume of less
than 10 ml on a steam bath. Analysis was completed by gas chromatography. Peak
heights were measured individually for DDT, DDE, and TDE. DDT and metabolites
were then summed to obtain the total DDT present. DDT, DDE, and TDE are
expressed as the sum of the orthopara and parapara fractions found in the
analyses.
Samples of embryos and larvae at hatch were homogenized in 10 ml of petroleum
ether in a glass tissue-grinding tube by using a teflon pestle. Samples
were extracted five to eight times depending upon total sample weight and were
concentrated as previously described for the adult fish. No cleanup
was necessary and no lipid analysis was performed. Residue analysis was
performed as for adult fish. Residue analysis on the clams was conducted
on duplicate 1-g samples of tissue from both contr.ol and l^C-DDT-exposed
clams before addition of the vitamin-mineral supplement. The samples
were extracted in a blender with 35% water/acetonitrile and then
partitioned with 100 ml of petroleum ether. Cleanup was performed as
described for the adult fish.
Liquid Scintillation
Radiometric methods were used to determine DDT residues attributed to the
food for all samples. Analysis was performed on a Packard Tri-Carb
Liquid Scintillation Spectrometer. Samples were prepared from the portion
of the extracted tissue residues not used in gas chromatographic analysis.
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The samples were evaporated to 0.2 ml in glass concentrator tubes and vere
then transferred to glass scintillation vials with four 1-ml washings of
toluene. Fifteen milliliters of Instagel* (scintillation cocktail) were
added to each vial. The vials were kept in the scintillation counter
overnight to allow them to cool and dechemiluminess before analysis.
A correlation was made between the gas chromatograms and scintillation
counts through a series of dilutions of the lliC-DDT stock solution. An
average count per minute per microgram of DDT was then calculated and used for
the determination of the micrograms of DDT attributed to the contaminated food
source. Individual sample counts per minute were corrected for background
radiation and sample volume removed for gas-chromatograph analysis before final
calculations were made. Final results were calculated on a whole-body wet-weight
basis.
STATISTICS
All survival and egg-hatchability data were transformed to arcsin ~\f%7
Two-way analysis of variance was applied to all survival, embryo-hatchability,
percentage lipid, and 30- and 60-day progeny growth data to determine the DDT
effect from food or water exposure. Dunnett's procedure (Steel and Torrie, 1960)
was used for comparison of treatment means with control means. Non-linear least
square estimation was used to determine values for the food and water parameters
that affected adult survival. Regression analysis was performed on tissue-residue
data obtained from the elimination study.
^Instagel was purchased from Packard Instrument Company, Inc., Downers Grove,
Illinois.
JO
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SECTION V
RESULTS
ADULT FISH
To simplify the presentation of results test treatments are coded as follows:
DDT Exposure Coding
Clean water, clean food (Control) C
Clean water, DDT food F
0.5 yg/1 DDT water, clean food 0.5 W
0.5 yg/1 DDT water, DDT food 0.5 W + F
2.0 yg/1 DDT water, clean food 2.0 W
2.0 yg/1 DDT water, DDT food 2.0 W + F
Results from duplicate chambers were combined, and the data are expressed as
the mean +_ standard error (S.E.) unless otherwise indicated.
Determined DDT water concentrations in the test exposures are presented in
Table 1.
Figures 1-3 show the total DDT tissue residues (DDT + DDE + TDE) at the
various sample periods. An equilibrium with dietary total DDT occurred within
28<-56 days (Figure 1). Figures 2-3 demonstrate the additive residue effect
of dietary total DDT when compared to residues from a water source only. In
general, total residues peaked by 56 days for fish exposed at F and 0.5 W and
by 112 days for the rest of the exposures. An equilibrium may have been
reached at 0.5 W within 56 days. In general, residue levels decreased rapidly
during the spawning period (112-224 days) and then increased after termination
of spawning activity. Residue levels fluctuated greatly, and apparently
neither sex was affected more than the other.
11
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TABLE 1. DDT WATER CONCENTRATIONS IN EXPOSURE CHAMBERS
Is}
Nominal DDT water concentration
(ug/l)
2.0 ¥
2.0 W + FC
0.5 W
0.5 ¥ + FC
FC
C (Control)
Measured concentration (yg/l)
N
Ul
hi
Ul
Ul
Ul
Ul
Mean
1.53 (O.35)a
1.U8 (0.30)
0.35 (0.11)
0.37 (0.11)
0.01 (0.03)d
0.00 (0.01)
Range
0. 82b-2. 30
0.82b-2.00
O.l6b-0.70
0.19l3-0.79
0.00-0.10d
0.00-0.066
a
Standard deviation in parentheses.
Syringe injector malfunction (leakage).
°Fed clam tissue with lUC-DDT (U5.6 mg/kg).
dLeaching from lUC-DDT food.
SPossible contamination from wrong food; occurred only once during
exposure.
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o>
3100
50
50
• C
100
150
200
250
TIME (days)
Figure 1. Total DDT residues (ng/g) in the controls and fish
exposed to DDT in the food. (Vertical lines
indicate standard error.)
0>
3100
Q
Q
50
f 0.5 W
50
100 150 200
TIME (days)
250
Figure 2. Total DDT residues (ug/g) in fish exposed to DDT in
the water (0.5 Wg/l) or in combination of food and
water. (Vertical lines indicate standard error.)
13
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4OO
350
300
O>
3
250
200
150
100
y
t~*'
• '
,/'/
^
I \
• 2.0 W+F
xi 2.0 W
50
50 100 150 200
TIME (days)
250
Figure 3. Total DDT residues (yg/g) in fish exposed to DDT in
the water (2.0 yg/1) or in combination of food and
water. (Vertical lines indicate standard error.)
14
-------
Residue levels after Ik days of exposure were for the most part greater for
fish exposed to DDT in both the water and diet than for those exposed in the
water only. After 266 days fish from the F exposure had a mean body burden
2.1+ times those exposed at 0.5 W, fish exposed at 0.5 W + F had mean residues
three times those exposed at 0.5 W, and mean residues in fish exposed at 2.0 W +
F were about two times greater than in fish exposed at 2.0 W.
The highest mean total DDT body burdens (yg/g) achieved for each exposure
group were as follows: C, 2.0 yg/g at lU days; F, 69 yg/g at 56 days;
0.5 W, 56 yg/g at 56 days; 0.5 W + F, 118 yg/g at 112 days; 2.0 W,
291 yg/g at 112 days; and 2.0 W + F, 337 yg/g at 266 days.
Total DDT residues are presented as DDT, DDE, and TDE in Table 2. DDE was the
principal constituent found after lit days of exposure, an indication that DDT
was rapidly metabolized. Lack of TDE in the residues of fish fed DDT-exposed
clam tissue until 112 days was caused by failure of the gas-chromatographic
column to differentiate p,p" TDE from o,p' DDT. Column changes permitted
differentiation after 56 days. In general, DDT levels decreased rapidly after
ih days. DDT, DDE, and TDE residues decreased at spawning time except at the
F exposure, where DDE alone increased. TDE residues were low at 22^ days for
fish exposed to DDT in the food. Analysis of the DDT-exposed clam tissue is
presented in Table 3. The clams metabolized very little DDT to DDE; the
principal metabolite was TDE.
The lUC-DDT content of the clam meat from the four exposures averaged 1*5.6+3.8
yg/g (n=lt) as determined by gas chromatography. This value indicates a
concentration factor of 25,000 times based upon the average measured DDT water
concentration. Total DDT in the clam tissue consisted of 68% DDT, 1% DDE, and
31% TDE.
Use of ll+C-labeled DDT in the contaminated food allowed the separation
of DDT contributed by the food by liquid-scintillation analysis (L.S.)
from the total amount of DDT as determined by gas-chromatographic
analysis (G.C.). Gas-chromatograph and liquid-scintillation results were
not identical as they should have been for the fish exposed to l^C-DDT-
contaminated food only. With n=19, the mean liquid-scintillation value
15
-------
TABLE 2. DDT, DDE, AND TDE RESIDUES (yg/g) IN FISH EXPOSED
TO VARIOUS TEST CONDITIONS (DUPLICATE SAMPLES COMBINED)2
Nominal
DDT water
concentration
(Pg/l)
c
F
0.5 W
0.5 W + F
2.0 W
2.0 W 4- F
Days of exposure
7
DDT
1.0
(0.5)b
16.2
(5.7)
26.1
(0.6)
47.4
(5.4)
94.2
(5.3)
92.8
(4.9)
DDE
0.7
(0.2)
12.0
(3.4)
24.9
(0.8)
39.0
(1.7)
66.8
(0.9)
50.3
(0.0)
TDE
0.1
(0.1)
0.0
0.0
0.0
0.0
0.0
14
DDT
1.0
(0.2)
28.2
(3.2)
18.4
(5.4)
22.5
(10.6)
48.5
(8.8)
77.8
(23.2)
DDE
1.0
(0.0)
24.1
(4.0)
24.3
(4.7)
29.5
(5.2)
88.3
(19.6)
54.7
(28 3)
TDE
0.0
1.0
(0.1)
0.0
1.4
(0.5)
1.4
(0.3)
3.8
(0.8)
28
DDT
0.6
(0.1)
23.8
(4.7)
7.2
(2.0)
35.9
(7.4)
37.0
(13.9)
43.3
(3.7)
DDE
1.0
(0.5)
35.3
(12.9)
25.5
(2.9)
53.7
(2.0)
92 . l)
(14.1)
113.4
(12.1)
TDE
0.0
1.1
(1.1)
0.7
(0.0)
3.1
(0.6)
4.0
(0.9)
7.4
(0.2)
56
DDT
0.2
(0.0)
18.5
(1.5)
13.2
(6.8)
17.7
(3.4)
25.1
(8.2)
44.6
(0.5)
DDE
0.3
(0.1)
49.4
(8.6)
18.8
(7.4)
63.3
(14.4)
86.6
(14.8)
149.2
(8.4)
TDE
0.0
1.0
(0.2)
0.6
(0.6)
1.4
(0.5)
1.7
(0.3)
3.1
(0.9)
112
DDT
0.1
(0.0)
7.0
(1.9)
6.1
(0.3)
14.2
(5.1)
58.0
(0.6)
49.8
(19.0)
DDE
0.2
(0.0)
38.5
(13.9)
36.8
(1.0)
120.2
(33.8)
230.9
(34.9)
246.6
(67.8)
TDE
0.1
(0.0)
15.0
(3.8)
3.7
(0.7)
20.4
(5.2)
19.3
(1.6)
52.2
(13.5)
224
DDT
0.1
(0.0)
5.2
(1.7)
3.5
(1.0)
4.9
(2.1)
9.5
(2.5)
18.0
(7.3)
DDE
0.1
(0.0)
47.5
(11.7)
36.7
(11.4)
41.7
(17.2)
93.6
(20.7)
113.3
(32.8)
TDE
0.0
12.5
(3.8)
0.8
(0.2)
7.9
(3.7)
9.1
(1.0)
6.2
(3.3)
266
DDT
0.0
5.0
(1.0)
2.7
(0.7)
7.5
(1.5)
15.8
(4.6)
28.5
(8.1)
DDE
0.1
(0.2)
33.3
(4.6)
21.7
(3.2)
54.1
(7.0)
136.9
(31.4)
291.7
(62.0)
TDE
0.0
22.4
(3.9)
1.0
(0.2)
18.2
(2.9)
3.8
(0.8)
17.1
(5.1)
Each sample is a composite of 5-10 fish.
OStandard error.
-------
TABLE 3. DDT, DDE, AND TDE RESIDUES (yg/g) IN THE FISH FOOD
Exposures
1
2
3
4
Time period when
used as food
(days)
0-112
112-224
112-224
224-266
DDT
14C-DDT
42. 63
38.2
31.2
11.0
DDE
-expose
0.6
0.5
0.2
0.2
TDE
d clair
14.3
2.7
9.7
29.0
Time of year
collected
is
Fall
Spring
Fall
Fall
Time of year
exposed to
14C-DDT
Spring (late)
Fall
Winter
Spring (early)
Clean clams
1
2
3
4
0-112
112-224
112-224
224-266
0.7
-
-
-
_b
-
-
-
-
-
-
-
Fall
Spring
Fall
Spring (late)
Fall
Winter
Fall j Spring (early)
Ovendried weight (1 1/2-2 hr; 110° C).
Not detectable.
17
-------
vas 110$ of the gas-chromatograph value with a standard deviation of 28.k%. To
correct for this difference L.S. values were adjusted to G.C. values by the
following formula: adjusted L.S. value = L.S. value obtained/ratio L.S. value
to G.C. value for F exposed fish. These data are presented in Table h and are
expressed as a percentage of the total residues as determined by gas-
chromatographic analysis. It appears that DDT in the food had a maximum input
within 28-56 days. The relative amount contributed by the DDT-contaminated
food was about 60% of the total DDT residues in fish exposed at 0.5 W + F and
about 30$ in fish exposed at 2.0 W + F.
Mean calculated accumulation of total DDT from food and water was 1.2 +_ 0.1
times for the DDT from the food (n=57), 99,000+J,000 times for DDT from the
water (n=39, 0.5 W and 2.0 W exposures combined), and 87,000+9,000 times from
the water (n=38, 0.5 W + F and 2.0 W + F exposures combined) after the food
contribution was substracted. If all water-exposure samples were combined,
a mean accumulation (n=77) of 93,000+6,000 times was obtained.
Lipid percentages were also determined for the test fish. Conversion of lipid
values to arcsin A/percent lipid and analysis by two-way analysis of variance
indicated that there was no significant difference (P=0.05) in percentage of
lipids between fish exposed to DDT in the water only and fish exposed to DDT
in the water and fed DDT-contaminated food.
Mortality results were analyzed by calculation of the accumulative probability
of fish survival over the different sample periods during the toxicity test.
This was accomplished by obtaining an estimate of the probability of a fish
surviving from time tj to tj given that it was alive at time tj and
exposure started at time t =0. This estimate is (l) Pj =Nj /nj, where
nj is the number of fish alive and exposed at time tj; it assumes that no
fish were removed during the interval. Since samples were taken at the end
of each interval, the probability of survival for each interval had to be
computed separately, and the probability (Ps) of survival for the entire
period is (2) Ps=7r=l Pj , where m is the number of sampling points. In Table
5 the computed probability of survival for each exposure is shown where m=7.
To test whether the addition of DDT-contaminated food altered the
probability of- 'survival, a two-way analysis of variance was run on the data
18
-------
TABLE 4. PERCENTAGE OF TOTAL DDT CAUSED BY THE
14C-DDT FOOD SOURCE (DUPLICATE SAMPLES COMBINED)
Nominal
DDT water
concentration
(pg/D
F
0.5 W + F
2.0 W + F
7 (n-2) .
100.0a(1.4)b
22.5 (0.0)
4.8 (2.1)
14 (n»2)
100. 0C
46.9 (19.4)
28.1 (15.7)
Day
28 (n=2)
100.0 (2.8)
76.4 (3.4)
38.2 (3.4)
s of exposure
56 (n=2)
100.0 (13.4)
81.7 (4.2)
50.7 (5.6)
112 (n=2)
100.0 (6.2)
55.4 (1.4)
33.9 (0.9)
244 (n»4)
100.0 (2.7)
72.2 (5.4)
28.7 (2.3)
266 (n=6)
100.0 (3.0)
64.7 (1.4)
17.6d(1.2)
Mean for
entire period
100.0 (1.6)
(n=19)
62.1 (4.2)
(n=20)
27.6 (3.4)
(n-18)
Liquid-scintillation values shown are adjusted.
( ) Standard error.
Cn=l.
dn=4.
TABLE 50 ESTIMATED PROBABILITY OF SURVIVAL FOR FISH EXPOSED TO
VARIOUS TEST CONDITIONS FOR 266 DAYS
Clean
food
14C-DDT
contaminated
food
Tank A
Tank B
Tank A
Tank B
Nominal DDT water concentration (pg/1)
0 1 0.5
0.8700
0.9052
0.6248
0.8860
0.7993
0.8697
0.7369
0.7392
2.0
0.5234
0.4211
0.1837
0.2541
19
-------
after arcsin ~\/Ps transformation was employed. The hypothesis that the
presence of DDT in the food does not change survival was rejected at the
P=0.025 level.
The accumulative reduction in the probability of survival is presented in
Figure k. Two definite periods of high mortality are indicated, the
juvenile stage during the first 28 days of exposure (test started with
about 1+5-day-old fish; therefore, the fish were 73 days old at 28
days' exposure) and at spawning time between 112 and 22H days of exposure.
Mortality was greater in fish fed DDT-contaminated food and it remained high
into 56 days' exposure before a plateau was reached, whereas the death, rate
among fish exposed at a corresponding DDT water concentration, but fed clean
food, reached a plateau at 28 days. The death rate was also greater
for fish fed DDT-contaminated food during the spawning period. Fish
that died during the spawning period were predominantly highly colored
adult males (about 83% males, 1.1% females). Residue levels in the dead fish were
only about 55$ of those in live sampled fish. Lipid percentages, however, were
very low (mean 1.07$ (n=23), all samples combined) when compared to the live fish
at the corresponding time period (mean 3.1$ (n=12), all samples combined).
The residue levels attributed to DDT in the food among fish that died was
all the DDT at the F exposure, 70% at the 0.5 W + F exposure, and 23$
at the 2.0 W + F exposure. These percentages are similar to those
found in the similarily exposed live fish.
EMBRYOS
Embryo-hatchability data are presented in Table 6. Data were transformed to
arcsin "Vpercent hatch and analyzed by two-way analysis of variance. Presence
of DDT in the parents' food did not significantly alter hatchability,
whereas DDT in the water did (P=0.05). Hatchability reduction, however,
was significant only for embryos from parent fish that were subjected to the
2.0 W exposure. Table 7 shows the total DDT residues at the various exposures
and also the percentage DDT from food for embryos from adult fish fed the DDT-
contaminated food. The percentages observed are similar to those for the
adult fish. Presence of DDT-contaminated food appears to be additive.
Addition of the residues found in embryos from fish at the F exposure to
residues found in embryos from fish at the 0.5 W and 2.0 W exposures will
20
-------
80
£ 60
_J
DO
O
o:
£L
LU 40
^^
1
20
• 2.O W
• 2.0 W+F
50 IOO 150 2OO 250
TIME (days)
Figure U. Accumulative pro"ba"bility of survival for fish that had
been exposed to DDT.
21
-------
TABLE 6. HATCHABILITY OF EMBRYOS AT VARIOUS TEST EXPOSURES
(DUPLICATE SAMPLES COMBINED)
NJ
Nominal
DDT water
concentration
(yg/D
c
F
0.5 W
0.5 W + F
2.0 W
2.0 W + F
Na
62
44
51
55
98
51
Number of
eggs set up
3,473
2,610
2,550
3,491
6,759
3,117
Number of
eggs hatched
3,089
2,241
2,152
3,114
5,009
2,440
Percentage
hatch
88.9
85.7
84.4
89.2
74.1b
78.3
Range
(percentage hatch)
54-100
38-100
46-100
48-100
12-100
44-100
Arc sin
-\/percentage hatch
70.54
67.78
66.74
70.81
59.41
62.24
rt
N=number of spawnings.
bSignificantly different from the control (aO.05) Dunnett's procedure (Steel and
Torrie, 1960).
-------
TABLE 7. TOTAL DDT RESIDUES (pg/g) IN EMBRYOS FROM FISH EXPOSED TO VARIOUS
TEST CONDITIONS, AND PERCENTAGE TOTAL DDT CONTRIBUTED
BY THE CONTAMINATED FOOD (DUPLICATE SAMPLES COMBINED)
Nominal DDT water
concentration (yg/1)
C
F
0.5 W
0.5 W 4- F
2.0 W
2.0 W + F
Number of
samples
15
12
16
15
23
18
Mean (yg/g)
0.4 (0.0)a
12.0 (1.1)
6.7 (0.7)
18.9 (3.1)
24.0 (2.0)
40.9 (3.8)
Range (yg/g)
0.1-0.6
6.4-18.8
3.3-11.9
5.4-43.8
12.2-50.4
21.9-78.0
Percentage due
to DDT in
the food
Liquid
scintillation
adjusted
-
100 (6.2)
-
68 (1.8)
-
28 (0.5)
( ) Standard error.
give residue levels close to those for embryos from fish at the 0.5 W + F
and 2.0 W + F exposures. Residues in the embryos from fish exposed to DDT
in the food and in the water are two times greater than in those from fish
exposed to DDT only in the water.
To determine the relative amount of DDT that might be transferred to the
embryo from the fish, control embryos were placed in the 2.0 yg/1 DDT
water exposure for 24 hr (embryos would not have been exposed to DDT in the
water any longer than this before collection for residue analysis). These
embryos had a residue level of 0.95 yg/g, which would probably be the maximum
that embryos spawned in this tank could have attained from the water. Residue
levels found in the latter embryos were much higher; therefore residues
found were mostly transferred from the adult fish.
23
-------
Separation of total DDT to DDT, DDE, and TDK is shown in Table 8. DDE
was the primary constituent. Some DDT from the water was metabolized
to TDE, but the amount is only about 10% of that attributed to the food
source. There is increased TDE with higher water exposure for the
embryos from the water plus food-exposed fish. Mean accumulation of DDT
was 0.26+0.02 times (n=45) from the food source for embryos from parent
fish at the F exposure, 17,000+1,000 times (ri"39) from the water for those
from parent fish exposed at 0.5 W and 2.0 W, and 19,000+2,000 times Cn-331 for
those from fish exposed in combination, to 0.5 W + F and 2.0 W + F after the
residues due to food were subtracted.
LARVAE AT HATCH
Total DDT residue data and percentages of residues caused by DDT-contaminated
food are presented in Table 9. When compared with embryo residues,
residues in larvae at hatch from parent fish fed dietary DDT are about
two times higher, whereas those from parent fish exposed to DDT only in
the water are about 3.6 times higher. Part of this difference is
explained by the reduction in weight of the larvae at hatch to almost one-half
that of the embryos. Therefore, if nearly all the residue were contained within
the developing larva, the calculated residue level would automatically be two
times greater at hatch when the embryo membrane and surrounding fluid is lost.
The percentage of total DDT resulting from DDT-contaminated food was about
8% lower for larvae at hatch than for embryos. Larvae were removed for analysis
daily, so it is possible that some may have been exposed to DDT in the water
for a maximum of 24 hr before being removed.
Separation of total DDT to DDT, DDE, and TDE is shown in Table 10. DDE
again is the principal constituent. All residues are greater than those
observed for embryos. High TDE levels in larvae from food-exposed parents
are attributed to the food source. Mean accumulation of DDT in larvae
from fish exposed to the two DDT sources was 0.53+0.03 times (n=5l6)
for dietary DDT (double that for embryos), 62,000+4,000 times (n~13)
for DDT water exposure alone, and 50,000+5,000 times (n=ll) for
exposure to DDT in the water minus that contributed by the food.
24
-------
TABLE 8. DDT, DDE, AND TDE RESIDUES (yg/g) IN EMBRYOS FROM FISH EXPOSED
TO VARIOUS TEST CONDITIONS (DUPLICATE SAMPLES COMBINED)
Nominal DDT
water concentration (yg/1)
C
F
0.5 W
0.5 W + F
2.0 W
2.0 W + F
DDT
0.08 (O.O)3
1.56 (0.1)
0.75 (0.1)
1.99 (0.4)
3.83 (0.5)
5.31 (Oo4)
DDE
0.35 (0.0)
7.76 (0.9)
5.70 (0.6)
13.66 (2.2)
19.61 (1.7)
31.32 (2.9)
TDE
_b
2.75 (0.2)
0.30 (0.0)
3.26 (0.6)
0.58 (0.1)
4.24 (0.6)
( ) Standard error.
Not detectable.
TABLE 9. TOTAL DDT RESIDUES (pg/g) IN LARVAE AT HATCH FROM FISH EXPOSED TO
VARIOUS TEST CONDITIONS, AND PERCENTAGE TOTAL DDT CONTRIBUTED BY
THE CONTAMINATED FOOD (DUPLICATE SAMPLES COMBINED)
Nominal DDT
water
concentration (yg/1)
C
F
0.5 W
0.5 W + F
2.0 W
2.0 W + F
Number
of
samples
6
4
4
6
9
5
Mean
0.17 (0.01)a
26.4 (2.9)
24.0 (2.2)
43.5 (4.3)
87.9 (5.9)
96,8 (17.8)
Range
0.11-0.19
19.1-31.5
17.8-27.2
33.5-62.4
54.0-113.2
65.4-166.5
Percentage due
to food
Liquid
scintillation
adjusted
-
100 (0.8)
-
60 (1.9)
-
20 (0.6)
( ) Standard error.
25
-------
N>
TABLE 10. DDT, DDE, AND TDE RESIDUES (yg/g) IN LARVAE AT HATCH FROM
FISH EXPOSED TO VARIOUS TEST CONDITIONS (DUPLICATE SAMPLES COMBINED)
Nominal DDT
water
concentration (pg/1)
C
F
0.5 W
0.5 W + F
2.0 W
2.0 W + F
DDT
a
2.96 (0.3)
3.77 (0.3)
5.70 (0.5)
19.85 (0.9)
20.11 (2.0)
DDE
0.17 (0.0)b
18.63 (2.6)
18.89 (2.3)
32.77 (4.5)
66.49 (5.3)
72.99 (15.4)
TDE
a
4.83 (0.2)
1.32 (0.0)
4.99 (0.6)
1.53 (0.1)
2.89 (0.7)
Not detectable.
( ) Standard error.
-------
PROGENY AT 30 AND 60 DAYS
Residue data and tissue-residue percentages caused by DDT-contaminated food
are shown in Table 11. The additive DDT food effect is again indicated.
Residues are slightly higher at 60 than at 30 days, except for fish at
the F exposure, where they are lower. The percentage of total DDT caused by
the food source is only 8% higher at 60 days than at 30 days. Progeny from
embryos from fish exposed to 2.0 W hatched and raised in control water for
30 days contained only 0.5 yg/g total DDT in their tissues. Progeny from embryos
from fish exposed to 2.0 W + F, also hatched and raised in control water and
fed clean food for 30 days, had residues of only 0.2 yg/g total DDT. As
determined by liquid-scintillation analysis, none of this DDT could be traced to
the DDT-contaminated food intake by parent fish. Progeny, however, from embryos
from the same parent fish that were hatched and raised in clean water but fed
DDT contaminated food for 30 days had total DDT residues of 31.6 yg/g, of which
93% could be attributed to the food.
Residues in 60-day progeny are not much different from residues in the
corresponding parent fish at 14 days' exposure (fish were ~59 days old) if
calculated on a percentage lipid basis. These values are as follows: 60-day
progeny fed DDT food, 8.41 yg/g, parent fish at 59 days old, 14.4 yg/g;
0.5 W 60 days, 7.1 yg/g, parent fish 59 days old, 10.7 yg/g; and 0.5 W + F
60 days, 17.9 yg/g, parent fish, 59 days old, 21.8 yg/g.
Separation of total DDT residues to DDT, DDE, and TDE are presented in
Table 12. In general, DDT residues decreased at most exposures between
30 and 60 days, whereas DDE residues increased. TDE residues also showed
a slight increase between 30 and 60 days, except at the 0.5 W + F
exposure where there was a 38% increase.
Mean accumulation of total DDT from the food was 0.70jf0.13 times
(n=8) for 30-day progeny and 0.75+0.09 times (n=4) for 60-day progeny.
Total DDT in the .water was accumulated 39,000+5,000 times (n=4)
for 30-day and 70,000+12,000 times (n=5) for 60-day progeny exposed
to DDT only in the water, whereas DDT in the water was magnified 70,000+
27
-------
TABLE 11. TOTAL DDT (ug/g) IN 30- AND 60-DAY-OLD PROGENY OF FISH EXPOSED TO
VARIOUS TEST CONDITIONS, AND PERCENTAGE OF TOTAL DDT CONTRIBUTED BY THE
CONTAMINATED FOOD (DUPLICATE SAMPLES COMBINED)
(a) 30-Day-old progeny
Parent fish
nominal
DDT water
concentration (pg/1)
C
Fb
0.5 W
0.5 W + Fb
2.0 W
2.0 W + F°
2.0 W + Fd
Number
of
samples
3
4
4
4
3
1
1
Mean
0.20 (O.O)3
35.70 (1.6)
13.70 (1.8)
46.4 (4.9)
0.5 (0.2)
0.2
31.6
Range
0.16-0.23
31.50-39.10
10.00-17.90
37.20-60.20
0.33-0.93
-
Percentage
caused by DDT
in the food
Liquid
scintillation
adjusted
-
100.0 (9.4)
-
60.0 (16.3)
-
0.0
93.0
( ) Standard error.
Progeny fed DDT-contaminated food.
p
Progeny hatched and raised in control water and fed clean food.
Progeny hatched and raised in control water and fed DDT food.
(b) 60-Day-old progeny
Parent fish
nominal
DDT water
concentration (pg/1)
C
Fb
0.5 W
0.5 W + Fb
Number
of
samples
3
2
5
2
Mean
0.21 (O.O)3
28.60 (5.7)
24.00 (4.0)
58.20 (2.4)
Range
0.15-0.30
22.9-34.2
14.3-38.1
55.8-60.6
Percentage
caused by DDT
in the food
Liquid
scintillation
adjusted
-
100 (4.1)
-
68 (0.4)
( ) Standard error.
Progeny fed DDT-contaminated food.
28
-------
N>
1C
TABLE 12. DDT, DDE, AND TDE RESIDUES (yg/g) IN 30- AND 60-DAY-OLD PROGENY OF
FISH EXPOSED TO VARIOUS TEST CONDITIONS (DUPLICATE SAMPLES COMBINED) (LARVAE
WERE EXPOSED TO SAME DDT EXPOSURE AS PARENT FISH)
Nominal DDT water
concentration
(M8/D
C
F
0.5 W
0.5 W + F
Progeny
exposure (days)
30
60
30
60
30
60
30
60
DDT
0.08 (0.01)a
0.08 (0.01)
8.07 (2.00)
4.72 (0.40)
4.64 (0.90)
6.08 (0.40)
12.51 (3.30)
10.34 (1.00)
DDE
0.08 (0.01)
0.20 (0.04)
15.38 (2.90)
11.73 (0.70)
8.54 (0.90)
17.34 (3.50)
18.60 (3.10)
26.83 (0.50)
TDE
0.04 (0.04)
12.21 (3.20)
12.33 (5.80)
0.51 (0.20)
0.55 (0.20)
15.27 (2.40)
21.08 (0.90)
( ) Standard error.
-------
2,000 times (n=3) for 30-day and 51,000+4,000 times (n=2) for 60-day progeny
exposed to DDT in the water after the food contributed portion was substracted.
DDT apparently did not affect the growth of the 30- and 60-day progeny.
Two-way analysis of variance on growth data indicated that there was no
significant growth difference (P=0.05) between the progeny exposed to
DDT in the food or water.
All larvae died within 5 days of hatch at the 2.0 W or 2.0 W + F
exposures. Control larvae, in groups of 40 each (n=4), transferred
to these concentrations also died within 5 days. Survival
data (Table 13) were transformed to arcsin ~V percent survival
and analyzed by two-way analysis of variance. A significant effect
(P=0.05) on survival was observed for DDT water exposure at both 30
and 60 days, but not for food exposure. Use of Dunnett's procedure
(Steel and Torrie, 1960) indicated that the exposures significantly
different from the controls were those at 2.0 W and 2.0 W + F and
those from parent fish exposed to 2.0 W + F, but with the progeny
hatched and raised in control water and fed clean food for 30 days.
This latter group experienced a twofold higher death rate than
progeny from parent fish exposed at 2.0 W that were hatched and raised in
control water and fed clean food. In this higher mortality group, progeny
in two groups (40 fish each) experienced zero survival and one group had 75%
survival. Total DDT residue in the survivors was 0.18 ug/g. Another
group of progeny from the same parent fish similarily hatched and raised
in control water but fed DDT-contaminated food had only 19.5% survival
with a residue level in survivors of 31.6 yg/g total DDT. Ninety-three
percent of this residue level could be directly attributed to their DDT-
contaminated food intake.
ELIMINATION STUDY
Elimination of total DDT from fathead minnows exposed to 0.5 W and 0.5 W + F
is shown in Figure 5. Relatively little elimination occurred in fish exposed
to 0.5 W, whereas a definite elimination (significant at the 0.10 level)
occurred in fish exposed at 0.5 W + F. In these fish about 60% of the mean
total micrograms of total DDT was lost within 56 days.
30
-------
TABLE 13. PERCENTAGE SURVIVAL OF 30- AND 60-DAY OLD
PROGENY FROM PARENT FISH EXPOSED TO VARIOUS TEST CONDITIONS
(LARVAE EXPOSED TO SAME DDT EXPOSURE AS PARENT
FISH) (A) 30-DAY SURVIVAL (B) 60-DAY SURVIVAL
(A)
Clean food
DDT-contaminated food
Tank a
Tank b
Grand mean
Tank a
Tank b
Grand mean
Nominal DDT water concentration (pg/1)
0 0.5 2.0a 2.0
73.8 (n=4)
55.8 (n=3)
66.1
73.0 (n=3)
69.9 (n=3)
71.2
55.0 (n=4)
65.6 (n=4)
60.3
57.5 (n=3)
36.7 (n=3)
47.1
25.0 (n=2)
62.3 (n=2)
41.0
23.6 (n=3)
0 (n=l)
18.8b
0 (n=3)
0 (n=3)
ob
0 (n=2)
0 (n=2)
ob
F values 3.8 df
1.8 df
4.07 F cal DDT water = 23.83
5.32 F cal DDT food =2.26
Progeny from parent fish were hatched and raised in clean water and fed clean food.
Values significantly different from the control larvae (duplicate chambers combined),
two-way analysis of variance and Dunnett's procedure (Steel and Torrie, 1960) n=8;
ta = 0.05 Dunnett's - 7,8 df 30 days.
5,6 df 60 days
(B)
Clean food
DDT-contaminated food
Tank a
Tank b
Grand mean
Tank a
Tank b
Grand mean
Nominal DDT water concentration (pg/1)
0
46.3 (n=2)
45.0 (n=l)
45.8
62.5 (n=l)
51.3 (n=l)
57.0
0.5
31.2 (n=2)
66.3 (n=2)
49.0
35.0 (n-1)
42.5 (n=l)
38.8
2.0
0
0
0
0
0
0
31
-------
fe200
o
I
o>
3
o
o
o
0 20 40
TIME (days)
Figure 5. Elimination of total DDT from fish exposed at 0.5 W
(•) and 0.5 W + F (<*'). Each point represents one fish.
32
-------
Liquid-scintillation analysis of the fish exposed to DDT-contaminated
food is shown in Table 14. Residues caused by food and water are expressed
as a percentage of the total DDT. With longer elimination time significantly
less total DDT remains that is attributed to the food, and the water
contribution becomes correspondingly greater.
Separation of total DDT to DDT, DDE, and TDE and subsequent regression
analysis of elimination data are presented in Figures 6-8. A significant
reduction in DDT and TDE occurred for fish at 0.5 W + F, but not for
those at 0.5 W (P=0.05). In the latter group DDT was metabolized
slightly and TDE levels remained unchanged. Essentially no DDE was
eliminated in the 0.5 W-exposed fish, whereas elimination did take place in
fish exposed at 0.5 W + F. Comparison of the TDE residues between the two
groups of fish indicates that almost all the TDE was from the food source.
33
-------
TA3LE 14, MEAN PERCENTAGE OF TOTAL DDT RESIDUES REMAINING IN FISH (EXPOSED TO
A COMBINATION OF DDT IN FOOD AND WATER) THAT WAS ATTRIBUTED
TO THE FOOD OR TO THE WATER
U)
Percentage due to DDT food uptake
Liquid scintillation adjusted
Percentage due to DDT water uptake
100-Liquid scintillation adjuste
Days after placed in control water
0
n=6
64.7 (1.4)a
35.3
7
n=5
68.7 (4.7)
31.3
14
n=5
74.2 (7.0)
25.8
28
47. f (3.1)
52.5
56
n=4
46.0b(1.5)
54.0
( ) Standard error.
Significantly different from 0 days (ANOVA and Dunnett's procedure;
P=0.05; F=2.9; F cal=9.1).
-------
40
30
O
0
20 40
TIME (days)
Figure 6. Elimination of DDT from fish exposed at 0.5 W (•) and 0.5 W + F (0)
Each point represents one fish.
35
-------
150
UJ
Q
Q
l
o»
100
50
O
O
O
O
O
0 20 40
TIME (days)
Figure 7. Elimination of DDE from fish exposed at 0.5 W (•) and 0.5 W + F (0).
Each point represents one fish.
36
-------
60
L±J
Q
I-
40
O
O
O
0
20 40
TIME (days)
o
Figure 8. Elimination of IDE from fish exposed at 0.5 W (•) and 0.5 W + F (0)
Each point represents one fish.
37
-------
SECTION VI
DISCUSSION
The fathead minnows did not achieve an equilibrium with DDT ^rom the
water at 2.0 pg/1, whereas they apparently did achieve equilibrium at 0.5
Mg/1. Lack of equilibrium was also reported by Hamelink et al. (1971) for
young-of-the-year largemouth bass exposed to DDT at 50 vig/1 and greater for
up to 80 days. However, this does not necessarily mean that the fish did not
approach an equilibrium with the toxicant at any instant in time. Dior ing
long-term studies factors such as lipid content, toxicant, stress, etc. can all
be expected to influence residue concentrations. Therefore the fish probably
did achieve equilibrium with the DDT in the water at various times during the
test. Greater residue fluctuations at the higher DDT exposure in our test
may have occurred because the fish were closer to their maximum accumulative
capabilities, and any change in body conditions would be more directly
reflected in residue levels. Our results indicate that uptake from the food
is additive to the amount taken up from the water and that equilibrium with
the food was reached within 56 days. These findings are similar to those
observed by other authors. Grzenda _et_ aJ^. (1970) reported that there was no
additional increase in body concentration in goldfish after dietary DDT
exposure (l8 yg/g) for 32 days. Macek et al. (1970) reported an equilibrium
with dietary DDT in the liver, brain, and skeletal muscle of rainbow trout
after 28 days' exposure. The portion of the total DDT tissue residues in
our test that could be directly attributed to the DDT-contaminated diet
was about 30% in fish exposed to a DDT water concentration equal to that at
which the food had been exposed and 60% in fish exposed to a water concentration
one-fourth that level. This appears to indicate that the main source of DDT
uptake is the water. Chadwick and Brocksen (1969) exposed freshwater sculpins
to dieldrin in diet and water and observed that a maximum of 16% of the
dieldrin was accumulated from the food. They also stated that accumulation
from the diet might be expected to be additive, but this was not so in their
38
-------
test. They reported residue levels that were not much different between fish
exposed in food and water and in water only, whereas in our test tissue
residues between the two exposures were different after llj days. Reinert (1967),
also in a study with dieldrin, observed that only about one-tenth as much of the
residues in guppies were accumulated through ingestion of contaminated food
when compared to residues in guppies exposed to a water concentration equal to
that in which the food was exposed. This test, however, was of a shorter
duration (32 days). In our test the highest residue level caused by DDT-
contaminated food was one-fourth that caused by DDT water exposure at the same
concentration to which the food was exposed.
Our mean DDT concentration factor was about 1.2 times for dietary DDT and
about 100,000 times for DDT from the water. These values are similar to
those observed by other authors. Hunt and Bischoff (i960) observed a 125,000
times concentration factor for brown bullheads from water, and Courtney and Reed
(1972) observed a concentration in the tissues of golden shiners exposed to
DDT at 0.3 yg/1 in the water of about 100,000 times after 15 days. Reinert
(1967) observed a 0.05 times food concentration factor in tissue residues of
guppies exposed to a diet of dieldrin-contaminated Daphnia (31 yg/g) for 32
days, but a concentration factor of 1.3 times for Daphnia fed dieldrin-
contaminated algae (71 yg/g). He stated, however, that the latter concentration
factor may have been caused by algae ingested but not yet assimilated, as alga
cells were observed in the Daphnia digestive tract. Epifanio (1973) observed
a concentration factor of 1.7 times when he fed dieldrin-contaminated Artemia
salina brine shrimp nauplii (0.213 yg/g) to crab larvae. Macek and Korn (1970)
observed a food concentration factor of 0.6 tiroes for brook trout exposed to
3 yg/g DDT in their diet for 120 days. Grzenda _e£ all, (197Q) estimated a total
mean body residue level of 14.2 }jg/g DDT in goldfish after they were fed a
diet containing 17.7 yg/g DDT for 192 days, which would indicate a concentration
factor of 0.8 times.
Some lUC-DDT that leached from the food into the water was observed by liquid-
scintillation, and a mean ihC-DDT water concentration of 0.065+0.007
yg/1 (n=12) was determined. If this level was bioaccumulated 100,000 times,
the highest level that could occur in the fish would be 6.5 yg/g, or about
10% of the total food-contributed residue. However, the total DDT
39
-------
concentration measured in the water "by gas-chromatographic analysis was
not much greater where the fish were exposed to DDT-contaminated food only.
If the control fish with a mean measured water concentration of 0.0029 yg/1
DDT are compared to fish exposed only to DDT in the food (F) with a mean
measured DDT water concentration of 0.0123 ug/1, the difference is -only
0.009 yg/1, a possible body residue of 0.9 yg/g or about 1.5$ of
residues caused by DDT in the water leached from the food (assuming a
100,000 times magnification). Therefore, we believe that the contribution
of labeled DDT through the water was negligible.
Our results indicate that DDT in the tissues decreased between ih and 266
days. During the same period DDE and TDE residues increased. Grzenda
et al. (1970) observed similar results for goldfish fed a DDT-contaminated
diet for up to 192 days. We also observed in our test that the TDE residues
were high in fish exposed to dietary DDT and were low in fish fed clean food.
This indicates that most of the TDE in the fish fed a contaminated diet
probably came from the food itself, which was quite high in TDE.
The clams metabolized DDT almost entirely to TDE and produced little DDE.
Some discrepancy was observed in the amounts of TDE produced between
exposures. Although all clam DDT exposures were held at the same temperature
(20° C), clams for exposures 1 and k were collected in the fall and exposed
in the spring, whereas clams for exposures 2 and 3 were collected in the
spring and fall and exposed in the fall and winter. More clam metabolism
would naturally occur in the spring than in the winter, and a seasonally
controlled mechanism might be responsible for metabolite differences among
the DDT-exposed clams. Another possible explanation is that differences
could have been caused by different ratios of clam species present. No check
was made as to the relative frequency of each species, although the genus
Lampsilis appeared to be the one most prevalent. The 1 yg/g DDT found in
the control fish was probably caused by the presence of 0.7 yg/g DDT in
the first batch of clean clam tissue fed to the fish. This explanation is
likely since the use of this clam food and the residue peak in the control fish
terminated at about the same time.
The proportion of TDE and DDE produced appears to vary with the.type of organism
exposed. Rats convert most DDT to TDE, whereas- humans usually metabolize DDT
40
-------
to DDE (O'Brien, 1967). Mollusks, as observed in this test and by Cooke
and Pollard (1973), metabolize DDT mainly to IDE, whereas fish metabolize
DDT essentially to DDE (Priester, 1965; Johnson and Pecor, 1969; Reinert and
Bergman, 1974). Ferguson et al. (1967), however, observed almost equal amounts
of DDT, DDE, and IDE in several pooled samples of resistent mosquitofish
(Gambusia affinis). Therefore, it is apparent that some fish can also
readily produce large amounts of TDE.
Some breakdown, of DDT to TDE may have occurred in our test when the clam
tissue was ovendried or when it was in frozen storage. Breakdown of
DDT to TDE in frozen storage was demonstrated by French and Jefferies
(1971). Our clam tissue, however, was used within 60 days, so post-mortem
breakdown is believed to have been negligible. Ovendrying also is not
believed to have caused much DDT breakdown. Smith et_ al. (1973) found
no large change in total TDE after fish steaks were baked at 177° C, and
Metcalf (1955) stated that pure DDT is stable up to 195° C.
Lipid values could be correlated with residue values, an indication that
DDT uptake was influenced by the lipid content of the fish. DDT-residue
levels declined rapidly during the spawning period. Lipid content of the
fish also decreased at this time in fish exposed to the 2.0 yg/1 DDT
water concentration whether they were fed clean or DDT-contaminated food.
This is probably a fairly common occurrence. Reinert and Bergman (197*0
observed that DDT residues were redistributed in the tissues of spawning-
run fish and that this redistribution was closely related to a general
decrease in the amount of fish oil.
The presence of DDT in the diet significantly reduced the probability
of survival for exposed fish. Two definite periods of death were
observed, the early larval stage up to 73 days of age and the
spawning period, when highly colored males were the most sensitive. Fish
that died at the spawning period were in relatively poor condition and did
not feed. They probably used their fat reserves, thereby causing a release
of stored DDT into the blood where the DDT could become toxic. Holden
(1962) stated that fish in poor condition or with low fat content were more
41
-------
susceptible to DDT toxicity. Our results support Holden's statement in that fish
that died had predominantly lower lipid values than live fish sampled at the
same time. Redistribution of DDT to the brain during weight loss with
resultant deaths has been observed by some authors. Dale _et_ al. (1962)
observed high brain levels of DDT in rats that were starved after being
fed DDT. Transfer of DDT to brains of birds and some resultant deaths were
observed by Bernard (1966), Ecobichon and Saschenbrecker (1969), and Van
Velzen _et_ al_. (1972). Redistribution of DDT and fat depletion in salmon and
trout were observed during the spawning run by Holden (1962) and Reinert and
Bergman (197M and in the laboratory in starved rainbow trout by Grant and
Schoettger (1972)- Desaiah et_ al. (1975) analyzed fish sampled at 56, 118,
225» and 266 days during our toxicity test and found that partial chronic
9 +
exposure to DDT significantly inhibited mitochondrial Mg^ ATPase activity
in the brain tissue of the fathead minnows.
In regard to toxicity in relation to the high concentrations of TDE in
the food, the oral toxicity of TDE to rats (U.S. Environmental Protection
Agency, 1972) is about 1/3 that of DDE. In bluegills, however, the static
acute data of Cope (1965) and Mayer (personal communication) indicate that
TDE is more toxic than DDE. Probably the relative toxicity of each varies
with the specific organism exposed. Both DDE and TDE, however, appear to be
less toxic than DDT (Oettingen and Sharpiess, 191*6; Rudd and Genelly, 1956;
O'Brien, 1967; Moyle and Skrypek, 1969). Therefore, even though our clam
tissue had high levels of TDE, our mortality results present a more accurate
example of the DDT effect in a normal aquatic food chain than if we had used
the more toxic unmetabolized DDT in a dry food mix.
To estimate the relative contribution of total DDT from the food to DDT from
the water in regard to its effect on mortality the following model was
assumed: the log of the total dose that a fish can tolerate has a normal
distribution which can be approximated closely by a logistic distribution.
Thus we have that (l) ln(P/l-P) = y+31nx, where x is the total dose delivered
to the fish and P is the probability of death given a dose at x. It was
further assumed that the water and food DDT act on the fish in a similar
manner and thus x is proportional to the sum of water and food exposure and
is expressed as (2) x=§ (xj+-px2), where 6 is a magnification and absorption
42
-------
factor in the target organ, xi=concentration of DDT in the water, X2=l if DDT
is in the food, zero if otherwise, and. P is the unknown amount of DDT ingested
from the food. Substituting the value for x into equation (l) we have that
(3) ln(P/l-P) = y+31n (6(x1+px2)) = a+gln (x!=px2), where a=Y+Bln6 is a
combination of parameters that is estimatable from the data. The probability
of death given a target dose of x may be estimated from our data utilizing
Abbott's formula from the relationship Ps=(l-Po) (l-P), where Po is the
probability of death of a control fish. Thus P=l-Ps/(l-Po) is an estimated
probability of death due to DDT, where Ps and 1-Po are taken as the average
values in Table 5- Table 15 shows these estimated P values, the actual mean
measured DDT water concentrations in duplicated exposure chambers, and the
values of the estimated parameters obtained using non-linear least square
estimation. The value e~a/3 is an estimate of LC50 caused by total exposure.
It is estimated as 1.1*596 yg/1 in DDT water with no DDT in the food, or 0.9270
yg/1 in DDT water with 1*5.6 yg/g DDT in the food. The amount of DDT in the
food delivered to the target organism in this study is estimated as 0.5325
units in water. Thus the percentage of DDT in the fish from DDT in the
contaminated food in the 0.5 and 2.0 yg/1 DDT water exposures is estimated
as 58.8 and 26.h, respectively. These estimates are very close to the measured
percentages determined by liquid-scintillation analyses as shown in Table h.
Figure 9 shows the relationship between probability of death and DDT in the
water with and without DDT in the food. At low DDT water concentrations the
importance of the DDT-contaminated food is greater, and as DDT water concentration
increases the importance of the food effect decreases. Dunnett's procedure
(Steel and Torrie, I960) was used to compare the combined duplicate test
exposure mean probability of survival with that in the controls. With P=0.05,
reduction in the probability of survival was significant only at the 2.0 W and
2.0 ¥ + F exposures. By back calculation it is estimated that reduction in
the probability of survival to 6k*5% (35-5% mortality) is necessary to be
significantly different,. From Figure 9 it can be seen that this mortality
level would fall at about O.*l25 yg/1 DDT in the water plus DDT in the food and
at 0.925 yg/1 DDT in the water without DDT in the food.
Our mortality data indicate that the presence of dietary DDT is more important
when DDT water concentrations are low. A similar situation in nature could
easily occur if food organisms built up tissue residues during pesticide
43
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TABLE 15. ESTIMATED P VALUES, AVERAGE MEASURED DDT WATER CONCENTRATIONS, AND
VALUES FOR ESTIMATED PARAMETERS FROM NON-LINEAR LEAST SQUARE ESTIMATION
FOR FISH EXPOSED TO VARIOUS TEST CONDITIONS
P
Estimated probability
of death caused by DDT
0.0598
0.4680
0.1489
0.1685
0.7534
ln(P/l-P)
-2.7550
-0.1280
-1.7430
-1.5960
1.1170
Xi
Average measured
DDT in the water (ug/1)
0.3517
1.5285
0.0123
0.3740
1.4839
X2
(0 if no DDT
(1 if DDT in
0
0
1
1
1
in food)
food)
= a+
Parameters
a
6
p
Estimates
of
parameters
-0.7661
2.0261
0.5325
Standard deviation
of
estimates
0.3802
0.4313
0.1946
44
-------
DDT in food
•
^"
No DDT in food
.8876
|+e_[- .76614 + 2.0261 In (x, + .5325 x2)J
x,» DDT water concentration
x2 -1 if DDT in the food
« 0 if no DDT in the food
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0
DDT WATER CONCENTRATION (ug/l)
Figure 9. Estimated probability of death for fish exposed to DDT at various
water concentrations and fed clean or DDT-contaminated food.
45
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contamination and then were eaten by predators that entered the area after
pesticide water concentrations had moderated, or if several successive food-chain
accumulations occurred. Dietary DDT exposure is important at relatively higher
water concentrations, although to a lesser degree than water exposure. This view
is also held by other investigators, such as Grzenda et_ al_. (19TO), who stated
that apparent DDT biological magnification by the food chain may not be as
significant as the length of time fish are exposed to pesticide residues in
the water; Reinert and Bergman (197M, who stated that rapid accumulation
of DDT residues in coho salmon was probably related in part to an increase in
the intake of DDT-contaminated alewifes; and Macek and Korn (1970), who
demonstrated that dietary DDT can be very important, especially when compared
with very low DDT water concentrations such as are found in Lake Michigan.
Embryo hatchability was reduced significantly only when parent fish were
exposed to DDT at 2.0 pg/1 in the water and fed clean food. However, since
DDT tissue residues were twice as high in embryos from fish exposed to DDT
both in the diet and at 2.0 yg/1 in the water, the significant reduction of
hatchability in the former group and not in the latter may have been the result
of adult fathead minnow variability. Residues caused by dietary DDT were
additive for all embryos from parent fish exposed to DDT in water and food.
Huisman _et_ al. (l9?l) reported that high residues of DDT, DDE, and TDE were
accompanied by low fertility in pike. Kleinert and Degurse (1973) demonstrated
a strong correlation between DDT content of embryos and larvae from walleyes in
Wisconsin lakes, but could not associate the presence of DDT with success of
the hatch. Burdick et al. (1972) demonstrated loss of brown trout and brook
trout larvae hatched from eggs taken from female fish fed dietary DDT at
different concentrations and durations of time. They also confirmed DDT as
the cause of 100% mortality of larvae reared from eggs from lake trout fed 6 yg
of DDT per gram of body weight per week. Average DDT egg residues less DDE
were 7.6l and 11.92 yg/g for 2 separate years. In our test mean total DDT
embryo residues less DDE were ^.31 yg/g from fish fed dietary DDT and 9-55 yg/g
from fish fed dietary DDT and also exposed to DDT in the water. Lack of 100%
larval mortality at these levels is probably explained by species variability,
as fathead minnows are generally less susceptible to DDT than salmonids. Larvae
from fish exposed at 2=0 W and 2.0 W + F reared at the same water concentration
as the parent fish experienced 100% mortality within 5 days. Part of this
46
-------
mortality may have teen caused by DDT sorbed from the water, as larvae from
these two parental groups transferred to control water for 30 days
demonstrated an average of only 59% mortality among those from parents
exposed to DDT in the water only and greater than 8l% mortality among those
from parent fish exposed to DDT in both the water and diet, an almost twofold
higher mortality. These mortality data agree closely with embryo residue results
that indicated almost twice as much DDT residue (1*0.9 vs. 2U-0 pg/g) in embryos
from parent fish exposed to DDT in both water and diet. Although residue levels
were high, the larvae could either readily eliminate the DDT or dilute the
residues through growth if placed in clean water „ After 30 days in clean water
these larvae, if they survived, had residues no greater than those in control
larvae .
Total DDT residues were higher at all life stages whenever parent fish
were exposed to DDT in both the water and diet than for water exposure
alone. Even though DDT concentration may be greater from the water, presence
of DDT also in the diet caused higher tissue residues and death rates than a
corresponding water exposure alone. Death rates were not significant for
larvae exposed to dietary DDT at 30 and 60 days, perhaps because some larvae-
could not adapt to clam tissue when active feeding commenced, and deaths occurred
among the control fish by starvation. This was reflected in a mean mortality
of 3^% among control larvae at 30 days.
Results in the elimination phase of our study were similar to those
observed by other authors. Grzenda _et_ aiu (1970 ) demonstrated a 50%
elimination of DDT in goldfish by 29 days after exposure to dietary DDT
for 192 days. Gakstatter and Weiss (1967) observed less than 50% DDT
elimination after 32 days of recovery for bluegills that had been exposed
to 0.03 mg/1 DDT in the water for 5-19 hr. Buhler et_ al „ (1969) observed
an elimination of k^-68% of absorbed DDT in 35 days for chinook salmon
and 19-35% elimination for coho salmon within a similar time period.
Macek _et_ aJL. (1970 ) predicted, from their results, a 50% elimination of
total body DDT and dieldrin in rainbow trout within 160 and ^0 days, respectively,
after dietary exposure for ikO days. This elimination of DDT appears rather
long, but it is most likely influenced by species variability. In our
study almost all the eliminated DDT came from the DDT attributed to a dietary
47
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source, an indication of preferential elimination. We do not believe that this
selectivity was caused by metabolite differences, because the DDT, DDE, and TDE
residues in fish that had been exposed to DDT only in the vater essentially
did not change. As simple kinetics will not explain these observations,
unknown physiological factors are important, perhaps related to the
different routes of entry.
We have demonstrated that even though DDT uptake may be faster from a water
source, the presence of DDT in the food can cause higher tissue residues and a
significant increase in mortality. The residues from dietary exposure were not
as large as those observed for water exposure, but nevertheless the food chain
must be considered an important component. A "just safe" water concentration
or maximum acceptable toxicant concentration (MATC) determined from mortality
results would be about 0.9 pg/1 DDT for water exposure alone and about O.k yg/1
DDT with the added presence of dietary DDT. This increase in toxicity is greater
than 50%. Application factors as defined by Mount and Stephan (1967), using a
96-hr TL50 value of kQ yg/1 DDT demonstrated in one of our acute fathead minnow
toxicity tests and the maximum acceptable toxicant concentrations above,
would be 0.0188 or 1/53 for fish exposed to DDT in the water alone and 0.0083
or 1/120 for fish exposed to DDT both in the water and diet. Consequently,
food as well as water sources of" exposure to certain materials must be considered
when toxicity tests are designed or the conclusions drawn from such tests are
evaluated.
48
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Frisa. 1972. Effect of rate and duration of feeding DDT on the reproduction
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Chadwick, G0 G. , and R. Wo Brocksen. 1969. Accumulation of dieldrin by
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immature roman snails Helix ppmatia L. when treated with pp'-DDT.
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on fish and wildlife. U.S. Fish Wildl. Serv. Circ. 226:51-64.
Courtney, C. H. , and J. K. Reed. 1972. Accumulation of DDT from food
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25th Ann. Conf. Southeastern Assoc. Game and Fish Comm. p.
Dale, W. Eo, To B. Gaines, and W» J. Hayes, Jr. 1962. Storage and
excretion of DDT in starved rats. Toxicol „ Applu Pharmacol. 4:89-106.
Desaiah, D. , L. K. Cutkomp, R. B. Koch, and A. Jarvinen. 1975. DDT:
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Eberhardt, L. L. , R. L. Meeks, and To J. Peterle. 197l» Food chain model
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EcoMchon, D. J., and P. W. Saschenbrecker. 1969. The redistribution of
stored DDT in cockerels under the influence of food deprivation. Toxicol.
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Epifanio, C. E. 1973. Dieldrin uptake by larvae of the crab, Leptodius
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Ferguson, D. E., J. L. Ludke, M. T. Finley, and G. G. Murphy. 196?.
Insecticide-resistant fishes: a potential hazard to consumers. J. Miss.
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French, M. C., and D. J. Jefferies. 1971. The preservation of biological
tissue for organochlorine insecticide analysis. Bull. Environ. Contam.
Toxicol. 6:h60-h63.
Gakstatter, J, H., and C. M. Weiss. 1967. The elimination of DDT-C14,
dieldrin-C llf, and lindane-Clt+ from fish following a single sublethal exposure
in aquaria. Trans. Amer. Fish. Soc. 96:301-307.
Grant, B. F., and R. A. Schoettger. 1972. The impact of organochlorine
contaminants on physiological functions in fish. Proc. Tech. Sess. Ann.
Mtg. Inst. Environ. Sci. 18:2^5-250.
Grzenda, A. R., D. F. Paris, and W. J. Taylor. 1970. The uptake, metabolism,
and elimination of chlorinated residues by goldfish (Carassius auratus) fed
a x \J-DDT contaminated diet. Trans. Amer. Fish. Soc. 99:385-395.
Hamelink, J0 L., R. C. Waybrant, and R. C. Ball. 1971. A proposal:
Exchange equilibria control the degree chlorinated hydrocarbons are
biologically magnified in lentic environments. Trans. Amer. Fish. Soc. 100:
207-21U.
Harrison, H. L., 0. L. Loacks, J. W. Mitchill, D. F. Parkhurst, C. R. Tracy,
D. G. Watts, and V. J. Yannacone Jr. 1970. Systems studies of DDT
transport. Science 170:503-508.
Hermanutz, R. 0., L. H. Mueller, and K. D. Kempfert. 1973. Captan toxicity
to fathead minnows (Pimephales •promelas), bluegills (Lepomis macrochirus),
and brook trout (Salvelinus fontinalis). J. Fish. Res. Board Can. 30:l8ll-l8l7.
Holden, A. V. 1962. A study of the absorption of Cll+-labelled DDT from
water by fish. Ann. Appl. Biol. 50:467-777.
Huisman, E. A., J. H. Koeman, and P. V. I. M. Wolff. 1971- An investigation
into the influence of DDT and other chlorinated hydrocarbons on the fertility
of the pike. Ann. Rpt. Organ. Improv. Freshwater Fish. p. 69-86.
Hunt, E. Go, and A. I. Bischoff. 1960. Inimical effects on wildlife of
periodic DDD applications to Clear Lake. Calif. Fish Game U6:91-106.
Johnson, H. E0, and C. Pecor. 1969. Coho salmon mortality and DDT in Lake
Michigan. Trans. N. Amer. Wildl. Conf. 3^:159-166.
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Johnson, T. B,, Ro Co Saunders, H. 0. Sanders, and R. S. Campbell. 1971.
Biological magnification and degradation of DDT and aldrin by freshwater
invertebrates. J. Fish. Res. Board Can. 28:705-709.
Kleinert, S. J., and P. E. Degurse. 1973. Survival of walleye eggs and fry
of known DDT residue levels from ten Wisconsin waters in 19&7- Wisconsin Dept.
Nat. Resources Res. Rep. 37. 30 p.
Macek, K. J., and S. Korn. 1970. Significance of the food chain in DDT
accumulation by fish. J0 Fish. Res. Board Can. 27:1^96-1^98<>
Macek, K. J», C. R, Rodgers, D. L. Stalling, and Sc Korn. 1970. The uptake,
distribution, and elimination of dietary 14C-DDT and 11+C-dieldrin in
rainbow trout. Trans. Amer. Fish. Soc. 99:689-695-
Martin, J. W» 1967. A method of measuring lengths of juvenile salmon from
photographs. Prog. Fish-Cult. 29:238-2^0.
Mayer, F0 L., Jr. (personal communication). Fish-Pesticide Research
Laboratory, Columbia, Missouri.
McKim, J. Mo, and D0 A0 Benoit. 1971. Effects of long-term exposures to
copper on survival, growth, and reproduction of brook trout (Salvelinus
fontinalis). J. Fish. Res. Board Can. 28:655-662.
Metcalf, R. L. 1955. Organic insecticides. Interscience, New York. 392 p.
Mount, D. !„, and W. A0 Brungs. 1967. A simplified dosing apparatus for
fish toxicology studies. Water Res. 1:21-29.
Mount, D. Io, and C. E. Stephan. 1967. A method for estimating acceptable
toxicant limits for fish-malathion and the butoxyethanol ester of 2,k-~D.
Trans. Amer0 Fish. Soc. 96:185-193.
Moyle, Jo B., and J. L. Skrypek. 1969. Levels of DDT, DDE, and aldrin in
muscle and brain tissue of some Minnesota fishes, 1962-1967. Minnesota Dept.
Conserv., Div. Game and Fish, Spec. Publ, 59- 15 p.
Murphy, P» G. 1971. The effect of size on the uptake of DDT from water by
fish. Bull. Environ. Contain. Toxicol. 33:693-700.
O'Brien, R. D0 1967. Insecticide action and metabolism. Academic Press,
New York. 332 p.
Oettingen, W. F0, and N. E. Sharpless. 19^6. The toxicity and toxic
manifestations of 2,2-Bis (p-Chlorophenyl)-!,!,1-Trichloroethane (DDT) as
influenced by chemical changes in the molecule. J. Pharm. Exper Therap
88:itOO-lil3.
Priester, E. L., Jr. 1965. The accumulation and metabolism of DDT, parathion,
and endrin by aquatic food-chain organisms. Ph.D. Thesis. Clemson Univ
Clemson, S.C. jk p. *'
51
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Reinert, R. E. 1967. The accumulation of dieldrin in an alga (Scenedesmus
obliquus), Daphnia (Daphnia magna), guppy (Lebistes reticulatus) food chain.
Ph.D. Thesis. Univ. Michigan, Ann Arbor, Mich. 76 p.
Reinert, R. E. 1970. Pesticide concentrations in Great Lakes fish.
Pest. Mon. J. 3:233-240.
Reinert, R. E., and H. L. Bergman. 1974. Residues of DDT in lake trout
(Salvelinus namaycush) and coho salmon (Oncorhynchus kisutch) from the Great
Lakes.J. Fish. Res. Board Can. 31:191-199.
Rudd, R. L., and R. E. Genelly. 1956. Pesticides: Their use and toxicity in
relation to wildlife. Calif. Dept. Fish and Game, Bull. 7. 209 p.
Smith, W. E., K. Funk, and M. E. Zabik. 1973. Effects of cooking on
concentrations of PCB and DDT compounds in chinook (Oncorhynchus tshawytscha)
and coho (0. kisutch) salmon from Lake Michigan. J. Fish. Res. Board Can.
30:702-706.
Sprague, J. B. 1969. Measurement of pollutant toxicity to fish. I.
Bioassay methods for acute toxicity. Water Res. 3:793-821.
Steel, R. G. D., and J. H. Torrie. 1960. Principles and procedures of
statistics. McGraw-Hill Book New York. 481 p.
Syrett, R. E,, and W. F. Dawson. 1972. An inexpensieve electronic relay for
precise water-temperature control. Prog. Fish-Cult. 34:241-242.
U.S. Department of Health, Education, and Welfare. 1971. Pesticide Analytical
Manual. Vol. I. Section 211. 13 f, 211.14d, and 212.13b. Food and Drug
Administration, Washington, D.C.
U.S. Environmental Protection Agency. 1972. Pesticide standards program in
the EPA. Pesticide Standards Data sheets EPA/FDA No. 28 and FDA No. 206.
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Van Velzen, A. C., W. B. Stiles, and L. F. Stickel. 1972. Lethal mobilization
of DDT by cowbirds. J. Wildl. Manage. 36:733-739.
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APPENDIX
RECOMMENDED BIOASSAY PROCEDURE FOR
FATHEAD MINNOW PIMEPHALES PROMELAS RAFINESQUE CHRONIC TESTS
April, 1971
(Revised January, 1972)
by
Environmental Research Laboratory-Duluth
(formerly the National Water Quality Laboratory)
Duluth, Minnesota 55804
ENVIRONMENTAL RESEARCH LABORATORY-DULUTH
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
DULUTH, MINNESOTA 55804
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RECOMMENDED BIOASSAY PROCEDURES
Preface
Recommended Bioassay Procedures are established by the approval of both
the Committee on Aquatic Bioassays and the Director of the National
Water Quality Laboratory. The main reasons for establishing them are:
(1) to permit direct comparison of test results, (2) to encourage
the use of the best procedures available, and (3) to encourage
uniformity. These procedures should be used by National Water Quality
Laboratory personnel whenever possible; unless there is a good reason
for using some other procedure.
Recommended Bioassay Procedures consider the basic elements that are
believed to be important in obtaining reliable and reproducible
results in laboratory bioassays. An attempt has been made to adopt
the best acceptable procedures based on current evidence and opinion,
although it is recognized that alternative procedures may be adequate.
Improvements in the procedures are being considered and tested, and
revisions will be made when necessary. Comments and suggestions are
encouraged.
Director, National Water Quality Lab, (NWQL)
Committee on Aquatic Bioassays, NWQL
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Recommended Bioassay Procedure for
Fathead Minnow Pimephales promelas Rafinesque Chronic Tests
April, 1971
(Revised January, 1972)
A. Physical system
1. Diluter: Proportional diluters (Mount and Brungs, 1967) should
be employed for all long-term exposures. Check the operation
of the diluter daily, either directly or through
measurement of toxicant concentrations. A minimum of five
toxicant concentrations and one control should be used for
each test with a dilution factor of not less than 0.30. An
automatically triggered emergency aeration and alarm system
must be installed to alert staff in case of diluter, temperature
control or water supply failure.
2. Toxicant mixing: A container to promote mixing of toxicant
bearing and w-cell water should be used between diluter and
tanks for each concentration. Separate delivery tubes
should run from this container to each duplicate tank.
Check at least once every month to see that the intended
amounts of water are going to each duplicate tank or chamber.
3. Tank: Two arrangements of test tanks (glass, or stainless
steel with glass ends) can be utilized:
a. Duplicate spawning tanks measuring 1 x 1 x 3 ft. long
with a one sq. ft. portion at one end screened off
and divided in half for the progeny. Test water is
to be delivered separately to the larval and spawning
chambers of each tank, with about one-third the water
volume going to the former chamber as to the latter.
b. Duplicate spawning tanks measuring 1x1x2 ft. long
with a separate duplicate progeny tank for each
spawning tank. The larval tank for each spawning
tank should be a minimum of 1 cu. ft. dimensionally
and divided to form two separate larval chambers with
separate standpipes, or separate 1/2 sq. ft. tanks
may be used. Test water is to be supplied by delivery
tubes from the mixing cells described in Step 2 above.
Test water depth in tanks and chambers for both a & b
above should be 6 inches.
4. Flow rate: The flow rate to each chamber (larval or adult)
should be equal to 6 to 10 tank volumes/24 hr.
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5* Aeration; Total dissolved oxygen levels should never be allowed
to drop below 60% of saturation, and flow rates must be increased
if oxygen levels do drop below 60%. As a first alternative flow
rates can be increased above those specified in A.4. Only
aerate (with oil free air) if testing a non-volatile toxic agent,
and then as a last resort to maintain dissolved oxygen at 60%
of saturation.
6. Cleaning: All adult tanks, and larvae tanks and chambers after
larvae swim-up, must be siphoned a minimum of 2 times weekly
and brushed or scraped when algal or fungus growth becomes
excessive.
7. Spawning subs tra te: Use spawning substrates made from inverted
cement and asbestos halved, 3-inch ID drain tile, or the equiva-
lent, each of these being 3 inches long.
^* Egg cup: Egg incubation cups are made from either 3-inch
sections of 2-inch OD (l 1/2-inch ID) polyethylene water hose
or 4-oz., 2-inch OD round glass jars, with the bottoms cut off.
One end of the jar or hose sections is covered with stainless
steel or nylon screen (with a minimum of 40 meshes per inch).
Cups are oscillated in the test water by means of a rocker arm
apparatus driven by a 2 r.p.m. electric motor (Mount, 1968).
The vertical-travel distance of the oips should be 1 to 1 1/2
inches.
9. Light; The lights used should simulate sunlight as nearly as
possible. A combination of Durotest (Optima FS)1»2 and wide
spectrum Grow-lux-^ fluorescent tubes has proved satisfactory at
the NWQL.
10. Pnotoperiod; The photoperiods to be used (Appendix A) simulate
the dawn to dusk times of Evansville, Indiana. Adjustments in
day-length are to be made on the first and fifteenth day of
every Evansville test month. The table is arranged so that
adjustments need be made only in the dusk times. Regardless
of the actual date that the experiment is started, the Evansville
test photoperiod should be adjusted so that the mean or estimated
hatching date of the fish used to start the experiment corresponds
to the Evansville test day-length for December first. Also,
the dawn and dusk times listed in the table need not correspond
to the actual times where the experiment is being conducted. To
illustrate these points, an experiment started with 5-day-old
larvae in Duluth, Minnesota, on August 28 (actual date), would
require use of a December 5 Evansville test photoperiod, and
the lights could go on anytime on that day just so long as they
remained on for 10 hours and 45 minutes. Ten days later (Sept. 7
actual date, Dec. 15 Evansville test date) the day-length
Mention of trade names does not constitute endorsement.
2
Duro-Test, Inc., Hammond, Ind.
3
Sylvania, Inc., New York, N. Y.
56
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would be changed to 10 hours and 30 minutes. Gradual changes
in light intensity at dawn and dusk (Drummond and Dawson, 1970),
if desired, should be included within the day-lengths shown,
and should not last for more than 1/2 hour from full on to full
off and vice versa.
11. Temperature: Temperature should not deviate instantaneously
from 25° C by more than 2° C and should not remain outside the
range of 24 to 26° C for more than 48 hours at a time. Temperature
should be recorded continuously.
12. Disturbance; Adults and larvae should be shielded from
disturbances such as people continually walking past the
chambers, or from extraneous lights that might alter the
intended photoperiod.
13. Construction materials; Construction materials which contact
the diluent water should not contain leachable substances and
should not sorb significant amounts of substances from the water.
Stainless steel is probably the preferred construction material.
Glass absorbs some trace organics significantly. Rubber should
not be used. Plastic containing fillers, additives, stabilizers,
plasticizers, etc., should not be used. Teflon, nylon, and
their equivalents should not contain leachable materials and
should not sorb significant amounts of most substances. Un-
plasticized polyethylene and polypropylene should not contain
leachable substances, but may sorb very significant amounts of
trace organic compounds.
14. Water; The water used should be from a well or spring if at
all possible, or alternatively from a surface water source.
Only as a last resort should water from a chlorinated municipal
water supply be used. If it is thought that the water supply
could be conceivably contaminated with fish pathogens, the
water should be passed through an ultraviolet or similar ster-
ilizer immediately before it enters the test system.
B. Biological system
!• Test animals; If possible, use stocks of fathead minnows from
the National Water Quality Laboratory in Duluth, Minnesota or
the Fish Toxicology Laboratory in Newtown, Ohio. Groups of
starting fish should contain a. mixture of approximately equal
numbers of eggs or larvae from at least three different females.
Set aside enough eggs or larvae at the start of the test to
supply an adequate number of fish for the acute mortality
bioassays used in determining application factors.
57
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2. Beginning test; In beginning the test, distribute 40 to
50 eggs or 1 to 5-day-old larvae per duplicate tank using a
stratified random assignment (see D.3). All acute mortality
tests should be conducted when the fish are 2 to 3 months old.
If eggs or 1 to 5-day-old larvae are not available, fish up to
30 days of age may be used to start the test. If fish
between 20 and 60 days old are used, the exposure should
be designated a partial chronic test. Extra test animals
may be added at the beginning so that fish can be removed
periodically for special examinations (see B.12.) or for
residue analysis (see C.4.).
3. Food: Feed the fish a frozen trout food (e.g., Oregon
Moist). A minimum of once daily fish should be fed ad
libitum the largest pellet they will take. Diets should
be supplemented weekly with live or frozen-live food
(e.g., Daphnia, chopped earthworms, fresh or frozen brine
shrimp, etc.). Larvae should be fed a fine trout starter
a minimum of 2 times daily, ad libitum; one feeding each
day of live young zooplankton from mixed cultures of
small copepods, rotifers, and protozoans is highly
recommended. Live food is especially important when
larvae are just beginning to feed, or about 8 to 10 days
after egg deposition. Each batch of food should be
checked for pesticides (including DDT, IDE, dieldrin,
lindane, methoxychlor, endrin, aldrin, BHC, chlordane,
toxaphene, 2,4-D, and PCBs), and the kinds and amounts
should be reported to the project officer or recorded.
4. Disease: Handle disease outbreaks according to their
nature, with all tanks receiving the same treatment
whether there seems to be sick fish in all of them or
not. The frequency of treatment should be held to a
minimum.
5. Measuring fish; Measure total lengths of all starting fish
at 30 and 60 days by the photographic method used by McKim
and Benoit (1971). Larvae or juveniles are transferred
to a glass box containing 1 inch of test water. Fish
should be moved to and from this box in a water-filled
container, rather than by netting them. The glass box
is placed on a translucent millimeter grid over a
fluorescent light platform to provide background
illumination. Photos are then taken of the fish over
58
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the millimeter grid and are enlarged into 8 by 10 inch
prints. The length of each fish is subsequently
determined by comparing it to the grid. .Keep lengths of
discarded fish separate from those of fish that are to be
kept.
6. Thinning; When the starting fish are sixty (+ 1 or 2) days
old, impartially reduce the number of surviving fish in
each tank to 15. Obviously injured or crippled individuals
may be discarded before the selection so long as the number
is not reduced below 15; be sure to record the number of
deformed fish discarded from each tank. As a last resort in
obtaining 15 fish per tank, 1 or 2 fish may be selected for
transfer from one duplicate to the other. Place five spawning
tiles in each duplicate tank, separated fairly widely to reduce
interactions between male fish guarding them. One should
also be able to look under tiles from the end of the tanks.
During the spawning period, sexually maturing males must be
removed at weekly intervals so there are no more than four
per tank. An effort should be made not to remove those
males having well established territories under tiles where
recent spawnings have occurred.
7. Removing eggs: Remove eggs from spawning tiles starting at
12:00 noon Evansville test time (Appendix A) each day.
As indicated in Step A.9., the test time need not correspond
to the actual time where the test is being conducted. Eggs
are loosened from the spawning tiles and at the same time
separated from one another by lightly placing a finger on
the egg mass and moving it in a circular pattern with
increasing pressure until the eggs being to roll. The
groups of eggs should then be washed into separate,
appropriately marked containers and subsequently handled
(counted, selected for incubation, or discarded) as soon as
possible after all eggs have been removed and the spawning
tiles put back into the test tanks. All egg batches must
be checked initially for different stages of development.
If it is determined that there is more than one distinct
stage of development present, then each stage must be
considered as one spawning and handled Separately as
described in Step B.8.
8. Egg incubation and larval selection; Impartially select
50 unbroken eggs from spawnings of 50 eggs or more and
place them in an egg incubator cup for determining
viability and hatchability. Count the remaining eggs and
discard them. Viability and hatchability determinations
must be made on each spawning (>49 eggs) until the number
of spawnings (>49 eggs) in each duplicate tank equals the
59
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number of females in that tank. Subsequently, only eggs
from every third spawning (>49 eggs) and none of those
obtained on weekends need be set up to determine hatch-
ability; however, weekend spawns must still be removed from
tiles and the eggs counted. If unforseen problems are
encountered in determining egg viability and hatchability,
additional spawnings should be sampled before switching to
the setting up of eggs from every third spawning. Every
day record the live and dead eggs in the incubator cups,
remove the dead ones, and clean the cup screens. Total
numbers of eggs accounted for should always add up to
within two of 50 or the entire batch is to be discarded.
When larvae begin-to hatch, generally after 4 to 6 days,
they should not be handled again or removed from the egg-
cups until all have hatched. Then, if enough are still
alive, 40 of these are eligible to be transferred
immediately to a larval test chamber. Those individuals
selected out to bring the number kept to 40 should be
chosen impartially. Entire egg-cup-groups not used for
survival and growth studies should be counted and
discarded.
9. Progeny transfer: Additional important information on
hatchability and larval survival is to be gained by
transferring control eggs immediately after spawning to
concentrations where spawning is reduced or absent, or
to where an affect is seen on survival of eggs or larvae,
and by transferring eggs from these concentrations to
the control tanks. One larval chamber in, or corresponding
to, each adult tank should always be reserved for eggs
produced in that tank.
10. Larval exposure: From early spawnings in each duplicate
tank, use the larvae hatched in the egg incubator cups
(Step B.8. above) for 30 or 60 day growth and survival
exposures in the larval chambers. Plan ahead in setting
up eggs for hatchability so that a new group of larvae is
ready to be tested for 30 or 60 days as soon as possible
after the previously tested group comes out of the larval
chambers. Record mortalities, and measure total lengths
of larvae at 30 and, if they are kept, 60 days post-
hatch. At the time the larval test is terminated they
should also be weighed. No fish (larvae, juveniles, or
adults) should be fed within 24 hr's. of when they are to
be weighed.
60
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11. Parental termination: Parental fish testing should be
terminated when, during the receding day-length photo-
period, a one week period passes in which no spawning
occurs in any of the tanks. Measure total lengths and
weights of parental fish; cheek sex and condition of
gonads. The gonads of most parental fish will have
begun to regress from the spawning condition, and thus
the differences between the sexes will be less distinct
now than previously. Males and females that are readily
distinguishable from one another because of their
external characteristics should be selected initially for
determining how to differentiate between testes and
ovaries. One of the more obvious external characteristics
of females that have spawned is an extended, transparent
anal canal (urogenital papilla). The gonads of both
sexes will be located just ventral to the kidneys. The
ovaries of the females at this time will appear transparent,
but perhaps containing some yellow pigment, coarsely
granular, and larger than testes. The testes of males
will appear as slender, slightly milkly, and very finely
granular strands. Fish must not be frozen before making
these examinations.
12. Special examinations: Fish and eggs obtained from the test
should be considered for physiological, biochemical, histo-
logical and other examinations which may indicate certain
toxicant related effects.
13. Necessary data: Data that must be reported for each tank
of a chronic test are:
a. Number and individual total length of normal and deformed
fish at 30 and 60 days; total length, weight and number
of either sex, both normal and deformed, at end of test.
b. Mortality during the test.
c. Number of spawns and eggs.
d. Hatchability.
e. Fry survival, growth, and deformities.
61
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C. Chemical system
1. Preparing a. stock solution; If a toxicant cannot be introduced
into the test water as is, a stock solution should be prepared
by dissolving the toxicant in water or an organic solvent.
Acetone has been the most widely used solvent, but dimethylformanide
(DMF) and triethylene glycol may be preferred in many cases.
If none of these solvents are acceptable, other water-miscible
solvents such as methanol, ethanol, isopropanol, acetonitrile,
dimethylacetamide (DMAC), 2-ethoxyethanol, glyme (dimethylether
of ethylene glycol, diglyme (dimethyl ether of diethylene glycol)
and propylene glycol should be considered. However, dimethyl
sulfoxide (DMSO) should not be used if at all possible because
of its biological properties.
Problems of rate of solubilization or solubility limit should be
solved by mechanical means if at all possible. Solvents, or as
a last resort, surfactants, can be used for this purpose, only
after they have been proven to be necessary in the actual test
system. The suggested surfactant is p-tert-octylphenoxynonaethoxy-
ethanol (p-1, 1, 3, 3-tetramethylbutylphenoxynonaethoxyethanol,
OPE,Q) (Triton X-100, a product of the Rohm and Haas Company, or
equivalent).
The use of solvents, surfactants, or other additives should be
avoided whenever possible. If an additive is necessary, reagent
grade or better should be used. The amount of an additive used
should be kept to a minimum, but the calculated concentration of
a solvent to which any test organisms are exposed must never exceed
one one-thousandth of the 96-hr. TL50 for test species under the
test conditions and must never exceed one gram per liter of water.
The calculated concentration of surfactant or other additive to
which any test organisms are exposed must never exceed one-twentieth
of the concentration of the toxicant and must never exceed one-tenth
gram per liter of water. If any additive is used, two sets of
controls must be used, one exposed to no additives and one exposed
to the highest level of additives to which any other organisms
in the test are exposed.
2. Measurement of toxicant concentration: As a minimum the
concentration of toxicant must be measured in one tank at each
toxicant concentration every week for each set of duplicate
tanks, alternating tanks at each concentration from week to
week. Water samples should be taken about midway between the
top and bottom and the sides of the tank and should not include
any surface scum or material stirred up from the bottom or sides
of the tank. Equivolume daily grab samples can be composited
for a week if it has been shown that the results of the analysis
are not affected by storage of the sample.
62
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Enough grouped grab samples should be analyzed periodically
throughout the test to determine whether or not the concentration
of toxicant is reasonably constant from day to day in one tank
and from one tank to its duplicate. If not., enough samples must
be analyzed weekly throughout the test to show the variability
of the toxicant concentration.
3. Measurement p_f other variables: Temperature must be recorded
continuously (see A.10=).
Dissolved oxygen must be measured in the tanks daily, at least
five days a week on an alternating basis, so that each tank is
analyzed once each week. However, if the toxicant or an additive
causes a depression in dissolved oxygen, the toxicant concentration
with the lowest dissolved oxygen concentration must be analyzed
daily in addition to the above requirement.
A control and one test concentration must be analyzed weekly for
pH, alkalinity, hardness, acidity, and conductance or more often,
if necessary, to show the variability in the test water. However,
if any of these characteristics are affected by the toxicant
the tanks must be analyzed for that characteristic daily, at
least five days a week, on an alternating basis so that each
tank is analyzed once every other week.
At a minimum, the test water must be analyzed at the beginning
and near the middle of the test for calcium, magnesium, sodium,
potassium, chloride, sulfate, total solids, and total dissolved
solids.
4. Residue analysis; When possible and deemed necessary, mature
fish, and possibly eggs, larvae, and juveniles, obtained from
the test, should be analyzed for toxicant residues. For fish,
muscle should be analyzed, and gill, blood, brain, liver, bone,
kidney, GI tract, gonad, and skin should be considered for
analysis. Analyses of whole organisms may be done in addition
to, but should not be done in place of, analyses of individual
tissues, especially muscle.
5. Methods: When they will provide the desired information with
acceptable precision and accuracy, methods described in Methods
for Chemical Analysis of Water and Wastes (EPA, 1971) should be
used unless there is another method which requires much less time
and can provide the desired information with the same or better
precision and accuracy. At a minimum, accuracy should be measured
using the method of known additions for all analytical methods
for toxicants. If available, reference samples should be
analyzed periodically for each analytical method.
63
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D. Statistics
1. Duplicates: Use true duplicates for each level of toxic agent,
i.e., no water connections between duplicate tanks.
2. Distribution of tanks: The tanks should be assigned to locations
by stratified random assignment (random assignment of one tank
for each level of toxic agent in a row followed by random assign-
ment of the second tank for each level of toxic agent in another
or an extension of the same row).
3. Distribution of_ test organisms : The test organisms should be
assigned to tanks by stratified random assignment (random assignment
of one test organism to each tank, random assignment of a second
test organism to each tank, etc.).
E. Miscellaneous
1. Additional information: All routine bioassay flow through methods
not covered in this procedure (e.g., physical and chemical
determinations, handling of fish) should be followed as
described in Standard Methods for tie Examination of Water and
Wastewater, (American Public Health Association, 1971), or
information requested from appropriate persons at Duluth or
Newtown.
2. Acknowledgments; These procedures for the fathead minnow
were compiled by John Eaton for the Committee on Aquatic
Bioassays. The participating members of this committee are:
Robert Andrew, John Arthur, Duane Benoit, Gerald Bouck,
William Brungs, Gary Chapman, John Eaton, John Hale,
Kenneth Hokanson, James McKim, Quentin Pickering, Wesley
Smith, Charles Stephan, and James Tucker.
3. References: For additional information concerning flow
through bioassays with fathead minnows, the following
references are listed:
American Public Health Association. 1971. Standard
methods for the examination of water and wastewater.
13th ed. APHA. New York.
Brungs, William A. 1969. Chronic toxicity of zinc to the
fathead minnow, Pimephales promelas Rafinesque. Trans. Amer.
Fish. Soc., 98(2): 272-279.
Brungs, William A. 1971. Chronic effects of low dissolved
oxygen concentrations on the fathead minnow (Pimephales promelas).
J. Fish. Res. Bd. Canada, 28(8): 1119-1123.
64
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Brungs, William A. 1971. Chronic effects of constant elevated
temperature on the fathead minnow (Pimephales promelas). Trans,
Amer. Fish. Soc. 100(4): 659-664.
Carlson, Dale R. 1967. Fathead minnow, Pimephales promelas
Rafinesque, in the Des Moines River, Boone County, Iowa, and
the Skunk River drainage, Hamilton and Story Counties, Iowa.
Iowa State Journal of Science, 41(3): 363-374.
Drummond, Robert A., and Walter F. Dawson. 1970. An
inexpensive method for simulating Diel patterns of lighting
in the laboratory. Trans. Amer. Fish. Soc., 99(2): 434-435.
Isaak, Daniel. 1961. The ecological life history of the
fathead minnow, Pimephales promelas (Rafinesque). Ph.D.
Thesis, Library, Univ. of Minnesota.
Markus, Henry C. 1934. Life history of the fathead minnow
(Pimephales promelas). Copeia, (3): 116-122.
\
McKim, J. M., and D. A. Benoit. 1971. Effect of long-term
exposures to copper on survival, reproduction, and growth
of brook trout Salvelinus fontinalis (Mitchill). J. Fish.
Res. Bd. Canada, 28: 655-662.
Mount, Donald I. 1968. Chronic toxicity of copper to
fathead minnows (Pimephales promelas, Rafinesque). Water
Research, 2: 215-223.
Mount, Donald I., and William Brungs. 1967. A simplified
dosing apparatus for fish toxicology studies. Water Research,
1: 21-29.
Mount, Donald I., and Charles E. Stephan. 1967. A method
for establishing acceptable toxicant limits for fish —
malathion and the butoxyethanol ester of 2,4-D. Trans.
Amer. Fish. Soc., 96(2): 185-193.
Mount, Donald I., and Charles E. Stephan. 1969. Chronic
toxicity of copper to the fathead minnow (Pimephales promelas)
in soft water. J. Fish. Res. Bd. Canada, 26(9): 2449-2457.
Mount, Donald I., and Richard E. Warner. 1965. A serial-
dilution apparatus for continuous delivery of various
concentrations of materials in water. PHS Publ. No- 999-
WP-23. 16 pp.
Pickering, Quentin H., and Thomas 0. Thatcher. 1970. The
chronic toxicity of linear alkylate sulfonate (LAS) to
Pimephales promelas, Rafinesque. Jour. Water Poll. Cont
Fed., 42(2): 243-254.
65
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Pickering, Quentin H., and William N. Vigor. 1965. The
acute toxicity of zinc to eggs and fry of the fathead
minnow. Progressive Fish-Culturist, 27(3); 153-157.
Verma, Prabha. 1969- Normal stages in the development
of Cyprinus carpio var. communis L. Acta biol. Acad. Sci.
Hung., 21(2): 207-218.
Approved by the Committee
on Aquatic Bioassays, NWQL
Approved by the Director, NWQL
66
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Appendix A
Test (Evansville, Indiana) Photoperiod
For Fathead Minnow Chronic
Dawn to Dusk
Time
Date
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
6:00 -
4:45)
4:30)
4:30)
4:45)
5:15)
5:45)
6:15)
7:00)
7:30)
8:15)
8:45)
9:15)
9:30)
9:45)
9:45)
9:30)
9:00)
8:30)
8:00)
7:30)
6:45)
6:15)
5:30)
5:00)
DEC.
JAN.
FEB.
MAR.
APR.
MAY
JUNE
JULY
AUG.
SEPT.
OCT.
NOV.
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
1
15
10:45)
10:30)
10:30)
10:45)
11:15)
11:45)
12:15)
13:00)
13:30)
14:15)
14:45)
15:15)
15:30)
15:45)
15:45)
15:30)
15:00)
14:30)
14:00)
13:30)
12:45)
12:15)
11:30)
11:00)
Day-length (hour and minute)
5-month pre-
spawning growth
period
4-month spawning
period
post spawning period
67
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TECHNICAL REPORT DATA
(Please read Instructions on the reverse before completing)
1. REPORT NO.
EPA-600/3-76-114
4. TITLE AND SUBTITLE
TOXICITY OF DDT FOOD AND WATER EXPOSURE TO
FATHEAD MINNOWS
3. RECIPIENT'S ACCESSION NO.
5. REPORT DATE
December 1976 (Issuing Date)
6. PERFORMING ORGANIZATION CODE
7. AUTHOR(S)
Alfred W. Jarvinen, Molly J. Hoffman, and Todd W.
Thorslund
8. PERFORMING ORGANIZATION REPORT NO.
9. PERFORMING ORGANIZATION NAME AND ADDRESS
Environmental Research Laboratory-Duluth
6201 Congdon Boulevard
Duluth, Minnesota 55804
10. PROGRAM ELEMENT NO.
1BA608; ROAP/Task 16AAK/010
11. CONTRACT/GRANT NO.
12. SPONSORING AGENCY NAME AND ADDRESS
Environmental Research Laboratory - Duluth, Minn.
Office of Research and Development
U.S. Environmental Protection Agency
Duluth, Minnesota 55804
13. TYPE OF REPORT AND PERIOD COVERED
Final
14. SPONSORING AGENCY CODE
EPA/600/03
15. SUPPLEMENTARY NOTES
16. ABSTRACT
Fathead minnows (Pimephales promelas) were exposed during a partial chronic
toxicity test to two DDT concentrations in the water, one in the diet, and combination
of water and diet for 266 days through a reproductive period of their life cycle.
Tissue-residue analyses were performed on test fish at preset intervals throughout the
exposure and also on embryos, larvae at hatch, and 30- and 60-day progeny. The
contribution of DDT from each source was monitored with gas-chromatography and liquid-
scintillation techniques. The diet was clams that had accumulated 14C-DDT when
exposed at a DDT water concentration similar to that in the high fish exposure.
Higher total DDT tissue residues were accumulated from the water than from the
diet. Residues contributed by dietary DDT were additive to those from the water,
Mean concentration factors were 1.2 times from the diet and 100,000 times from the
water. Mortality was higher in fish exposed to DDT in both water and diet than in
fish exposed to only one or the other of these sources. DDT in the diet significantly
reduced the probability of survival of the test fish (P=0.025). Estimated maximum
acceptable toxicant concentrations for DDT are 0.9 yg/1 for fish exposed to DDT in the
water only or 0.4 yg/1 for fish exposed to DDT in both water and diet.
KEY WORDS AND DOCUMENT ANALYSIS
DESCRIPTORS
Bioassay*
Aquatic animals
Minnows*
Clams
Pesticides*
DDT*
Freshwater fishes
Insecticides
Food chain
Toxicity
Carbon 14
b.lDENTIFIERS/OPEN ENDED TERMS
Aquatic life
Chlorinated hydrocarbon
Bioaccumulation
Tissue Residues
COS AT I Field/Group
6A
6C
6F
7B
7C
7E
3. DISTRIBUTION STATEMENT
RELEASE TO PUBLIC
19. SECURITY CLASS (ThisReport)
UNCLASSIFIED
21. NO. OF PAGES
76
20. SECURITY CLASS (This page)
UNCLASSIFIED
22. PRICE
EPA Form 2220-1 (9-73)
68
ftU.S. GOVERNMENT PRINTING OFFICE: 1977-757-056/5524
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