United States
Environmental Protection
Agency
Environmental Research
Laboratory
Narragansett Rl 02882
EPA 600 3 80053
June 1980
Research and Development
The Effect of
Different Pollutants
on Ecologically
Important Polychaete
Worms
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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 nine series. These nine broad cate-
gories were established to facilitate further development and application of en-
vironmental technology. Elimination of traditional grouping was consciously
planned to foster technology transfer and a maximum interface in related fields.
The nine series are:
1. Environmental Health Effects Research
2. Environmental Protection Technology
3. Ecological Research
4. Environmental Monitoring
5. Socioeconomic Environmental Studies
6. Scientific and Technical Assessment Reports (STAR)
7. Interagency Energy-Environment Research and Development
8. "Special" Reports
9. Miscellaneous Reports
This report has been assigned to the ECOLOGICAL RESEARCH series. This series
describes research on the effects of pollution on humans, plant and animal spe-
cies, and materials. Problems are assessed for their long- and short-term influ-
ences. Investigations include formation, transport, and pathway studies to deter-
mine 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 600/3-80-053
September 1980
THE EFFECT OF DIFFERENT POLLUTANTS
ON ECOLOGICALLY IMPORTANT POLYCHAETE WORMS
by
Donald J. Reish
California State University
Long Beach, California 90840
Grant No. 800962
Project Officer
Donald K. Phelps
Environmental Research Laboratory
Narragansett, Rhode Island 02882
ENVIRONMENTAL RESEARCH LABORATORY
OFFICE OF RESEARCH AND DEVELOPMENT
U.S. ENVIRONMENTAL PROTECTION AGENCY
NARRAGANSETT, RHODE ISLAND 02882
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DISCLAIMER
This report has been reviewed by the Environmental Research Laboratory,
Narragansett, U.S. Environmental Protection Agency, and approved for publica-
tion. Approval does not signify that the contents necessarily reflect the
views and policies of the U.S. Environmental Protection Agency, nor does
mention of trade names or commercial products constitute endorsement or
recommendation for use.
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FOREWORD
The Environmental Research Laboratory of the U.S. Environmental
Protection Agency is located on the shore of Narragansett Bay, Rhode Island.
In order to assure the protection of marine resources, the laboratory is
charged with providing a scientifically sound basis for Agency decisions on
the environmental safety of various uses of marine systems. To a great
extent, this requires research on the tolerance of marine organisms and
their life stages as well as of ecosystems to many forms of pollution stress.
In addition, a knowledge of pollutant transport and fate is needed.
This report describes the results of a 3.5 year study to establish
laboratory colonies of polychaetous annelids and to measure the effects of
heavy metals and petroleum hydrocarbons on these organisms through a com-
plete life cycle. This report covers the period of time from late 1972
through June 1976. Certain phases of the investigations have been updated
since the termination of the grant to reflect additional data which has
been accumulated since June, 1976.
iii
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ABSTRACT
The procedures for culturing marine polychaetous annelids from egg to
egg under laboratory conditions were described. A manual was prepared de-
tailing the procedures used in culturing 12 species of polychaetes. The
polychaetes which have been successfully cultured and the number of cycles
completed in the laboratory are: Neanthes arenaceodentata (50+), Capitella
capitata (50+), Ctenodrilus serratus (50+), Opkn/otrooha diadema (50+),
0. pueTpilis (20+), Dinophilus sp. (50+), Dexiospira brasiliensis (3),
Polydora ligni, (3), Boccard-iaprobosoidea (3), Cirviformia luxwri-osa (1),
C. spirdbraneha (1), and Halosydna johnsoni (1).
The effects of heavy metals and the water soluble fractions of petro-
leum hydrocarbons were measured over 96 hours, 28 days, and with some of the
toxicants, over a complete reproductive cycle for some of these species of
polychaetes. Mercury and copper were the most toxic of the six metals tested
and cadmium was the least toxic. The 28-day LCso was less than the 96-hour
value in most experiments. Larval stages were more sensitive than the adults
to heavy metals. Dexiospira was the most sensitive species and Cirri-formia
luxuriosa was the most tolerant. Suppression of reproduction occurred with
each species studied when exposed to heavy metals; the concentrations at
which this occurred was less than the 28-day LC5Q.
The water soluble fraction of a refined oil was more toxic than a crude
oil. Capi-tella was the most tolerant species to the oils and Opfan/otroeha
diadema was the most sensitive one tested. Suppression of reproduction was
observed for both Ctenodrilus and Oplwyotrocha di-adema.
An interlaboratory calibration experiment using Capitella was described
and the publication resulting from this study has been published elsewhere.
IV
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CONTENTS
Page
Disclaimer ii
Foreword iii
Abstract iv
List of Figures vi
List of Tables ix
Acknowledgments x
1. Introduction 1
2. Laboratory Culture Procedures with Polychaetous Annelids 3
3. The Effects of Heavy Metals on Polychaetous Annelids . . 7
4. The Effects of Petroleum Hydrocarbons on Polychaetous
Annelids 10
5. An Interlaboratory Calibration Experiment Utilizing
Cap'i'bella oap-itata 11
6. Appendices 18
A. Culture and Bioassay Procedures for Polychaetous
Annelids: Neanthes arenaceodentata 18
B. Culture and Bioassay Procedures for Polychaetous
Annelids: Cap'l'tella. capi.ta.ta 36
C. Culture and Bioassay Procedures for Polychaetous
Annelids: Ctenodri,1us serratus 54
D. Culture and Bioassay Procedures for Polychaetous
Annelids: Ophyrotrocha diadema and 0. puerilus ... 62
E. Culture and Bioassay Procedures for Polychaetous
Annelids: Janua (Dexi.ospi.ro.) brasi.li.ensi,s 75
F. Culture and Bioassay Procedures for Polychaetous
Annelids: Polydora ligni and Boccardi, proboscidea . 83
G. Culture and Bioassay Procedures for Polychaetous
Annelids: Di.nophi.lus sp 103
H. Culture and Bioassay Procedures for Polychaetous
Annelids: Cirriformia luxuviosa and C. spirobrancha 111
I. Culture and Bioassay Procedures for Polychaetous
Annelids: Ealosydna johnsoni 123
J. List of Publications and Thesis Supported by This
Research Grant 133
K. List of Laboratories Which Have Utilized Specimens
Cultured in California State University, Long Beach . 136
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FIGURES
No. _
— Page
1. Neanthes ax>enaceodentata, anterior end, with everted
proboscis 28
2. NeantheSj neuropodial heterogomph falcigerous seta with
hook at tip of the blade 29
3. Neanthes of the same sex in fighting position 30
4. Neanthes3 three segmented stage 31
5. Neanthesj four segmented stage 32
6. Neanthes j 12 segmented stage 33
7. NeantheSf juvenile, with 21 segments, which has left the
parent's tube and commenced feeding 34
8. One gallon (3.78 A) aquarium system fitted with an outside
filter system used to culture Neanthes adults, especially
those specimens approaching sexual maturity 35
9. Capitella eapitata, male, dorsal view of anterior end.
Note the genital hooks in setigerous segments 8 and 9 ... 46
10. Capi-teZlaj female, dorsal view of anterior end 47
11. Cap'itella, female, incubating eggs, lateral view. Note
the mucoid tube with the fertilized eggs lining the
inside of the tube 48
12. Capitella.; trochophore stage 49
13. Cap'ite1'la3 metatrochophore stage 50
14. Capitel'la, juvenile 51
15. Gallon (3.78 £) jar aquarium system for rearing Capi-tella.
The colony is located within the mass of Enteromorpha at
the bottom of the jar. Air is supplied by an aquarium
stone connected to an air compressor 52
VI
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No. Page
16. Capitella, bifurcated abnormal larva induced by sublethal
amounts of a heavy metal 53
17. Ctenodrilus sewatus, dorsal view of entire worm 59
18. Ctenodrilus, simple seta 60
19. Ctenodrilus with five buds forming by transverse
fission 61
20. OphryotTocha diadema, dorsal view of entire worm
(after Akesson, 1976) 70
21. Opkryotrocha; egg mass (after Akesson, 1976) 71
22. Opkpyotrochaf larva from egg mass (after Akesson, 1976). . . 72
23. Ophryotrochotj larva from egg mass (after Akesson, 1976). . . 73
o
24. OphryotrochcLj released larva (after Akesson, 1976) 74
25. Jania (Dexi,ospi,x>a)bz>as'ili,ensis3 tube 81
26. Dex-Losp-iraj dorsal view of entire worm removed from tube . . 82
27. PoZydora ligni, anterior end, dorsal view showing location of
palps, nuchal antenna and fifth setiger 92
28. Polydordj posterior end, dorsal view showing disc-like
pygidium ..... 93
29. Polydora, setae of modified fifth setiger showing one
spine with a subapical tooth and companion seta with
forked tip 94
30. Polydova, portion of a string of egg capsules as they
appear inside the tube of the female 95
31. Po1ydova3 three setiger larva just after release from
the female's tube 96
32. Polydova, 14 setiger larva ready to metamorphose into
the adult form 97
33. Boocard'ia pvdbosoidea, anterior end dorsal view 98
34. BooQardia* setae of the modified fifth setiger 99
35. Boooardia^ portion of a string of egg capsules 100
VII
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No. Page
36. Boccardia, three setiger larva 101
37. BoccardLa, 15 segmented larva 102
38. Dinophilus sp., dorsal view of entire worm 109
39. Dinophilus sp., egg capsule 110
40. Cirriformia luxuri-osa, anterior end, dorsal view 116
41. Cirriformia spirabranoha, anterior end, dorsal view 117
42. Ci,rri,formia luxioriosa, outline of segment 150
(after Moore, 1904) 118
43. CirrifoYmia spirdbrancha, outline of segment 150
(after Moore, 1904) 119
44. Cirrifoz-mia luxuriosa, late trochophore stage 120
45. Cirriformia luxuri-osa, metamorphosed juvenile 121
46. Ciwifozvnia luxuriosa, late juvenile stage 122
47. Ealosydna johnsoni, anterior end, dorsal view 129
48. Ealosydna, trochophore larva 130
49. Ealosydna, metatrochophore larva 131
50. Ealosydna, metamorphosed juvenile 132
Vlll
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TABLES
No. Page
1. List of Polychaetous Annelids Which Are or Were in
Laboratory Culture ..................... 13
2. Toxicity of Heavy Metals (mg/1) to Polychaetous
Annelids .......................... 14
3. The Effect of Heavy Metals on Reproduction in Laboratory
Reared Polychaetous Annelids ................ 15
4. Toxicity of the Water Soluble Fraction of Two Oils on
Polychaetous Annelids .................... 16
5. The Effects of the Water Soluble Fractions of Two Oils on
Reproduction in Two Species of Polychaetous Annelids .... 17
IX
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ACKNOWLEDGMENTS
I wish to express my thanks especially to my former graduate students
who contributed so much to the success of this study; these include
Robert Scott Carr, John H. Dorsey, Kathleen M. King, James Michael Martin,
Douglas E. Morgan, Philip S. Oshida, Fred M. Piltz, Stanley A. Rice,
Mark Michael Rossi, John F. Shisko, Diana Vermillion, and Jack Q. Word. My
appreciation is due to Dr. Gerard Belland and Dr. Denise Bellan-Santini
(Station Marine d'Endoume, Marseille, France) and Mrs. Carol Pesch and
Dr. John H. Gentile (EPA, Narragansett) for their participation in the
interlaboratory bioassay experiment. Thanks are due to Dr. Donald K. Phelps
(EPA, Narragansett) and Dr. C. M. Tarzwell (EPA, Narragansett, Retired) for
their assistance and interest in carrying out the objectives of this
research. The drawings were done by Lhisa Reish.
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SECTION 1
INTRODUCTION
Polychaetous annelids are segmented worms that are present in all
oceanic regions of the world and in all habitats. A few species have been
found in fresh water, usually upstream from estuaries. Polychaetes are
abundant in the intertidal environments, especially in sediments consisting
of silts and clays. The subtidal benthos is their preferred habitat since
they are most numerous here, both in terms of numbers of species and total
specimens. Polychaetes are also the most numerous macrofaunal component of
the soft bottom community; the percentage of polychaete species and specimens
present is generally between 30 and 50 regardless of geographical region or
depth (Knox, 1977). Polychaetes play an important role in the movement of
sediments in much the same manner as earthworms do on land. Polychaetes also
play an important role in providing food for birds and fish (Reish and Ware,
1976).
The study of marine pollution can be divided into two areas, field
studies and laboratory studies. Laboratory studies have generally focused on
the toxic effect of a single compound on an economically important organism
such as fish, with the results generally expressed as a 96-hour IC^Q, the
concentration at which 50 percent of the organisms die in 96 hours. Applica-
tion factors, generally 0.01, were applied to this figure to estimate the
long-term effect level. With the discovery of the effects of DDT on repro-
duction in birds, it became apparent that in order to obtain a clearer under-
standing of the effects of a toxicant, toxicity tests must be conducted over
an entire life cycle of the organism, including the effects on its offspring.
Because of their importance in marine environments, polychaetes were a log-
ical choice with which to study long-term effects under laboratory conditions.
i
The majority of marine invertebrates, including polychaetes, had never
been cultured previously in the laboratory; those studies that had been done
with invertebrates were pursued only through larval stages. For the most
part, mass culture methods had never been considered for marine organisms,
except possibly those of economic importance. In order to measure the long-
term effect of a pollutant on the reproduction, fertilization, larval
development, settlement, growth, and reproduction of the F generation, it is
necessary to have large quantities of specimens available on a routine basis.
Therefore, the two primary objectives of this study were as follows:
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1. To develop techniques for culturing several species of polychaetes
through their life cycle under laboratory conditions, particularly
for those species that could reproduce on a more or less continuous
basis under laboratory conditions.
2. To study the short-term and long-term effects of various toxicants,
such as heavy metals and petrochemicals, on those polychaete species
that were in laboratory culture, with emphasis on studying the
effect of these toxicants on reproduction as measured by the number
of eggs or offspring produced.
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SECTION 2
LABORATORY CULTURE PROCEDURES
WITH POLYCHAETOUS ANNELIDS
Prior to the initiation of this study, the author had two species of
polychaetes under laboratory culture; these were Neanthes catenaeeodentata
and Capitella oapitata. The N. arenaoeodentata colony was established in
1964 and had undergone about 35-40 generations in the laboratory by the
initiation of this study in late 1972. The original colony was established
from six worms collected from Los Angeles Harbor. The C. capitata colony
was started from a single female collected from Los Angeles Harbor in 1968.
C. capitata has a faster life cycle and had undergone about 40 generations
by the initiation of this study.
A total of twelve species, including N. orenaceodentata and C. eapitata,
were under laboratory culture, of which eleven had gone through at least one
complete life cycle in the laboratory and six had completed at least twenty
life cycles. C-irriformia luxuviosa was the largest worm and after two years
had not yet completed its life cycle. Those species that were cultured in
the laboratory during the course of this research are summarized in Table 1.
Additional data included in Table 1 were the number of life cycles each
species had completed, the length of the life cycle, and the status of the
colony in 1979. A manual of culture techniques for twelve species of poly-
chaetes has been included with this report (Appendix A-I) and can be found
elsewhere (Reish, 1976). Additional information concerning the culture
techniques and the usefulness of the particular species in bioassay studies
is noted below.
N. orenaceodentata (Appendix A). This species is a convenient labora-
tory bioassay organism. The adults are sufficient size and weight (2-4 cm;
50-100 mg wet weight) to permit handling with ease and to make body burden
analysis for toxicants possible. The life cycle, of three to four months
duration at 20°C, is of sufficient length to permit long-term studies.
Short-term tests can be conducted with either juveniles or adults. Survival
of controls is almost always 100 percent, and death is clearly defined. The
female lays about 150-400 large eggs (500 U in diameter) in the male's tube
and then she dies. There is no planktonic larval stage. The male cares for
the young until they leave in about one month at the 18-21 segmented stage.
The data obtained in reproductive tests provide information on whether or
not N. avenaoeodentata will reproduce at a particular concentration and the
number of eggs laid (or juveniles emerged) per female. The data can be sub-
jected to a variety of statistical tests to determine if the results are of
significance (American Public Health, 1976).
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N. arenaceodentata is the species favored by the author for bioassay
work. An established colony of 3,000-5,000 specimens requires about 30-40
hours per month to maintain. A further advantage of working with this spe-
cies is the ease in shipping live specimens to other geographical regions.
Five adult specimens are placed in a 11 x 23 cm plastic bag together with
24 ml of seawater and a small amount of resoaked Enteromorpha. The bags are
then placed in a box and sent by air mail. Successful shipments (numbering
about 80 as of December 1979) of N. arenaceodentata have been sent to labor-
atories located on the east coast of the United States, in the Gulf States,
on the Pacific coast, and in Canada.
C. capitata (Appendix B). The life cycle of this species is 30-40 days
at 20°C. The female lays about 200-400 eggs in her tube, which she incubates
for four to five days. The young larvae leave the tube of the parent and
may swim for a few hours before settlement and metamorphosis. The number of
eggs laid by the female makes it possible to obtain data on the effects of a
toxicant on reproduction and number of eggs produced. The induction of ab-
normal larvae (bifurcated posterior ends) is a method of measuring sublethal
effects (Reish, and others, 1974; Reish, 1977). Short-term bioassays can be
conducted with both trochophores and adults. Survival of controls has been
at least 90 percent, and with practice, death of specimens can be readily
distinguished. Long-term bioassays can be initiated with young Capitella
and carried out through one or more life cycles. Capi>te11a is an easily
cultured species; a colony of 3,000-5,000 worms requires about 15 hours per
month to maintain. Living specimens have been shipped by air mail to the
east coast of the United States and to Texas, Canada, and France with a good
survival rate. Approximately 25-40 specimens are placed in a 11 x 23 cm
plastic bag together with 25 ml of seawater and a small amount of resoaked
EnteromoTpha. The bags are then placed in a box and mailed.
Ctenod.Ti.lus serratus (Appendix C). Ctenodrilus is a small polychaete
that reproduces asexually by transverse fission in 12-15 days at 20°C. All
specimens were derived from a single worm collected in Los Angeles Harbor
in 1972. It is a convenient species with which to conduct toxicity tests on
reproduction since the life cycle is completed in less than a month. Sur-
vival of control specimens has been at least 95 percent. A large colony can
be cultured in five 3.78 & aquaria with only three to five hours per month
required for maintenance.
0 Ophryotrocha diadema (Appendix D). This species was described by
Akesson (1976) from specimens collected in Los Angeles Harbor in 1972. The
colony has been maintained since this original collection. 0. diadema is a
protandic hermaphrodite that completes its life cycle in 20-25 days at 20°C.
Bioassay procedures similar to those described for Ctenodvilus are used.
Survival of control specimens over a 96-hour experiment has been at least
90 percent. A colony of 3,000-5,000 specimens requires about 12-15 3.78 S,
aquaria and about ten hours per month to maintain.
Opkryotrocha puerilis (Appendix D). This species has been cultured in
Sweden by Akesson for several years. The present colony was established
from specimens brought to Long Beach by one of AJeesson's colleagues in 1974.
The methods of culturing 0. puerll^s are similar to those of 0. diadema3 but
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the life cycle is shorter in the former species, hence its more frequent use
in bioassays by the author.
Dexiospira brasi-Ziensis (Appendix E) . This is one of the many common
small spirorbid-type serpulid polychaetes that builds a tightly coiled cal-
careous tube. Specimens were collected from the surface of mussels or algae
in Alamitos Bay. D. brasi-Ziensis broods a small number of embryos in its
operculum. The number of embryos or the amount of calcareous deposition in
the tube can be related to the concentration of toxicant. While this species
requires only a month to complete its life cycle, it requires several hours
per week to maintain a small colony of less than 1,000 since the phytoplank-
tonic organism DunaZieZZa must also be cultured to provide food for
D. brasiZiensis.
Potydova 1-Lgni (Appendix F) . This is a commonly encountered species in
southern California waters, both as a bottom-dwelling adult or as a plank-
tonic larvae. Both stages were collected from nearby Bolsa Chica. The lar-
vae have been used for 96-hour bioassays, but they are not suitable for
longer tests because of'the low survival rate in culture. The number of eggs
per capsule and the number of capsules per spawning period may be convenient
sublethal parameters to study. Both larvae and adults require living
DunaZ-ieZZa as food. Several hours per week are required to maintain a colony
of about 400 because of the time required to culture the DunaZieZZa and for
feeding.
Boeccopdia probosc-idea (Appendix F) . This species of spionid polychaete
is encountered less frequently in southern California waters than
PoZydora Z-igni. All details concerning culturing and conducting bioassays
with this species are identical to those given for PoZydora Zigni in
Appendix F.
Dinophilus sp. (Appendix G) . This is a small species of an archiannelid
polychaete that appeared in some of the stock colonies of Neanthes and
CapiteZZa. Presumably it was present in the seawater transported to the
campus in 1973 by a tank truck from Marineland of the Pacific. It probably
represents an undescribed species. It is about the same size as Ophryotroehz
but can be distinguished by the presence of one anal cirrus rather than two
as in Opkryotroeha. This minute species has about a five-day life cycle,
therefore it is not a suitable test organism for a 96-hour bioassay. How-
ever, Dinophilus is especially useful for studying the effects of a toxicant
over two or three life cycles.
Cirriformia Zuxuriosa and Cirriformia spirabrancha (Appendix H). These
two closely related species are several centimeters in length and require
one to two years to complete their life cycle. Culture methods are identical
for both species. Specimens of C. 1uxw?iosa were collected from the mussel
community in Alamitos Bay, and C. spirabrancha were collected from intertidal
muddy sand flats in Alamitos Bay in Long Beach. Large numbers of specimens
may be kept in various sized aquaria, but for best results a five cm layer of
coarse sand, which prevents the worms from becoming tangled with one another,
should be placed in the aquaria. This is especially important since both
species have numerous long filiform gills arising from the notopodial region
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along the entire length of the worm, that makes the worms difficult to
separate. Adults are fed a mixture of finely chopped, dried Enteromorpha
and commercially prepared fish food flakes. A stock colony of 1,000 adults
of either species requires five to eight hours of maintenance per week.
Since these two species are large in size and have a long life cycle, they
would be excellent benthic organisms to conduct long-term bioassays and
determine body burden levels of a toxicant.
Halosydna johnsoni (Appendix I). One complete life cycle has been
carried out in the laboratory, the first such instance in the largest family
of polychaetes, the Polynoidae. Specimens were collected from the mussel
community in Alandtos Bay in Long Beach. Each female lays up to 100,000
eggs, which makes this species particularly well suited for larval testing.
The adults require a considerable amount of individual attention since they
mast be kept in separate containers because of their cannibalistic
tendencies. Adults can be maintained in a small glass tubing. Frozen brine
shrimp are fed to each specimen twice a week. This species lives much
better at 17°C than at 20°C. About 15 hours a week are required to maintain
about 500 specimens.
The feasibility of rearing polychaetes in large numbers under laboratory
conditions for use in short-term or long-term bioassays over at least one
complete life history has been demonstrated. Large populations of up to ten
species of polychaetes can be maintained by one full-time laboratory
assistant. However, it is advantageous to have more than one technician
trained on culture techniques in the event of illness, vacations, etc. Many
of these species have been successfully transported to other laboratories for
use in various types of investigations.
6
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SECTION 3
THE EFFECTS OF HEAVY METALS
ON POLYCHAETOUS ANNELIDS
The effects of six heavy metals were studied, at least in part, on nine
different species of polychaetes (Table 2) . Metals included were cadmium
(CdCl2) , chromium (CrOs) , copper (CuS04'5H20) , mercury HgCl2) , lead
(Pb(CH3COO)2'3H20) , and zinc (ZnS04-7H20) . It was necessary to add sodium
citrate as a chelating agent to the copper, zinc, and lead solutions; a sec-
ond control was used that contained this agent at the highest concentration
used. Data generated included 96-hour and 28-day LCso values (Table 2) and
effect on reproduction as measured by the number of offspring produced. The
results of much of this work dealing with the effects of heavy metals on
polychaetes have been published elsewhere (Appendix K) .
96-HOUR AND 28-DAY LC DATA
The results of these experiments can be summarized as follows:
1. Comparisons of the 28-day LCso to the 96-hour LCso data indicated
that within a species most of the 28-day values were less than the
96-hour values. The 96-hour and 28-day LCso data for copper and
mercury were previously reported for most of the copper species
(Reish, and others, 1976). Similar 96-hour and 28-day LCso values
for the species exposed to mercury may be attributable to the
static exposure conditions since mercury is a volatile element.
Thus , the mercury concentrations at 28 days were most likely less
than at 96-hours. No explanation is offered to account for the
similar results obtained with the copper experiments.
2 . In the limited number of experiments conducted with larvae , the
larval stage was more sensitive than the adult to heavy metals
(Reish, 1977, 1978; Reish and Carr, 1978). Trochophores were more
sensitive than the adults to cadmium, copper, lead, mercury, and
zinc but were more tolerant to chromium. Juvenile Neanthes were
more sensitive than adults to all metals except cadmium and
chromium. Trochophores of Eaiosydna. were more sensitive to chrom-
ium, copper, and zinc, but in contrast, were less sensitive to
mercury (Table 2) .
3. Mercury and copper were the most toxic of the six metals to all
species tested. Cadmium was the least toxic (Reish and others,
1976) .
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4. While a complete comparison is impossible since not all the experi-
ments indicated in Table 2 were completed, some generalities can be
made on the relative sensitivity of these species of polychaetes to
the metals tested.
a. Dexiospwa was the most sensitive species in the two
instances where data are available (Cu, Zn) ; the 28-day
data were about one order of magnitude lower than those
values for the other species (Table 2) .
b. C'i'Pvifo'rm'ia. luxwp'iosa. was the most tolerant species for
which data are available (Cu, Zn, Cd) (Table 2} .
c. While the relative sensitivities of the species were gener-
ally constant, one noteworthy exception was observed,
Halosydna was very tolerant to five of the six metals tested
but very sensitive to copper.
REPRODUCTION
The effect of these six metals on reproduction was studied in detail
with Ctenodvilus and Ophryotrooha which possess a short life cycle, and to a
limited extent with Capitella and Neanthes (Table 3) . The data included in
this table are the highest concentration at which reproduction occurred and
the concentration at which a statistically significant suppression of repro-
duction was noted. The findings can be summarized as follows:
1. The effect of these six metals on the reproduction of Ophryotrocha
and Ctenodrilus is in general similar; however, Ophryotrooha tends
to be the more sensitive species except to chromium. Ophryotrocha
was much more sensitive to zinc than Ctenodrilus (Reish, 1978;
Reish and Carr, 1978).
2. Supression of reproduction in Capltella occurred at a lower concen-
tration of copper (0.05 mg/1) than in either Ophpyotrocha or
Ctenodvilus (0.1 mg/1). Abnormal larvae were induced in Capitetla
at an even lower concentration, 0.01 mg/1 (Reish and others, 1974).
3. Reproductive suppression in Neanthes occurred at a very low concen-
tration of chromium, .0125 mg/1 (Oshida and others, 1976) .
4. Comparison of the 96-hour LCso data to the concentration at which
reproductive suppression occurred in Ctenodpilus and Ophryo trochee
is of special significance (Tables 2 and 3) . The concentration at
which reproductive suppression occurred was from one to two orders
of magnitude less than the 96-hour LCso f°r Ophryotrocha . In
Ctenodvilus the range in which reproductive suppression occurred
was from slightly lower than the LCso to two orders of magnitude
different (Reish and Carr, 1978) . With mercury, however, the con-
centration for suppression for both species was approximately the
same as the 96-hour LCso. These results with mercury may be attri-
butable to the static exposure conditions since mercury is a
8
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volatile element. However, no explanation is offered to account for
similar results with copper.
5. For each metal, there was a sublethal concentration at which repro-
duction occurred but was significantly reduced. Also, in the case
of Cap-itella there was a lower concentration (with chromium, copper,
and zinc) at which the reproductive rate was not affected but abnor-
mal larvae were induced (Reish and others, 1974; Reish, 1977).
Interestingly, the magnitude of difference between the 96-hour LC5Q and
that concentration at which there was no significant suppression of repro-
duction was less than one for animals exposed to mercury and copper but was
greater than one to cadmium, chromium, lead, and zinc. As stated above the
similar results in tests with mercury may be attributed to the volatile
nature of the element since the solution was not renewed during the
experiment. The action of cadmium, chromium, lead, and zinc may take a long-
er period of time to cause an effect, or the animal has an ability to excrete
at least some of the metal.
In summary, this study has shown that it is feasible to conduct toxicity
studies with polychaetes with reproduction as the parameter that is measured.
Since reproduction is obviously necessary for the survival of the species, to
be able to determine at what levels the organism can or cannot reproduce is
of vital importance to assess the effects of pollution.
-------
SECTION 4
THE EFFECTS OF PETROLEUM HYDROCARBONS
ON POLYCHAETOUS ANNELIDS
The toxicty of water-soluble fractions of No. 2 fuel oil and South
Louisiana crude oil to five species of polychaetes was determined. These
species were Cap-itella capitata^ Ctenodrilus serratus, Ophpyotrocha diadema^
Dexiospira bvas-il-iensis 3 and Cirr-iformia spirabrancha. Toxicity was re-
ported as 96-hour and 14-day LCso values for these five species (Table 4) .
In addition, the effect of these two oils on reproduction was measured for
Ctenodrilus serratus and Ophryotrocha diadema (Table 5). The results of these
experiments can be summarized as follows: (see also Carr and Reish, 1977.)
1. The water-soluble fraction of No. 2 fuel oil (a refined oil) was
more toxic to all species of polychaetes, except C-vrviformia sp.,
at both 96 hours and 14 days than South Louisiana crude.
2. In many of the experiments with both No. 2 fuel oil and South
Louisiana crude the 96-hour testing period was an insufficient
length of time to show any toxic effect to these species of
polychaetes .
3. Cap-itella was the most tolerant species tested to both oils (iden-
tical to Cirriformia sp. exposed to No. 2 fuel oil) and
Ophryotrocha sp. was the most sensitive species tested.
4. A significant suppression of reproduction was observed for
Ctenodrilus and Opkryotrocha exposed to both oils. The percent
concentration of oil at which reproductive suppression occurred was
always less than the 14-day LCso percent concentration level.
In summary, as in the case with the heavy metals study, long-term
studies with sublethal amounts of the water-soluble fraction of petrochem-
icals (the part that persists in the environment) can be conducted routinely
under laboratory conditions.
10
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SECTION 5
AN INTERLABORATORY CALIBRATION
EXPERIMENT UTILIZING
Capitella capitata
An interlaboratory calibration toxicity test involving three labora-
tories (California State University, Long Beach; U.S. EPA Environmental
Research Laboratory, Narragansett, Rhode Island; and State Marine d'Endoume,
Marseille) was planned and carried out with the assistance of Mrs. Carol
Pesch of the Environmental Research Laboratory. Dr. Gerald Bellan and
Dr. Denise Bellan-Santini were the participants from Marine d'Endoume.
Capitella capitata was selected as the test organism, and cadmium (CdCl2) was
selected as the toxicant. This is the first interlaboratory calibration ex-
periment that has been conducted with a marine organism. Since the paper
that describes this experiment and results has been published elsewhere,
only the reference to it will be given: Reish, D.J., C.E. Pesch, J.H.
Gentile, G. Bellan and D. Bellan-Santini. 1978. Interlaboratory Cali-
bration Experiments Using the Polychaetous Annelid Capitella capitata.
Marine Environ. Res. 1:109-118.
11
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REFERENCES
Akesson, B. 1976. Morphology and Life Cycle of Ophryotrocha diadema, a New
Species from California. Ophelia 15-23:35.
American Public Health Association. 1976. Standard Methods for the
Examination of Water and Wastewater. 14th Edition. APHA, AWWA, WPCF,
Washington B.C. 1193 pp.
Hinegardner, R. T. 1969. Growth and Development of the Laboratory Cultured
Sea Urchin. Biol. Bull. 127:465-475.
Knox, G. A. 1977. The Role of Polychaetes in Benthic Soft-bottom
Communities. In: Essays on Polychaetous Annelids in Memory of
Dr. Olga Hartman, D. J. Reish and K. Fauchald, eds. Allan Hancock
Foundation, Univ. of Southern California, Los Angeles, pp 547-604.
Oshida, P. S. , A. J. Mearns, D. J. Reish, and C. S. Word. 1976. The Effect
of Hexavalent and Trivalent Chromium on Neanthes arenaceodentata
(Polychaeta:Annelida). So. Calif. Coastal Water Research Project. Tech.
Mem. No. 225, 58 pp.
Reish, D. J. 1976. The Establishment of Laboratory Colonies of Polychaetous
Annelids. Thalassia Jugoslavia. 10:181-195.
Reish, D. J. 1977. Effects of Chromium on the Life History of Capitella
oap'itata. In: Physiological Responses of Marine Biota to Pollutants.
F. J. Vernberg, A. Calabrese, F. P. Thurberg, and W. B. Vernberg, eds.
Academic Press, New York, pp. 199-207.
Reish, D. J. 1978. The Effects of Heavy Metals on Polychaetous Annelids.
Rev. Int. Oceanogr. Med. 49:99-104.
Reish, D. J., J. M. Martin, F. M. Piltz, and J. Q. Word. 1976. The
Induction of Abnormal Polychaete Larvae by Heavy Metals. Marine
Pollution Bull. 5:125-126.
Reish, D. J., and R. Ware. 1976. The Food Habits of Marine Fish in the
Vicinity of Fish Harbor in Outer Los Angeles Harbor. In: Marine
Studies of San Pedro Bay, California. Part 12. Allan Hancock
Foundation and Office of Sea Grant Programs, Univ. of Southern
California, pp. 113-128.
Reish, D. J., and R. S. Carr. 1978. The Effect of Heavy Metals on the
Survival, Reproduction, and Life Cycles for Two Species of Polychaetous
Annelids. Marine Pollution Bull. 9:24-27.
12
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Table 1
List of Polychaetous Annelida Which Are or Were in Laboratory Culture
Species
He an the s arenaceodentata
Capitella capitata
Ctenodrilus serratus
Ophryotrocha diadem^
pphryotrocha puerilja
Dinophilus sp.
Dexiospira brasiliensis
Polydora ligni
Boccardia proboscidea
Ctrrifornda luxurlosa
Clrriformla spirabrancha
Halosydna brevisetosa
Stage Cultured
Life Cycle
__ Life Cycle
Life Cycle
Life Cycle
Life Cycle
Life Cycle
Life Cycle
Life Cycle
Life Cycle
Two Years
Life Cycle
Life Cycle
Number of
Generations
50+
50+
50+
50+
20+
50+
3
3
3
incomplete
1
1
Length of Life Cycle
(In Days)
120
30
10-15
10-15
20-25
5
30
50
100
probably two years
475
270-300
Manual of
Procedures
Appendix 1
Appendix 2
Appendix 3
Appendix 4
Appendix 4
Appendix 5
Appendix 6
Appendix 7
Appendix 8
Appendix B
Appendix 8
Appendix 8
Status of
Culture 1979
Maintained
Maintained
Maintained
Maintained
Maintained
Maintained
Maintained
Not in Culture
Not in Culture
Not in Culture
Not in Culture
Not in Culture
-------
Table 2 Toxicity of Heavy Metals to Polychaetous Annelids
(in mg/1)
Species Cadmium Chromium Copper Lead
Neanthes arenaceodentata
Adults
Juveniles
Capitella capitata
Adults
Troohophores
Ophryotrocha diadema
Dinophilus sp.
Halosydna johnsoni
Adults
Trochophores
Cirriformia luxuriosa
Adults
Juveniles
Cirriformia spirabranoha
Ctenodrilus serratus
Pexiospira brasiliensis
96 hr
LC50
12.0
12.5
7.5
0.22
4.2
0.8
13.0
15.0
4.3
28 day
LC50
4.0
3.0
0.7
6.1
3.5
96 hr
LC50
1.0
1.0
5.0
8.0
7.5
0.82
4.7
1.2
4.3
28 day
LC50
0.55
0.7
0.28
1.45
96 hr
LC50
0.3
0.3
0.2
0.18
0.16
0.026
0.15
0.1
0.9
0.35
0.75
0.33
0.96
28 day
LC50
0.25
0.14
0.2
0.084
0.65
0.2
0.2
0.03
96 hr
LC50
10.0
7.5
6.8
1.2
14.0
2.5
6.25
7.2
28 day
LC50
3.2
2.5
1.0
—
___
6.0
Mercury Zinc
96 hr
LC50
0.22
0.1
0.1
0.014
0.09
0.3
0.028
0.04
28 day
LC50
0.17
0.09
0.1
0.27
™ -
96 hr
LC50
1.8
0.9
3.5
1.7
1.4
0.22
6.0
2.2
> 15.0
7.1
3.0
28 day
LC50
1.4
0.9
1.25
3.0
8.6
0.16
-------
Table 3 The Effect of Heavy Metals on
Reproduction in Laboratory Reared
Polychaetous Annelids
Metal and
Species
Cadmium
Ctenodrilus serratus
Ophryotrocha diadema
Chromium
Neanthes arenaceodentata
Capitella capitata
Ctenodrilus serratus
Ophryotrocha diadema
Copper
Capitella capitata
Ctenodrilus serratus
Ophryotrocha diadema
Lead
Ctenodrilus serratus
Ophryotrocha diadema
Mercury
Ctenodrilus serratus
Ophryotrocha diadema
Zinc
Capitella capitata
Ctenodrilus serratus
Ophryotrocha sp.
Dexiospira brasiliensis
Highest Concentration
at Which Reproduction
Occurred
2.5 mg/1
1.0
0.05
0.4
0.5
0.1
0.1
0.1
0.1
1.0
0.5
0.05
0.1
0.1
2.5
0.5
0.05
Concentration at
Which Reduction in Significant
Reproduction Occurred
2.5 mg/1
1.0
0.0125
0.1
0.05
1.0
0.05
0.1
0.25
1.0
5.0
0.05
0.1
0.05
0.5
0.5
0.01
15
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Table 4 Toxicity of the Water Soluble Fraction of
Two Oils on Polychaetous Annelids
Species
Capitella capitata
Ophryotrocha diadema
Cirriformia spirabrancha
Ctenodrilus serratus
Dexiospira brasiliensis
Fuel Oil #2
96 hour
LC50
no deaths
at 100%
67% in
30% oil
no deaths
at 100%
47
91
14 day
LCSO
70%
28
70
38
56
So . Louisiana Crude
96 hour
LCSO
no deaths in
100% oil
65
no deaths in
100% oil
96% survival
in 100% oil
14 day
LCSO
70% survival
in 100% oil
55
50
72% survival
in 100% oil
-
16
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Table 5 The Effects of the Water Soluble Fractions
of Two Oils on Reproduction in Two Species of Polychaetous Annelids
Oil
So. Louisiana Crude
Fuel Oil tt2
Highest Concentration
With Reproduction at
21 days
Ctenodrilus
75% WSF
25
Ophryotrocha
75% WSF
10
Concentration at Which a
Significant Difference in
Reproduction Occurred
Ctenodrilus
50% WSF
25
Ophryotrocha
50% WSF
10
17
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APPENDIX A
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELIDS:
Neanthes arenaceodentata
CULTURE TECHNIQUES
Equipment and Supplies
The equipment and supplies needed to maintain a continuous supply of
Neanthes arenaceodentata at the rate of 1,000 to 2,000 per month are neither
elaborate nor expensive. Approximately 20 m^ of either shelf or table sur-
face is needed to rear this quantity of worms. While it is not essential, a
constant temperature laboratory set at 19±1°C, insures against the possible
effects of either abnormally high (>25°c) or low (<17°C) temperatures.
N. arenaceodentata is capable of withstanding temperature fluctuations of
19±4°C without any measurable stress. Natural, filtered seawater at normal
salinity (35 °/oo) is a satisfactory source of seawater; however, success
has been obtained using artifical sea salts. N. arenaceodentata can tolerate
salinity variations of 35±5 °/oo without adverse affects. A running seawater
system is not required; approximately 1,500 liters of seawater are needed per
month to raise 1,000 to 2,000 specimens. A central compressed air system is
the most convenient source of air, but several aquaria pumps could supply
sufficient air to culture N. arenaceodentata. A dissecting microscope with
light is required to examine specimens prior to use in an experiment. A hand
lens and flashlight are helpful in examining the condition of the eggs and
larvae. Glass containers required include 3.78 X- (1 gallon) jars, standard
petri dishes (may be plastic) and 37.8 & (10 gallon) to 56.7 & (15 gallon)
aquaria. Additional supplies needed include 12 cm diameter plastic covers
for the glass jars, forceps, fine brushes, plastic tubing, air stones, an
outside aquaria filter system, charcoal, and glass wool. Quantities of the
dried green alga Enteromorpha sp. and commercially prepared alfalfa flour are
needed for a food supply.
Life History
N. arenaceodentata is widely distributed throughout the world; it has been
collected in Europe, New England, Florida, California, Baja California, and
the Central Pacific. It is present within the sediment, especially in estu-
arine environments. It may be present in large numbers; as many as 1,000/m2
have been observed. Specimens live in the benthos where they construct tubes
consisting of sediment, food, and feces, which is held together by mucus.
18
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Specimens measure up to about four cm in length and are tan to yellow-
tan in color. Since many species of Family Nereidae appear alike under low
magnification, Neanthes can be distinguished from the other species by the
possession of a continuous ring of paragnaths around areas V - VII of the pro-
boscis (Figure 1) and by a hooked tip present on the blade of the neuropodial
heterogomph seta (Figure 2).
It is impossible to distinguish immature males from females on the basis
of morphology. However, a behavioral difference occurs that can be utilized
to distinguish sexes. Males will fight males and females will fight females.
This fighting behavior consists of extending the jaws to grasp the opposing
worm (Figure 3). They can be cannibalistic. A male and female placed to-
gether will come alongside each other and lie side by side. If one specimen
has constructed a tube, the two will lie within the tube until eggs are laid.
It is possible to detect this behavior difference in young specimens possess-
ing 20-25 segments. In order to determine the sex of an immature worm, place
it in a petri dish with a female containing developing eggs within her coelom
and observe the behavior of the two when they come in contact with one
another. If they come to lie side by side, then the immature worm is a male.
The eggs of Neanthes are formed within the walls of the parapodia.
Shortly thereafter, they break free and mature within the coelom. The muscle
cells, especially the longitudinal muscles, slough off and are digested by
the eleocytes that transfer material to the maturing eggs. The mature eggs
of Neanthes are large, measuring 500-600 y in diameter. During maturation of
the egg, about 75 percent of the body weight of the female is transferred to
the eggs. The eggs are probably passed through bilateral breaks in the body
wall between successive parapodial lobes. The fertilized eggs are clumped
in the central part of the tube around the mid-body region of the male.
Development of the young proceeds within the tube until the 18-21 seti-
gerous segment stage (Figures 4-6). The developing eggs are incubated by
the male who circulates water with his body undulations. These undulations
provide a continuous source of oxygenated water that is essential for devel-
opment of the eggs. The fertilized eggs are yellow as the result of the
large amount of yolk material, but as the yolk supply is utilized during
development, the young become tan in color. Neanthes lacks the free-swimming
trochophore larval stage characteristic of most polychaetes. The shape of
the larval body is distorted as a result of the large quantities of yolk
material; in fact, the yolk bodies may be observed in the future digestive
tract of larvae up to nearly the 18 setigerous segment stage (Figure 4-6).
The young worms leave the tube of the parent when they attain 18-21 setiger-
ous segments (Figure 7). Shortly after leaving the parent's tube, they
construct a tube and commence feeding. Under laboratory conditions, it takes
three to four months for Neanthes to complete its life cycle at about 20-22°C.
Presumably, a longer time is required in the field because of lower
temperatures. Under laboratory conditions, the male is capable of reproduc-
ing at least a second time after the conclusion of the initial incubation
period.
19
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Techniques of Handling Neanthes
Adults—
Stock colonies of adults can be maintained at room temperature in aquaria
of about 37.8 to 56.8 liter size with a maximum population concentration of
approximately 75-100 worms per aquaria. The aquarium must be provided with
at least two air stones and an outside filter system for aeration (Figure 8).
Add about 1.5 g of alfalfa flour and/or Entevomovpha per 56.8 £ aquarium per
week. The alfalfa flour should be mixed with seawater prior to use.
If all specimens are going to be used for experimental studies within a
month, it will not be necessary to change the water in the aquarium; however,
the water level should be checked periodically and double distilled water
added to keep the level constant. The air stem in the outside filter system
must be checked weekly and cleared of accumulated salts. If the specimens
are not being utilized within a month, the water should be changed at about
four to six weeks.
Large quantities of the green alga Enteromorpha sp. can be collected
from estuaries during high tide. This genus is widespread in temperate
estuarine environments of the world and frequently flourishes in the spring
or fall months. The alga is hand-collected and washed at the site to remove
as much adhering debris and sediments as possible. It is then spread out on
chicken wire and allowed to air-dry. After drying, it can be stored in plas-
tic bags for an indefinite period of time. In areas of high humidity it may
be necessary to dry it in a warming oven (30-35°C). The dried Enteromorpha
is soaked in seawater prior to refeeding the worms. The alga should be
kneaded so that the individual branches of the alga separate from one
another. The alga is then ready to feed to the worms.
Reproductive Specimens—
Females with maturing eggs in the coelom appear yellow-orange. Since
sexual maturity will be reached about two months after the aquarium is estab-
lished with juvenile worms, all specimens should be removed from the aquarium
and females with eggs should be set aside in a petri dish. Remove a large
worm with a fine paint brush from the aquarium, and place it in the dish with
the female. Determine the sex of the second worm by the fighting response.
If the unknown worm is a male, establish a 3.78 & jar with the couple, and
feed them Enteromorpha. Examine them two to three times a week until the
eggs are laid. Note when the eggs were laid and continue to feed the worms
on schedule. Examine the developing embryos every other day beginning at two
weeks for emergence from the parent's tube. Since the male can become canni-
balistic after the emergence of his offspring, it is important to remove the
parent. The male may be utilized a second time or discarded. Do not use it
as an experimental animal.
20
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Larvae—
No special care is required for specimens having less than 18-21 seti-
gerous segments. These nonfeeding larvae are cared for by the male parent
within his tube.
Immature Worms—
After the young worms leave the tube of the parent, they should be
removed as soon as possible to prevent being eaten by the parent. Place 50
to 75 young worms in a 56.8 & aquarium with about 50 & of seawater. Feed
them 1.5 g of alfalfa flour initially and once weekly. Cut down the amount
of food fed if the majority of the food has not been eaten.
Problems in Culturing—
Most of the difficulties in culturing Neanthes can be attributed to the
lack of regular, systematic care. Since there may be a large number of vari-
ous sized containers such as petri dishes, 3.78 & jar aquaria, and 17.8 to
56.8 & aquaria, it is recommended that each container be numbered and records
kept. It is possible to minimize some of the routine record keeping by keep-
ing basic data written on the outside of the container. Arranging the
containers by stages will facilitate routine observations and feeding. It is
recommended that all routine culture care be done on a specific day each week.
Feeding problems—The two primary feeding problems are insufficient
soaking and kneading of the algae and overfeeding. Both problems lead to the
appearance of a white fungal growth over the surface of the algae that the
worms cannot penetrate to feed. As a general rule, it is better to underfeed
than to overfeed. The individual filaments of the soaked and kneaded
Entevomovpha should separate from one another when placed in an aquarium. If
all the food is in one clump, the worm will not feed.
Enemies—Fungal growth can cause the worm to abandon its tube and build
a second one elsewhere. Microorganisms, such as protozoa and copepods, may
cause problems by eating the food or attacking the worm, but this has not
been proven as yet. Amphipods may feed upon developing embryos since larvae
disappear from the parent's tube whenever these crustaceans become estab-
lished in the aquarium.
Cannibalism—As discussed above, like sexes fight, and it is possible
for one to eat the other. Like sexes should never be placed together within
a petri dish. Also, do not overcrowd worms in aquaria.
Abandonment of fertilized eggs by male—About 10 to 15 percent of the
time the male may abandon his tube of developing eggs. Eggs left in a tube
abandoned by a male will not survive. This results in the death of the
zygotes, and attempts to provide aeration for these abandoned eggs have been
successful only with a magnetic stirring device (Hinegardner, 1969)- The
cause of this abandonment is unknown.
21
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Parent male eating developing eggs or larvae—About 10 to 15 percent of
the time male parent will eat his offspring, especially during the late stages
of development. The amount of food present does not seem to alter this be-
havior. The cause of this behavior is unknown. Since upwards to one-third
of the fertilizations may result in either abandonment or cannibalism of the
offspring, the only apparent solution of this problem is to allow for this
amount of loss in culturing this species.
TOXICITY TEST
Equipment and Supplies
In addition to the equipment and supplies listed above, 500 ml Erlenmeyer
flasks or other suitable containers, for example a Carolina dish, culture
dishes, etc. and white enamel pans are needed.
96-Hour Experiments with Juveniles or Adults
Remove clumps of Neanthes from the stock colony, and place them in a
white pan filled with enough seawater to reach a depth of one cm. Allow the
worms to free themselves from their tube masses if possible so as to minimize
the amount of handling. Touching one end of the tube mass with a brush may
hasten the worm's exit. Transfer the worm to a petri dish containing sea-
water. Several worms can be placed within the same dish if they are not left
together for more than one hour. Separate out any female with developing
eggs; the female will appear orange in color and after some practice all but
the ones with ova in the early stages of development can be recognized with-
out the use of a microscope. Do not use any females with developing eggs as
test specimens. Females with developing ova can be utilized for reproductive
stock. Examine all worms under a dissecting microscope for presence of ova
or injuries. Discard all injured specimens. Place a single worm in a 500 ml
Erlenmeyer flask containing 100 ml seawater and toxicant. Close the flask
with a rubber stopper.
Use 20 specimens per concentration and a minimum of five concentrations
of toxicant plus control. Examine each worm daily for death. Discard all
specimens at the end of the test.
Death in Neanthes—
Death in Neanthes is defined as the absence of movement when the flask
is gently rolled or when a worm is gently poked. A dead Neanthes typically
appears white; its proboscis is everted exposing its jaws, and it has aban-
doned its mucoid tube.
Many behavioral changes in Neanthes can be observed that will assist the
experimenter in determining the health of the worm. If the concentration is
highly toxic, the worm will react violently by everting its proboscis and
moving its body in twisting motions. If this activity begins when the worm
is initially placed with the toxicant, then death will follow within an hour
or less. A similar behavioral response is elicited if Neanthes is placed
directly into fresh water. If the twisting activity begins several hours or
more later, the movement will not be as violent and death may take days.
22
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Tube building also is an indication of the well-being of a worm. A
healthy Neanthes will construct a mucoid tube on the bottom or along the side
of the container. Slightly stressed worms will construct the mucoid tube
along the side of the container at the air-water interface. With additional
stress or effect of a toxicant the worm will fail to construct a mucoid tube.
Long-Term Experiment with Adults (28 days)
Separate out and examine the worms to be used in the experiment as out-
lined above. For a 28-day toxicity test the solution should be renewed a
minimum of every four days. It will be necessary to feed Neanthes because
of the length of the test. Soak dried Enteromorpha sp. in a petri dish con-
taining seawater. Knead the alga so that it is well soaked. Use about 0.1 g
of dried Enteromovpha sp. (0.2 g wet weight) per worm. The Enteromorpha sp.
should be soaked in the same concentration of seawater plus toxicant in which
it is going to be used. Place the appropriate amount of kneaded EnteTomonrpha
sp. for one Neanthes in a petri dish at the specific concentration of toxi-
cant where the worm will be tested. Place a single Neanthes in the center of
algal mass. Continue this procedure until all experimental worms have been
so separated. Transfer algae and worm to experimental flask, and commence
the experiment. Examine daily for the first 96 hours, then at 7, 14, 21, and
28 days for death. (This experiment can serve also as a 96-hour experiment.)
Discard all specimens at the conclusion of the experiment.
Death—
The description of death in Neanthes given under the 96-hour toxicity
test applies to this long-term experiment with some additional behavioral
changes as the result of the introduction of Enteromorpha sp. as food. If
Neanthes is able to construct a tube in a moderately toxic solution it will
build one along the sides of the container at the air-water interface. It
may or may not incorporate the alga in its tube; if it does, it may or may
not feed upon the alga. The amount eaten per unit time may be less than ob-
served for specimens in the control. Neanthes in control solutions or non-
toxic solutions will construct mucoid tubes in the alga on the bottom of the
container. Fecal pellets will be noted within a 24-hour period.
Long-Term Experiments to Study the Effects of a Toxicant on Reproduction,
Egg Production, and F Larval Survival—
Place Neanthes tube masses in white enamel pans and separate out speci-
mens weighing about 10-15 mg wet weight, (35-45 setigerous segments). The
worms have been out of the parental tube for seven to ten days. Place four
worms in a petri dish; examine all specimens under the dissecting microscope;
and discard all injured ones. Place the four worms in a 3.78 £ glass jar
containing 2,500 ml of toxicant. It may be necessary to use a brush to free
the worms from the petri dish. Add 0.3-0.4 g wet weight (0.2 g dry weight)
of kneaded Enteromorpha sp. to each jar. Aerate with an air stone. It is
convenient to use about 15 aquaria (60 worms) per concentration to take into
account any unusual sex ratio or death of worms at the higher concentrations.
Add food once a week. For the second and third feeding, add about 0.45 g wet
weight (0.3 g dry weight). For feedings beyond the third week, add 0.6 g wet
weight (0.4 g dry weight). Do not add any food if the worms have failed to
23
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eat during the previous week. For the best results in long-term static bio-
assays, the water should be renewed every three weeks.
If the purpose of the experiment is to study the effect of the toxicant
on egg production, then examine each female for the presence of developing
ova within her coelom. All jars containing females with developing ova
should be marked to facilitate examination every two to three days to see
whether or not the eggs have been laid. When the eggs have been laid, care-
fully remove the tube mass containing the eggs to a petri dish, and count
under a dissecting microscope the number of eggs present. Discard the eggs
after counting. Maintain the jar if another male and female are present; if
not, either transfer the specimens to another jar within the same concentra-
tion series or destroy.
If the purpose of the experiment is to study the effect of the toxicant
on the FI or F2 generation, follow the procedure outlined above through the
egg laying stage, but do not remove the tube mass containing the eggs.
Instead, remove all other tube masses from the jar with a pair of long for-
ceps. Record the date of egg-laying and begin observing the jar on a daily
basis after 15 days for emergence of juvenile worms from the parent's tube
mass. Such juveniles should possess a dark line extending throughout much
of their length. This is food visible in the gut and indicates that the
worm has commenced feeding. These juveniles can be removed from the jar,
counted, and either discarded or used to establish a new jar according to the
procedures outlined above, if the study is planned to extend through the
F2 generation.
24
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BIBLIOGRAPHY
Abati, J. L.,and D. J. Reish. 1972. The effects of lowered dissolved oxygen
concentrations and salinity on the free ainino acid pool of the polychae-
tous annelid Neanthes arenaoeodentata. Bull. So. Calif. Acad. Sci.
71:32-39.
Anderson, J. W. 1977. Responses to sublethal level of petroleum hydro-
carbons; are they sensitive indicators and do they correlate with tissue
contamination? In: Fate and Effects of Petroleum Hydrocarbons in
Marine Organisms and Ecosystems. D. A. Wolfe, ed. Pergamon Press,
New York, pp. 95-114.
Anderson, J. W. 1977. Effects of petroleum by hydrocarbons on the growth of
marine organisms. Rapp. P.-v. Reun. Cons. int. Her 171:157-165.
Cripps, R. A., and D. J. Reish. 1973. The effect of environmental stress on
the activity of malate dehydrogenase and lactate dehydrogenase in
Neanthes arenaceodentata (Annelida:Polychaeta). Comp. Biochem. Physiol.
46B:123-133.
Davis, W. R., and D. J. Reish. 1975. The effect of reduced dissolved oxygen
concentration on growth and production of oocytes in the polychaetous
annelid Neanthes avenooeodentata. Rev. Int. Oceanogr. Med. 37-38:3-16.
Herpin, R. 1926. Recherches biologiques sur la reproduction et le develop-
pement de quelques Annelides polychetes. Soc. Sci. Nat. 1'ouest France,
Nantes, Bull. ser. 4, vol. 5, 250 p.
Mearns, A. J. 1974. Toxicity studies of chromium. So. Calif. Coastal Water
Res. Proj., Annual Rept. pp. 15-18.
Oshida, P. S. 1976. Effects of chromium on reproduction in polychaetes. So.
Calif. Coastal Water Res. Proj., Annual Rept., pp. 161-167.
Oshida P. S. 1977. A safe level of hexavalent chromium for a marine invert-
brate. So. Calif. Coastal Water Res. Proj., Annual Rept., pp. 169-180.
Oshida, P. S., A. J. Mearns, D. J. Reish, and C. S. Word. 1976. The effects
of hexavalent and trivalent chromium on Neanthes avenaceodentata
(Polychaeta:Annelida). So. Calif. Coastal Water Res. Proj., TM No. 225,
58 pp.
25
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Oshida, P. S., and D. J. Reish. 1974. The effect of various water tempera-
tures on the survival and reproduction in the polychaetous annelids:
Preliminary Report. In: Marine Studies of San Pedro Bay, Calif. Part 3.
Allan Hancock Foundation, pp. 63-77.
Oshida, P. S., and D. J. Reish. 1975. Effect of chromium on reproduction in
polychaetes. So. Calif. Coastal Water Res. Proj., Annual Rept.
pp. 55-60.
Pesch, C. E., and D. Morgan. 1978. Influence of sediment in copper toxicity
test with the polychaete Neanthes arenaceodentata. Water Res.
12:747-751.
Petrich, S. M., and D. J. Reish. 1979. Effects of aluminum on survival and
reproduction in polychaetous annelids. Bull. Environm. Contain. Toxicol.
23:698-702.
Raps, M. E., and D. J. Reish. 1971. The effects of varying dissolved oxygen
concentrations on the hemoglobin levels of the polychaetous annelid
Neanthes arenaceodentata. Marine Biol. 11:363-368.
Reish, D. J. 1957. The life history of the polychaetous annelid Neanthes
caudata (delle Chiaje), including a summary of development in the
Family Nereidae. Pacific Sci. 11:216-228.
Reish, D. J. 1966. Relationship of polychaetes to varying dissolved oxygen
concentrations. Third International Conf. Water Pollution Res., Munich.
3:199-216.
Reish, D. J. 1970. The effects of varying concentrations of nutrients,
chlorinity, and dissolved oxygen on polychaetous annelids. Water Res.
4:721-735.
Reish, D. J. 1974 (1976). The establishment of laboratory colonies of
polychaetous annelids. Thalassia Jugoslavia. 10:181-195.
Reish, D. J. 1977. The role of life history studies in polychaete system-
atics. In: Essay on polychaetous annelids in memory of Dr. Olga Hartman.
D. J. Reish and K. Fauchald,eds. Allan Hancock Foundation, Univ. of
So. California, pp. 461-476.
Reish, D. J. 1978. The effects of heavy metals on polychaetous annelids.
Rev. Int. Oceangr. Med. 49:99-104.
Reish, D. J., and M. C. Alosi. 1968. Aggressive behavior in the polychae-
tous annelid Family Nereidae. Bull. So. Calif. Acad. Sci. 67:21-28.
Reish, D. J., J. M. Martin, F. M. Piltz, and J. Q. Word. 1976. The effect
of heavy metals on laboratory populations of two polychaetes with com-
parisons to the water quality conditions and standards in southern
California. Water Res. 10:299-302.
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Reish, D. J., T. V. Gerlinger, C. A. Phillips, and P. Schmidtbauer. 1977.
Toxicity of formulated mine tailings on marine polychaeta. Rept. to
Environment Canada, Winnipeg, Manitoba. Marine Biological Consultants,
Inc., Costa Mesa, California, 104 pp.
Reish, D. J., and T. E. Richards. 1966. A culture method for maintaining
large populations of polychaetous annelids in the laboratory. Turtox
News. 44:16-17.
Reish, D. J., and G. C. Stephens. 1969. Uptake of organic material by
aquatic invertebrates, V. The influence of age on the uptake of glycine-
C14 by the polychaete Neanthes arenaceodentata. Marine Biol. 3:352-355.
Rossi, S. S. 1976. Interactions between petroleum hydrocarbons and the
polychaetous annelid Neanthes arenaceodentata: Effect on growth and
reproduction; fate of diaromatic hydrocarbons accumulated from solution
on sediments. Doctoral Dissertation, Texas ASM University, College
Station, Texas, 94 pp.
Rossi, S. S. 1977. Bioavailability of petroleum by hydrocarbons from water,
sediments, and detritus to the marine annelid Neanthes arenaceodentata..
Proc. of 1977 oil spill conference, New Orleans, La., American
Petroleum Institute, Washington, D.C., 621-625.
Rossi, S. S., and J. W. Anderson. 1976. Toxicity of water-soluble fractions
of No. 2 fuel oil and South Louisiana crude oil to selected stages in
the life history of the polychaete, Neanthes arenaoeodentata. Bull.
Envir. Contain. Toxicol. 16:18-24.
Rossi, S. S., and J. W. Anderson. 1977. Effect of No. 2 fuel oil and South
Louisiana crude oil water soluble fractions on hemoglobin compensation
and hypoxia tolerance in the polychaetous annelid, Neanthes
arenaceodentata. Marine Sci. Communs. 3:117-131.
Rossi, S. S., and J. W. Anderson. 1977. Accumulation and release of fuel-oil
derived diaromatic hydrocarbons by the polychaete Neanthes
arenaeeodentata. Marine Biol. 39:51-55.
Rossi, S. S., and J. W. Anderson. 1978. Effects of No. 2 fuel oil water-
soluble-fractions on growth and reproduction in Neanthes arenaceodentata
(Polychaeta:Annelida). Water, Air, Soil Pollut. 9:155-170.
Rossi, S. S., J. W. Anderson, G. S. Ward. 1976. Toxicity of water-soluble
fractions of four test oils for the polychaetous annelids, Neanthes
avenaoeodentata and Capitefla capi-tata. Environ. Pollut. 10:9-18.
Rossi, S. S., and J. M. Neff. 1978. Toxicity of polynuclear aromatic hydro-
carbons to the polychaete Neanthes arenaceodentata.. Marine Pollut.
Bull. 9:220-223.
27
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Figure 1. Neanthes apenaoeodentata, anterior end, with everted proboscis.
28
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Figure 2. N. arenaceodentata, neuropodial heterogomph f alcigerous seta with
hook at tip of the blade.
29
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U)
o
Figure 3. N. arenaeeodentata of the same sex in fighting position.
-------
Figure 4. N. arenaceodentata, three segmented stage.
31
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Figure 5. N. arenaceodentata, four segmented stage.
32
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Figure 6. N. arenaoeodentata, 12 segmented stage.
33
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Figure 7. N. arenaoeodentata, juvenile, with 21 segments, which has left
the parent's tube and commenced feeding.
34
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Figure 8. One gallon (3.78 &) aquarium system fitted with an outside filter
system used to culture Neanthes sp. adults, especially those
specimens approaching sexual maturity.
35
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APPENDIX B
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELID:
Capitella oapitata
CULTURE TECHNIQUES
Equipment and Supplies
The equipment and supplies required to maintain a population of 3,000-
5,000 specimens of Capitella are neither elaborate nor expensive. Approxi-
mately three to four m^ of shelf space is needed to culture and maintain this
size population of Capitella. For best results, it is advantageous, although
not essential, to culture Capitella in a constant temperature laboratory
maintained at 19±1°C. Elevated temperature, especially above 23-24°C may
cause fouling of the food, which leads to the death of the specimens within
the colony. Growth rate below about 17°C is markedly reduced leading to an
increase in length of time required for completion of its life cycle. Natur-
al, filtered seawater at normal salinity (35 °/oo) has been the most
satisfactory media to cultur^ Capitella; however, it is capable of withstand-
ing lower salinities (25 °/oo). A running seawater system is not required;
approximately 100-150 liters of seawater are needed per month to maintain a
population of 3,000-5,000 specimens. A piped compressed air system fitted
with many outlets is the most convenient source of air for Capitella; how-
ever, a few aquaria pumps could supply sufficient air. A dissecting
microscope with a light source is required for examination of specimens prior
to use in experiments. Glass containers required include 3.78 & jars and
standard sized glass or plastic petri dishes. Additional supplies include
12 cm diameter plastic covers for the jars, jeweler's forceps, fine brushes,
plastic tubing, and air stones. Dried quantities of the green alga
Entevomorp'ha sp. and dried fish food flakes are needed as a food supply.
Life History
Capitella is a cosmopolitan species that is generally found in estuarine
waters. It has been described as a non-competitive or opportunistic species
because it flourishes in the absence of other polychaete species (Barnard,
1970). Capitella is found in large numbers in the vicinity of domestic out-
fall sewers, and it has been used as an indicator of such altered environments
(Pearsons and Rosenberg, 1978). Grassle and Grassle (1976) have described
six sibling species of Capitella that are distinguished by the morphology of
the anterior and posterior ends, the arrangement of teeth on the hooded
36
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hooks, the weight of the specimens, method of reproduction, the diameter of
eggs, the number of eggs laid, and the length of larval life. Using the
sibling species classification of Grassle and Grassle, the present population
of Capitella belongs to Type I.
Specimens generally measure less than two cm in length although larger
individuals occur especially in the vicinity of domestic outfalls. Capitella
is one of the few species of polychaetes that exhibit sexual dimorphism.
Specialized genital hooks appear on the mid-dorsal region of setigerous seg-
ments eight and nine in males (Figure 9). Females lack these setae (Figure
10). Young adults may be sexed easily by observing the anterior region under
the dissecting microscope and checking for the presence or absence of these
genital hooks.
Copulation occurs, but it is rarely observed. Sperm are transferred to
the female with fertilization occurring either internally or at the time of
discharge of the eggs. The tube of Capi-tetta is a loosely constructed struc-
ture that consists of fecal material, substrate, and potential food. The
tube is open at either end, and the specimen is capable of building another
one if the initial tube is abandoned. The female places the fertilized eggs
around the inner surface of her tube (Figure 11). The eggs remain fixed in
this position, presumably by a mucoid secretion, until the trochophore stage.
There is a considerable amount of variation in the number of eggs laid, but
under laboratory conditions, it usually ranges from 200 to 400.
The female incubates the fertilized eggs during the early developmental
stages. Incubation consists of periodic body undulations of the female that
circulate water through the tube. These undulations are apparently essential
for development because attempts to provide aeration to developing zygotes
following female abandonment have been unsuccessful. The fertilized eggs are
initially white and measure about 250 y in diameter. As development contin-
ues, the zygotes become darker, and at the trochophore stage they appear
grey-green.
The trochophore stage is reached about four to six days after egg-laying
(Figure 12). Trochophores are capable of moving freely within the tube
either by ciliary movement or by contraction of longitudinal muscles. The
trochophore may either swim free of the tube and become planktonic or proceed
directly into the metatrochophore stage (Figure 13) and begin to form its own
tube as a side branch from the parent tube. The planktonifc trochophore, if
it occurs, is of short duration, and it soon settles to the substrate and
develops into the metatrochophore stage.
The trochophore and metatrochophore stages last one to two days before
resembling a juvenile adult (Figure 14), Growth is rapid and temperature
dependent. Eggs begin to develop in the coelom of the maturing female in
about 20 days, and fertilized eggs are laid at 25-40 days at 20°C.
Both sexes are capable of reproducing more than once. Females have been
observed to have had three successful egg layings under laboratory conditions.
The second egg laying occurs about five to ten days after the first brood
has left the tube. Under laboratory conditions, both sexes appear to lose
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their vigor with increasing age as indicated by body color. The characteris-
tic internal blood red color of young adults becomes dull red in color with
increasing age.
Techniques of Handling Capitella
Adults—
Stock colonies of adults can be maintained at 20°C in 3.78 £ (1 gal)
jars (Figure 15). Add 2,500 ml filtered seawater to each jar. Provide area-
tion with an air stone connected to a plastic tube and air supply. Adjust
the pressure in the system to a low level. The top of the jar can be covered
with a glass or plastic top to minimize evaporation. Many adults (10-30)
can be placed in a single jar since CapitelZa does not exhibit cannibalistic
behavior. Add about 0.15 g dried Enteromorpha sp. per week to each jar.
Commercial fish flakes, Tetramin, or Biorell are useful food; however, prior
to use these flakes should be ground into a fine powder. Add about 0.1 g of
this fine powder per jar per week.
If specimens are not being removed from a particular jar for experimen-
tation, the contents of one jar should be divided into two or three
additional jars about every three weeks. Decant most of the seawater from
the jar until about 100 ml remains. Pour the remaining water and worm tube
masses into a few petri dishes and allow them to go undisturbed for several
minutes. Capitella will usually crawl free from its-mucoid tube. Remove the
specimen with a fine brush and place into a new dish with seawater. Water
should be changed and food added at this time. If specimens are being re-
moved from a particular jar for experimentation, it will not be necessary to
divide the population; however, the water should be changed about every three
weeks. The colony can be reduced in size, but still maintained at a reduced
expenditure of time, by placing some of the jars in a cold bath maintained
at 15-17°C. Add food to each jar about twice a month. A colony can be
maintained for about two months under these conditions without examination or
changing the water. The colony can be built up by returning the jars to
20°C and following the above culture procedures.
Trochophore Larvae—
Many of the techniques used to handle trochophores (Figure 12) and meta-
trochophores were outlined under the adult culture section above. No special
techniques are required for handling the larvae in a mixed population.
Trochophores can be freed from their parent's tube under the dissecting
microscope and 25 pipetted (with a Pastuer pipette) into a separate jar. The
initial food should consist of about 0.1 g of finely ground Enteromorpha sp.
or powdered Tetramin.
Metatrochophore Larvae—
The metatrochophore larvae in Type I of Capitella is short-lived and
does not require any special techniques (Figure 13).
Food Preparation—
The green alga, Enteromorpha sp. is collected from estuaries during high
tide. This genus is widespread in the temperate estuarine environments of
the world and frequently flourishes in the spring or fall. The alga is
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hand-collected and washed at the site to remove as much adhering debris and
sediments as possible. Then it is spread out on chicken wire and allowed to
air-dry. When it is dry, it is collected and stored in plastic bags for an
indefinite period of time. In areas of high humidity it may be necessary to
dry it in a warming oven (30-35°C). Prior to feeding, Enteromorpha sp.
should be soaked in seawater and kneaded.
Enteromorpha sp. powder is prepared by grinding up the alga in a mortar
and pestle until fine. The powder may be added directly to the jar or mixed
with seawater prior to feeding. Tetramin is ground into a powder in the same
way as Enteromorpha sp.
Problems in Culturing—
Most of the difficulties and failures in culturing Capitella can be
attributed to the lack of regular, systematic care. Since there may be a
large number of jars involved, it is recommended that the basic data be
written on the outside of the jar. Arranging the jars by stages facilitates
the routine observations and feedings. It is recommended that all routine
culture care be done on a specific day each week.
Feeding problems—The two primary feeding problems are feeding too much
food or not soaking or kneading the alga sufficiently prior to use. Both
situations lead to appearance of a white fungal growth over the surface of
the alga that the worms cannot penetrate to reach the food. The individual
filaments of the soaked and kneaded Enteromorpha sp. should be able to sep-
arate from one another when placing it in the jar. White fungal growth can
also occur when too much Tetramin powder is added. Each jar should be exam-
ined prior to feeding to determine whether or not additional food is required.
Enemies—Fungal growth can result from overfeeding, improper condition-
ing of food, or insufficient dissolved oxygen. All these causes of fungal
growth can be prevented by proper and periodic care. Microorganisms, espe-
cially ciliated protozoans, nematodes, and copepods, are almost always
present in the stock colonies, and they are almost impossible to eliminate.
Whether or not they feed upon Capitella is unknown, but they compete for food.
If the population of these organisms becomes large, they can be minimized by
removing the clumps of worms from the aquarium just prior to feeding and
placing them in a new container with filtered seawater.
Abandonment of Fertilized Eggs by Female—Occasionally the female may
leave her tube of fertilized eggs. Attempts to provide aeration for these
abandoned eggs have been unsuccessful. The only known cause of this condi-
tion is either rough handling or too much handling when examining the worm
under the dissecting microscope. Abandonment of the tube also occurs within
the stock colony; the causes of this problem are unknown, and no solution is
known. Fortunately, this situation does not occur frequently enough to war-
rant searching for a solution.
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TOXICITY TEST PROCEDURES
Equipment and Supplies
In addition to the equipment and supplies listed above, the following
supplies are needed to conduct toxicity tests with Capitella: 500 ml Erlen-
meyer flasks (or Carolina dishes, petri dishes, etc.), stender dishes, and
white enamel pans.
Collection and Examination of Experimental Animals
Remove clumps of Capitella from stock colonies, and place them in white
enamel pans with sufficient seawater to reach a depth of one cm. Since
Capitella lives within a mucoid tube, it is convenient to allow them
to crawl free of their tubes. If a large number of specimens is required
it is advantageous to have several pans containing tube masses of Ca.pite'i'la,.
As the worm frees itself from the tube, remove it with a fine brush, and
place it in a petri dish. Proceed to the next pan, and continue to examine
each pan periodically until enough specimens have been obtained. Examine
each specimen under the dissecting microscope, and remove any female contain-
ing developing ova within her coelom. Developing ova appear as bilateral,
segmentally arranged masses of white tissue ventrally located beginning
about segment 12. These females can be utilized for establishing reproduc-
tive colonies. Remove and destroy any injured specimens.
If trochophore larvae are to be used as experimental organisms, examine
clumps of Capi-bella under the dissecting microscope, and look for females
incubating embryos (Figure 11). If the embryos appear white, set the female
and her eggs aside for establishing a separate stock colony. If the embryos
appear grey-green, place the female and her young in a separate stender dish.
Carefully tease open the tube and allow the trochophore to swim free of the
tube. Remove the female and the tube. If 200 larvae or more are required,
it will be necessary to obtain larvae from more than one female. If so,
pipette all larvae into a common container, then distribute the larvae to the
experimental chambers.
96-Hour Experiments with Juvenile or Adult Capitella
Use either petri dishes or.500 ml Erlenmeyer flasks as experimental con-
tainers. Place a single worm i*n a petri dish containing 25 ml of toxicant or
an Erlenmeyer flask containing 100 ml of toxicant. Cover the container. Use
20 specimens per concentration and a minimum of five concentrations of toxi-
cant plus control. Examine each specimen daily for death, and discard all
specimens after 96 hours.
Death in Cccpitella—Dead Capitella may be difficult to distinguish from
moribund ones. Death is defined as a lack of movement in response to gentle
poking. Dying Capi-tella may fragment into two pieces. In this instance use
the anterior end to determine death. A dead Capitella generally appears
whiter than a living one and also slightly enlarged. It is usually lying on
the bottom of the container free from any mucoid tube.
40
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Long-Term Experiments with Juveniles (28 days)
Since the life history is short in Cap-Ltella, it is necessary to use
younger worms for conducting a 28-day experiment. Separate out the appropri-
ate aged specimens from white enamal pans. Examine each specimen under the
dissecting microscope for injuries or females with developing egg masses.
Place the selected specimens in containers as outlined for the 96-hour toxi-
city test. It will be necessary to feed the specimens but only at the
initiation of the experiment. Soak dried Enteromorpha in a petri dish con-
taining seawater. Knead the algae so that it is well soaked. Use about O.lg
of dried Enteromorpha sp. (0.2 wet weight) per worm. The Enteromorpha
should be soaked in the same concentration of seawater plus toxicant in which
it is going to be used. Place the appropriate amount of kneaded Enteromorphx
sp. for one worm in a petri dish or 500 ml Erlenmeyer flask with 25 ml or
100 ml of toxicant, respectively. Examine for deaths daily. The solution
should be renewed every four days. Sublethal behavioral modifications in
Capitella include the inability to feed or construct a mucoid tube. A
healthy Capi-tella typically builds its tube within the Enteromorpha sp. on
the bottom of the conainer; a stressed Capitella will construct its tube, if
indeed it does, along the side of the dish at the air-water interface. Dis-
card all specimens at the completion of the experiment.
96-Hour Experiments with Trochophore Larvae
Place ten trochophore larvae in each stender dish filled with 15 ml of
the appropriate toxicant. Use ten dishes per concentration and a minimum of
five concentrations plus control. Add 1.0 ml of a ground Enteromorpha sp.-
seawater mixture of 1:100 by weight to each stender dish. The Enteromorpha
sp. is required as food for the larvae and also to facilitate settlement and
metamorphosis. Examine each dish daily and record the number of living
specimens present. Frequently, a dead trochophore will have disintegrated
within the 96-hour period and therefore cannot be seen.
Long-Term Experiments through FI Generation
Since this species possesses a short life history and abnormal larvae
(Figure 16) can be induced in \the presence of a toxicant, Capitella is a
convenient species with which to conduct a whole life cycle toxicity test.
The procedures outline a static test; if periodic water changes are to be
made, care must be exercised not to disturb females incubating eggs in their
tubes.
Separate out trochophore larvae as outlined above. A total of 100 lar-
vae are needed per concentration. Pipette 25 larvae into a 3.78 A jar that
contains 2,500 ml of seawater and toxicant. Add 0.1 g ground Enteromorphz
to the jar and feed once a week making certain that most of the food had
been eaten the previous week. Do not feed if most of the food is present.
Use four jars per concentration and a minimum of five concentrations plus
controls. The 96-hour and 28-day toxicity tests should be conducted prior
to this reproductive test to determine the toxicant concentrations. Examine
41
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the worms after 15 days for females with incubating embryos and every two to
three days thereafter until the conclusion of the experiment (about 40 days).
To examine the reproductive state of the worms, decant most of the water into
another glass container, remove all the material from the bottom of the jar
and place it into one or more petri dishes, and examine this material under
the dissecting microscope for presence of females with developing embryos in
their tubes. Remove these females and young and place in a separate dish and
count the eggs. Replace all males and non-reproducing females in the jar
with the appropriate toxicant concentration.
The data obtained from such an experiment furnish information to relate
the concentration of the toxicant to survival, development of egg masses,
reproduction, number of embryo produced, and the number of abnormal larvae
produced.
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BIBLIOGRAPHY
American Public Health Association, 1976, Bioassay procedures for marine
polychaete annelids (Tentative). Standard Methods for the Examination
of Water and Waste Water. Amer. Public Health Assoc., Amer. Water
Works Assoc., and Water Pollut. Control Fed., 14th Edition, pp. 785-793.
Barnard, J. L. 1970. Benthic ecology of Bahia de San Quintin, Baja
California. Smith. Cont. Zool. No. 44, 60 pp.
Bellan, G. , D. J. Reish, and J. P. Foret. 1971. Action toxigue d'un
detergent sur le cycle de developpement de la polychete Capitella
occp-ltata (Fab.). Comptes Redues Acad. Sci., Paris. 272:2476-2479.
Bellan, G., D. J. Reish, and J. P. Foret. 1972. The sublethal effects of
a detergent on the reproduction, development, and settlement in the
polychaetous annelid Cap-itella capitata. Marine Biol. 14, 183-188.
Claparede, E. and E. Metschnikoff. 1869. Beitage zur Kenntnis der Entwick-
lungsgeschichte der Chaetopoden. Zeitschr. Wiss. Zool. 29:163-205.
Day, J. H. 1934. The development of Capite Hides giardi Mesnil. Dove Mar.
Lab., Cullercoats. Ser. 3, 4:31-37.
Eisig, H. D. 1887. Die Capitelliden des Golfes von Neapel. Fauna und
Flora des Golfes von Neapel. 16:1-906
Eisig, H. 1898. Zur Entwickelungsgeschichte der Capitelliden. Zool. Stat.
Neapel, Mitt. 13:1-292.
Foret, J. P. 1972. Etude des effets along terme de quelques detergents
(issus de la petroleochimie) sur le sequence du. developpement de deux
especes de polychetes sedentaires: Seolelepis fuliginosa (Claparede)
et Capitella capitata. (Fabricius). .Thesis, Univ. Marseille-Luminy.
125 p.
Foret, J. P. 1975. E'tude des effects a long terme de quelques detergents
sur la sequence du developpement de la polychaete sedentaire Capitella
oagi-tata (Fabricius). Tethys. 6:751-778.
Foret-Montardo, P. 1970. ^tude de 1'action des produits de base entrant
dans la composition des detergents issus de la petrolechimie vis-a-vis
de quelques invertebres benthiques marins. Tethys. 2:567-614.
43
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Grassle, J. G. , and J. P. Grassle. 1974. Opportunistic life histories and
genetic systems in marine benthic polychaetes. Jour. Marine Res.
32:253-284.
Grassle, J. F., and J. P. Grassle. 1976. Sibling species in the marine
pollution indicator Cccpitella capitata (Polychaeta). Science.
192:567-579.
Grassle, J. F., and J. P. Grassle. 1977. Temporal adaptations in sibling
species of Capi-bella. In: Ecology of Marine Benthos, B. C. Coull, ed.
Belle W. Baruch Library in Marine Science, Univ. of So. Carolina Press.
6:177-189.
Guerin, J. P. 1971. Utilisation de nourritures artificielles pour 1'elevage
des jeunes stades d'invertebres benthiques. Tethys. 2:557-566.
Hofker, J. 1930. Faunistische Beobachtungen in der Zuidersee wahrend der
Trockenlegung. Zeitschr. Morph. Okol. Tiere. 18:199-215.
Leschke, M. 1903. Beitrage zur kenntis der pelageschen Polychaetenlarven
der Kieler Fohrde. Wiss. Meeresunters. N.F. Abt. Kiel. 7:113-134.
Oshida, P. A., and D. J. Reish. 1974. The effect of various water tempera-
tures on' the survival and reproduction in polychaetous annelids:
Preliminary report. In: Marine Studies in San Pedro Bay. Part 3.
D. F. Soule and M. Oguri, eds. Allan Hancock Foundation, Univ. of So.
Calif., pp. 63-77.
Pearson, T. H., and R. Rosenberg. 1978. Macrobenthic succession in relation
to organic enrichment and pollution of the marine environment. Oceangr.
Mar. Biol. Ann. Rev., 16:229-311.
Petrich, S. M., and D. J. Reish. 1979. Effects of aluminum and nickel on
survival and reproduction in polychaetous annelids. Bull. Environ.
Contain. Toxicol. 23:698-702.
Rasmussen, E. 1956. Faunistic and biological notes on marine invertebrates
III. The reproduction and larval development of some polychaetes from
Isefjord> with some faunistic notes. K. Danke Vidensk. Selsk. Biol.
Medd. 23:1-84.
Reish, D. J. 1961. Tftie use of the sediment bottle collector for monitoring
polluted marine waters. Calif. Fish and Game. 47:261-272.
Reish, D. J. 1970. The effects of varying concentrations of nutrients,
chlorinity, and dissolved oxygen on polychaetous annelids. Water Res.
4:721-735.
Reish, D. J. 1974 (1976). The establishment of laboratory colonies of poly-
chaetous annelids. Thalassia Jugoslavia. 10:181-195.
44
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Reish, D. J. 1977. The role of life history studies in polychaete system-
atics. In: Essays on polychaetous annelids in memory of Dr. Olga
Hartman, D. J. Reish and K. Fauchald, eds. Allan Hancock Foundation,
Univ. of So. Calif, pp. 461-476.
Reish, D. J. 1978. The effects of heavy metals on polychaetous annelids.
Rev. Int. Oceanogr. Med. 49:99-104.
Reish, D. J., and J. L. Barnard. 1960. Field toxicity tests in marine
waters utilizing the polychaetous annelid Capitella capitata (Fabricius)
Pacific Nat. (21):l-8.
Reish, D. J., and T. V. Gerlinger. 1977. Toxicity of formulated mine tail-
ings on marine polychaeta. Marine Biological Consultants, Inc.,
Costa Mesa, Calif. 102 pp.
Reish, D. J., J. M. Martin, F. M. Piltz, and J. Q. Word. 1976. The effect
of heavy metals on laboratory populations of two species of polychaetous
annelids with comparisons to the water quality conditions and standards
in southern California marine waters. Water Res. 10:299-302.
Reish, D. J., C. E. Pesch, J. H. Gentile, G. Bellan, and D. Bellan-Santini.
1978. Interlaboratory calibration experiment using the polychaetous
annelid Capitella capitata. Marine Environmental Res. 1:109-118.
Reish, D. J., and T. L. Richards. 1966. A culture method for maintaining
large populations of polychaetous annelids in the laboratory. Turtox
News. 44:16-17.
Thorson, G. 1946. Reproduction and larval development of Danish marine
bottom invertebrates with special reference to the planktonic larvae in
the sound (0resund). Komm. Danmarks Fis.-Havundergelse. Ser. Plankton,
Meddel. 4:1-523.
Stora, G. 1972. Contribution a 1*etude de la notion de concentration
lethale limite moyenne (CLso) appliquie a des invertebres marins 1.
Etude methodologique. Tethys. 4:597-644.
Tenore, K. R. 1977. Growth of Capitella capitata cultured on various levels
of detritus derived from different sources. Limn. Oceanogr. 22:936-941.
Warren, L. 1976. Acute toxicity of inorganic mercury to Capitella. Mar.
Pollut. Bull. 7:69-70.
45
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Figure 9. Capitella aapitata, male, dorsal view of anterior end. Note the
genital hooks in setigerous segments 8 and 9.
46
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Figure 10. C. oapitata, female, dorsal view of anterior end.
47
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00
Figure 11. C. oapitata, female, incubating eggs, lateral view.
fertilized eggs lining the inside of the tube.
Note the mucoid tube with the
-------
Figure 12. C. aapi,tata, trochophore stage.
49
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Figure 13. C. capitata, metatrochophore stage.
50
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Figure 14. C. capitata, juvenile.
51
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Figure 15. Gallon (3.78 1) jar aquarium system for rearing C. oapi-tata.
The colony is located within the mass of food at the bottom of
the jar. Air is supplied by an aquarium stone connected to an
air compressor.
52
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Figure 16. C. cap€tata, bifurcated abnormal larva induced by sublethal
amounts of a heavy metal.
53
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APPENDIX C
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELID:
Ctenodx"ilus serratus
CULTURE TECHNIQUES
Equipment and Supplies
Since Ctenodrilus serratus is a minute polychaete, the equipment,
supplies, and space required are minimal. Approximately one irr of shelf or
table surface is required to rear 3,000 to 5,000 or more specimens per month.
A constant temperature room set at 19±1°C, provides an ideal environment to
culture this worm; however, CtenodvLlus can reproduce at temperatures as high
as 25-26°C or as low as 15°C. Because of the small size of this worm, it is
advantageous to use filtered seawater to minimize the introduction of micro-
organisms that may compete for food. The tolerance of Ctenodrilus to reduced
salinities is unknown; this population has been cultured in normal salinity
seawater (35 °/oo). A static system is used to culture this species. Less
than 100 liters of seawater are required per month to yield a monthly popu-
lation of 5,000 specimens. A central compressed air system is the most
convenient source of air for Ctenodrilus, although a population of this size
could be supplied with one air pump. Ctenodrilus are cultured in 3.78 H
(1 gal) jars. Additional supplies needed include 12 cm diameter plastic or
glass covers for the jars, forceps, fine brushes, petri dishes, pasteur
pipettes, plastic tubing, air stones, graded screens, and a blender. Equip-
ment includes a drying oven and dissecting and compound microscopes. Dried
green alga, EntevomoTpha, is used as food.
Collecting Techniques and Life History
This species is a minute polychaete that is widespread throughout the
temperate region of the world. It is difficult to observe in the field be-
cause of its small size. The most convenient way to collect Ctenodfilus is
to bring clumps of fouling organisms from boat floats or pilings into the
laboratory. Examine a mussel under the dissecting microscope, and look for
a small dark purple worm (Figure 17). This species lacks parapodia and
moves over the surface in an earthworm-like fashion. Remove the worm with a
pasteur pipette, and place it in a petri dish containing seawater.
Place a worm on a slide with a cover slip and examine under a compound
microscope. The most conspicuous feature of the worm is its characteristic
seta. The seta (Figure 18) is simple, and each is distally expanded and
54
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provided with four to six serrations along one side. The number of segments
varies, but generally there are about 6-11 setigerous segments. Number of seta
is usually three to four per side of a segment. It has been reported that
Ctenodrilus reproduces in a variety of ways: transverse fission, protandric
hermaphrodism, and internal gestation. A sexual transverse fission is the
most common method of reproduction. Prior to reproduction by transverse
fission, the dorso-anterior margin of each segment bearing setae becomes
enlarged (Figure 19). This becomes the future prostomium. Growth of the
future prostomium continues, and eventually each segment separates to form a
new individual. A complete life cycle takes approximately 14-21 days.
Techniques of Handling Ctenodrilus
Since this species has such a short life history, it is unnecessary to
use special techniques with the different stages. Stock colonies can be
maintained at 19°C with a minimum of time spent. Place several worms in a
petri dish with some food. Enteromorp'ha sp. , a green macroalga, has been
found to be the best food'to use to culture this species. The Enteromorpha
sp. is collected from estuarine areas, washed repeatedly in seawater, and
then allowed to air-dry. EnteTomovp'ha can then be stored indefinitely in
plastic bags. The food is prepared by drying Enteromorpha sp. in an oven at
35°C for about 15 minutes. This crisp Enteromorpha sp. is then placed in a
blender and chopped into fine powder. The powder is shaken through a series
of fine screens to a size of less than 0.06 mm. This powdered food can also
be stored for an indefinite period. In addition to petri dishes, several
worms can be cultured in a single 3.78 £ jar. If several jars are being
maintained, the Enteromovpha sp. powder can be weighed out and mixed with
seawater at the amount of 350 mg per 25 ml of seawater. Each jar is fed at
the rate of above one pasteur pipette drop of this mixture per worm per two
weeks. Make a fresh mixture for each feeding. The specimens can be main-
tained in a jar for about one month. If each 3.78 £ is established with
25-50 worms, a potential yield of 500 to 1,500 specimens may be realized from
each jar per month.
Problems in Culturing—
No special problems have been encountered in culturing Ctenodrilus. It
is important to establish new stock jars on a monthly basis to minimize the
growth of bacteria, protozoans, and copepods. By the use of microporous-
filtered seawater, the buildup of microorganisms is minimized. There are no
known enemies of CtenodrHus.
TOXICITY TEST PROCEDURES
Equipment and Supplies
In addition to the equipment and supplies listed above, 50 mm petri
dishes and a millipore filter system are required.
55
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96-Hour Experiments
Decant most of the water from a stock Ctenodrilus jar. Pour the remain-
ing water into one or more petri dishes. Examine under a dissecting
microscope. Because Ctenodrilus is dark purple use a white background to
facilitate recognition of the various stages. Use only those specimens that
are short (Figure 17) and are not undergoing the early stages of formation
of reproductive buds (Figure 19). The early stages of transverse fission are
recognized by the formation of the future prostomia. Transfer all non-
reproductive specimens with a pasteur pipette into a separate petri dish.
Examine all specimens for injuries and again for early signs of reproduction.
Discard any specimens not suitable for the toxicity test. Small plastic
petri dishes measuring 50 mm in diameter are the most convenient experimental
container, although stender dishes have been used with equal success. Add
20 ml of the appropriate concentration of toxicant to the experimental con-
tainers. Place five specimens per container, and use ten dishes per
concentration to give a total of 50 worms per concentration. Examine daily
and record the deaths. Discard all worms at the end of the experiment.
Death in Ctenodrilus—
Because of its minute size, death in Ctenodrilus is sometimes difficult
to ascertain. A healthy Ctenodrilus is usually moving; therefore, the condi-
tion of these worms is easily distinguished. However, a moribund Ctenodrilus
usually does not move, and it is necessary to observe such a specimen for a
few moments. A dead Ctenodrilus usually appears swollen and may be stuck to
the bottom of the container. It is possible for Ctenodrilus to be completely
decomposed within a 96-hour period, so it is necessary to count living speci-
mens. Be sure to check the water-air interface since Ctenodrilus has the
tendency to crawl to the surface of the water.
Long-term Experiments through Reproduction
Collect the specimens and set up as outlined for the 96-hour toxicity
test. Add five pasteur pipette drops of the Enteromorpha sp. powder-seawater
mixture to each experimental container. Solutions should be renewed every
four days, but because of the small size of Ctenodrilus the number of speci-
mens within each container should be counted before and after each change.
At the end of the experimental period, usually 28 days, count the number of
specimens in each container and record. In addition, note the number of
specimens undergoing bud formation (Figure 19). This latter observation may
be of value in the interpretation of data. Discard all specimens at comple-
tion of the experiment.
It is possible to combine the 96-hour and reproductive experiment into
one experiment. In this case set up the experiment as described for the 96-
hour one. After examination for deaths at 96 hours, add food to each
container in the concentration as noted above and continue the test.
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BIBLIOGRAPHY
American Public Health Association, 1976. Bioassay procedures for marine
polychaete annelids (Tentative). Standard methods for the examination
of water and waste water. Amer. Public Health Assoc., Amer. Water
Works Assoc., and Water Pollut. Control Fed., 14th Edition, pp. 785-793.
Carr, R. S., and D. J. Reish. 1977. The effect of petroleum hydrocarbons
on the survival and life history of polychaetous annelids. In: Fate
and Effects of Petroleum Hydrocarbons in Marine Organisms and
Ecosystems. D. A. Wolfe, ed. Pergamon Press, New York, pp 168-173.
Galvagi, E. 1905. Histolie des Genus Ctenodirilus Clap. Zool. Inst. Wien,
Arb. 15:47-80.
Herland-Meewis, H. 1958. La reproduction asexue chez les annelides. Ann.
Biol. 34:133-166.
Korschelt, E. 1931. Art und Dauer der ungeschlectlichen Fortpflanzung bei
Ctenodrilus. Zool. Anz. Leipzig. 93:227-238.
Deters, N. 1923. Uber das verhalthis der naturlichen zur kunstlichen bei
Ctenodrilus serratus. 0. Schmidt. Zool. Jahrb. (Physiol.) 40:293-350.
Petrich, S. M., and D. J. Reish. 1979. Effect of aluminum on survival and
reproduction in polychaetous annelids. Bull. Environm. Contain. Toxicol.
23:698-702.
Reish, D. J. 1977. The role of life history studies in polychaete system-
atics. In: Essays on polychaetous annelids in memory of Dr. Olga
Hartman, D. J. Reish and K. Fauchald, eds. Allan Hancock Foundation,
Univ. of So. Calif, pp. 461-476.
Reish, D. J. 1978. The effects of heavy metals on polychaetous annelids.
Rev. Int. Oceanogr. Med. 49:99-104.
Reish, D. J., and R. S. Carr. 1978. The effect of heavy metals on the sur-
vival, reproduction, development, and life cycles for two species of
polychaetous annelids. Marine Pollut. Bull. 9:24-27.
Reish, D. J., and T. V. Gerlinger. 1977. Toxicity of formulated mine tail-
ings on marine polychaeta. Marine Biological Consultants, Inc., Costa
Mesa, Calif. 102 pp.
57
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Sokolow, I. 1911. Ueber eine neue Ctenodrilus Art und ibre Vermehrung.
Zeits. Wiss. Zool. 97:546-603. (also in: Zool. Anz. 38:222-226)
58
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Figure 17. Ctenodrilus serratus, dorsal view of entire worm.
59
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Figure 18. C. serratus, simple seta.
60
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Figure 19. C. serratus with five buds forming by transverse fission.
61
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APPENDIX D
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELIDS:
Ophryotrocha diadema and Ophryotrocha puerilis
CULTURE TECHNIQUES
Equipment and Supplies
The equipment and supplies required for a continuous supply of
Ophryotrocha diadema and 0. puer'i'L'is at the rate of 1,000 specimens each per
month are minimal. Approximately one m^ of shelf space is needed for each
species. A constant temperature room is useful for culturing these species
since best results have been obtained at 19±1°C. Specimens will reproduce
successfully vin normal saline water (35 °/oo). Salinity tolerances have not
been investigated. Both species of Ophryotrocha are cultured in a static
system. Approximately 1,000 liters of seawater are required per month to
culture both. A compressed air system is the most convenient method to
supply air to each aquarium, although two air pumps could supply sufficient
air for these cultures. A dissecting microscope with a good light source is
needed to examine the condition of the specimens. Supplies for culturing
include 3.78 $, (1 gal) jar, pasteur pipettes, petri dishes, 12 cm diameter
plastic or glass covers for jars, forceps, plastic tubing, air stones, and a
microporous filter (45 y) system. Food consists of a powder prepared from
commercial fish food such as Biorell.
Life History
The genus Opkryotrocha is a minute polychaete that is widespread through-
out temperate regions of the world. It is difficult to observe in the field
because of its size. The most convenient way to collect Optoyotrocha is to
bring clumps of fouling organisms from boat floats or pilings into the labor-
atory and place them in a pan of seawater. After a period of time, examine
the animals that crawl up along the side of the pan under the microscope, and
look for Ophryotrocha (Figure 20). Remove with an extra fine brush, and
place in a petri dish containing seawater.
Species of the genus OphvyotYodha are very difficult to identify and
require the help of an expert. Historically, all species of Ophicyotvooha.
were referred to as 0. pueyil-is, but the experimental studies of several
European workers have indicated that several species are involved. This
species was originally collected from Los Angeles Harbor. Living specimens
were sent to Dr. Bertil Akesson in Sweden. He identified both 0.
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and an unidentified species from this material that he described as 0.
diadema (Figure 20).
Specimens of 0. puevilis are a subculture of a colony maintained in
Sweden by Akesson. They were brought to Long Beach by a colleague of
Akesson in 1974.
Members of the genus Opkryotrocha are either protandric hermaphrodites
or dioecious. Opkryotroaha diadema belongs to that group of species that are
protandric hermaphrodites with a restrictive number of male anterior segments
and a restrictive number of female posterior segments. Presumably, self fer-
tilization does not occur. The various stages in the life cycle of 0.
diadema are illustrated in Figures 21-24.
Positive identification of the species of Ophryotrocha depends upon a
knowledge of the reproductive biology of the species. Species are separated
on the basis of being either monecious or dioecious, the number of eggs, the
nature of the egg capsule, if present, and the chromosome number.
Techniques of Handling Ophryotrooha
The techniques described herein apply equally to both 0. diadema and 0.
puepilis; however, the life cycle is shorter for 0. diadema. Any plans for
culturing large quantities of these two species for experimentation must
take this difference into account. Since these species have such short life
histories and since the early stages of development take place within the egg
capsule, it is not necessary to use special techniques with the different
stages. Stock colonies should be maintained at 19±1°C. Add 2,500 ml of fil-
tered seawater to a 3.78 H jar. Best results are obtained when the filtered
seawater is additionally filtered through a microporous filter system. Pro-
vide aeration with an air stone connected with plastic tubing to an air
supply. Cover the jar with a plastic lid. Place 20-50 specimens per jar and
feed with about 0.05 g of powdered Biorell about every 10-14 days. Akesson
uses frozen spinach as a food supply for his cultures of Ophryotrocha. The
spinach is fragmented in a mixer and then washed several times with seawater.
The suspension of spinach in seawater is then fed to the cultures. Since
these species reproduce rapidly^ the population reaches its peak in about
three to five weeks, then declines. Each jar should be reestablished every
four to five weeks. The reproductive capacity of OpTwyotToaha can be slowed
by transferring the jars to a colder temperature (either a cold room or a
cold bath). It is possible to delay reestablishment of the jars by as much
as two to three months if the colony is maintained at 15-16°C.
Problems in Culturing—
Most of the difficulty in culturing Ophcyotrooha can be attributed to the
lack of regular care, especially the neglect of reestablishing new cultures
at four to five weeks. Records of the date of establishment of each jar
should be kept to minimize this potential problem. A fungal growth appears
whenever the colony is overfed. This fungal growth can lead to a poor yield
or loss of the colony within the jar. If the fungal growth does appear, the
colony should be reestablished. It is especially advantageous to use micro-
porous-filtered seawater in all jars to minimize growth of microorganisms.
63
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Enemies — Ophryotrocha are generally parasitized with gregarines
(Protozoa : Sporozoa) . While these internal parasites are not lethal to the
host, they lead eventually to a decrease in vitality. The buildup of the
gregarines generally occurs in jars that have been established for five weeks
or longer. Regular care will minimize the problem but not eliminate it.
TOXICITY TEST PROCEDURES
Equipment and Supplies
Stender dishes are needed to conduct toxicity tests with Ophryotroeha .
96-Hour Experiments
Decant all but 100 ml of seawater from a stock jar. Transfer the remain-
ing water into petri dishes, and place them under the dissecting microscope
for examination and selection of test organisms. Remove only the smaller
sized specimens and place in a separate petri dish. Place five specimens in
a stender dish together with 25 ml of seawater and the toxicant. Use ten
dishes per concentration and a minimum of five concentrations plus controls.
At the end of the 96-hour test period count the number of living specimens
in each container and record. Discard all specimens at the end of the
experiment.
Long-Term Experiment
Follow the identical procedures as outlined for the 96-hour bioassay
experiment. In fact, the same experiment can serve as both the short-term
and long-term experiment. At 96 hours, add to each dish five pasteur pipette
drops of a suspension of a 1 percent solution (by weight) of Biorell powder
in seawater. Solutions should be renewed every four days, but because of
the small size of Ophryotrocha , the number of specimens within the dish
should be counted before and after each change. After 28 days, count the
number of specimens in each jar. A separate count can be made of the number
of eggs or larvae within egg capsules. Discard all specimens at the termin-
ation of the experiment.
Death in
Because of its small size, it is usually difficult or impossible to de-
tect sick or dying specimens. The lack of fecal pellets on the bottom of the
dish in long-term experiments is an indication of either death or distressed
specimens. Dead Ophryotrooha are generally found on the bottom of the dish.
If decomposition has occurred, then only the black jaws can be seen.
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BIBLIOGRAPHY
Since specific identification of Ophryotrocha is difficult at best, all
references to experimental studies in the genus are included in this
.bibliography.
Akesson, B. 1967. On the biology and larval morphology of Ophryotrocha
puerilis Claparede and Metschnikob (Polychaeta). Ophelia. 4:110-119.
o
Akesson, B. 1970. Sexual conditions in a population of the polychaete
Ophryotvooha labron-ica La breca and Bacci from Naples. Ophelia.
7:167-176.
Akesson, L. 1970. Ophryotrocha Idbvonioa as a test animal for the study
of marine pollution. Helgolander Wiss. Meeresunters. 20:293-303.
Akesson, B. 1972. Sex determination in Opkryotrocha Zabronioa (Polychaeta,
Dorvilleidae). Fifth European Marine Biology Symposium, Padova,
pp. 163-171.
o
Akesson, B. 1972. Incipient reproductive isolation between geographic
populations of Ophpyotvocha Idbvon-ioa (Polychaeta, Dorvilleidae).
Zoologica Scripta. 1:207-210.
Akesson, B. 1973. Morphology and life history of Opkryotrocha masculata
sp. n. (Polychaeta, Dorvilleidae). Zool. Scr. 2:141-144.
Akesson, B. 1973. Reproduction and larval morphology of five Ophryotrocha
species (Polychaeta, Dorvilleidae). Zool. Scr. 2:145-155.
Akesson, B. 1975. Reproduction in the genus Ophryotrocha (Polychaeta;
Dorvilleidae). Pubbl. Staz. zool. Napoli. 39 (suppl.):377-398.
Akesson, B. 1975. Bioassay studies with polychaetes of the genus
Opkryotrocha as test animals. In: Sublethal Effect of Toxic Chemicals
on Aquatic Animals. O. J. M. Koeman and J. J. T. W. A. Strik, eds.
Elsevier, pp. 121-155.
Akesson, B. 1976. Morphology and life cycle of Ophryotroaha diadema, a new
species from California. Ophelia. 15:23-35.
Bacci, G. 1951. Efetti di ripetute amputazioni sulle fasi sessuali di
Ophryotrocha puerilis. Boll. Zoll. 18:193-196.
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Bacci, G. 1951. Existence of true males and females in a hermaphrodite
population of OpTwyotrocha pueril-is. Experientia. 7:222.
Bacci, G. 1955. Controllo genetico della inversions sessuale in individui
isolati di Opkpyotrocha. Caryologia (Suppl.) pp. 966-968.
Bacci, G. 1955. La variabilita dei genotipi sessuale negli animali
ermafroditi. Pubbl. Staz. zool. Napoli. 26:110-137.
Bacci, G. 1964. Equililrio genetico dei sessi e variabilita sessuale in
Opkryotrocha pueril-is. Boll. Zool. 31:1093-1097.
Bacci, G. 1965. Sex determination and genie balance of Ophryotrocha
puerilis, a hermaphrodite polychaete worm. Nature, London.
297:1208-1209.
Bacci, G., and O. Bortesi. 1960. Conferma della variabilita dei gentipi
sessuali nel polichaet ermafrodita Ophryotrocha puerilis siberti.
N.C. Accad. Lincei. 28:92-94.
Bacci, G., and O. Bortesi. Existence of multiple sex genotypes in the
hermaphrodite polychaete worm Ophryotroeha puerilis siberti. Nature,
London, 190:838.
Bacci, G., and 0. Bortesi. 1961. Pure males and females from hermaphroditic
strains of Ophryotrocha puevilis. Experientia. 17:229-230.
Banse, K. 1963. Polychaetous annelids from Puget Sound and the San Juan
Archipelago, Washington. Proc. Biol. Soc. Wash. 76:197-208.
Bortesi, O. 1964. Differenziamento citosessuale in ceppi mascolinizzati
e femmenelizzati di Opkpyotrocha puerilis. Boll. Zool. 31:1103-1109.
Bortesi, 0. 1965. Ibridazione fra sottospecie di Opkryotrocha puerilis.
Boll. Zool. 32:1141-1150.
Braem, F. 1894. Zur Entwickhengsgescheate von Opkryotrocha puerilis. Z.
wiss. Zool. 57:187-223.
Braem, F. 1908. Ueber die Aendrung des Geschlechts durch Auszere
Beeinflussung and uber die Regeneration des afterdarms bei Ophryotroeha.
Anat.Anz. 33:19-27.
Brown, G., and K. M. Ahsanullah. 1971. Effect of heavy metals on mortality
and growth. Marine Poll. Bull. 2:182-187.
Carr, R. S., and D. J. Reish. 1977. The effect of petroleum hydrocarbons
on the survival and life history of polychaetous annelids. In: Fate
and Effect of Petroleum Hydrocarbons in Marine Organisms and Ecosystems,
D. A. Wolfe, ed. Pergamon Press, New York, pp. 168-173.
66
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Claparede, E., and E. Mecznikow. 1869. Beitrage zur Erhenntrusder
Entwicklungsgeschichte der Cahetopoden. Z. wiss. Zool. 16:163-205.
Clark, R. B. 1965. Edocrinology and the reproductive biology of poly-
chaetes. Oceanogr. Marine Biol., Annual Reviews 3:211-255.
Dehorne, A. 1910. Le mecanisme de la reduction numerique dans la sperma-
togenese de Ophryotrocha puerilis. Clpd.-Meck. Zool. Anz.
36:209-222.
Dohle, W. 1967. Zur Morphologie und Lebensweise von Ophryotrocha graoilis
Huth 1923 (Polychaeta, Eunicidae). Keiler Meeres; forsch. 23:68-74.
Dusing, H. 1961. Die Umwandlung des Kavapparates in Abhangigkeit von
sexuellen Zustand bei OphvyotTOcha pueril'is Claparede & Metschnikoff
Naturwissenschaften. 48:532-533.
Emanuelsson, H., and 0. Hehy. 1978. Inhibition of putrescene synthesis
blocks development of the polychaete Ophryotroeha labroniea. Proc.
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Gregoire, V., and W. Deton. 1885. Contribution a I1etude de la spermato-
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und Geschlechtsumwandlung von Ophryotrocha puerilis. Zool. Jahbr.,
Jena. allg. Zool. Physiol. 56:389-439.
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La Greca, M., and G. Bacci. 1962. Una nuover specie di Opfcyotrocha delle
coste timeniche. Boll. Zool. 29:13-23.
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mique. Bull. Soc. Zoll. Fr. 79:466-469.
Mviller, H. 1962. liber die Sexualitat des Polychaeten Ophryotrocha puerilis,
ibre determination und ihrer Einfluss aus Drusen - tatgkeit und
Kanapparatenwicklung. Zool. Morph. Okol. tiere. 52:1-32.
Oshida, P. A., and D. J. Reish. 1974. The effect of various water tempera-
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Preliminary report. In: Marine Studies in San Pedro Bay, California,
Part 3. D. F. Soule and M. Oguri, eds. Allan Hancock Foundation, Univ.
of Southern California, pp. 63-77.
Parenti, U. 1960. Citossonaia del genere Ophryotrocha (Annelida,
Polychaeta). Rend Ace. Naz. Lincei. 28:386-389.
Parenti, U. 1960. Self fertilization in Ophryotrocha labroniea.
Experimentia. 16:413-414.
Parenti, u. 1961. Ophyrotrocha pueri-Hs siberti,, 0. hartmanni. ed. 0. bacci
nelle aeque de Roscoff. Cab. Biol. Mar. 24:437-445.
Parenti, U. 1961. Al differenzeamento citosessuale in Ophryotroeha
Idbronica. Monit. Zool. Stat. 69:106-119.
Parenti, U. 1962. Variabilita sessuale di una nuova sottospecie di
Optirytrooha hartmanni del mediterraneo. Rend. Ace. Naz. Lincei.
33:78-83.
Parenti, U. 1964. Inversione sessuale limitata a due metameric in
Ophpyotrocha graeilis. Boll. Zool. 31:25-31.
Parenti, U. 1964. Opkryotrocha gracilis ed. 0. baoo-i nelle acque. di
Plymouth: nuova deserizione e notizie biologiche. Boll. Zool.
31:33-39.
Pfannenstiel, H-D. 1972. Eine neue Ophryotrocha-Art (Polychaeta,Eunicidae)
aus tapan. Helgolander wiss. Meeresunters. 23:117-124.
Pfannenstiel, H-D. 1977. 1st der polychaet Ophryotrocha labronica ein
proterandrischer Hermaphrodet. Marine Biol. 38:169-178.
Pfannenstiel, H-D. 1977. Experimental analyses of the "paar kultuneffekt"
in the protandric polychaete, Opkryotroeha puerilis. Jour. Exper. Mar.
Biol. Ecol. 28:31-40.
Pfannestiel, H-D. 1977. Endokrinologsche und genetische Untersuchungen an
einer proterandrichen Population des Polychaeten Opkryotrocha labvonica.
Marine Biol. 39:319-329.
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Reish, D. J., and R. S. Carr. 1978. The effect of heavy metals on the
reproduction, development, and life history for two species of poly-
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Artem-ia salina and Ophryotrocha labvonica to copper sulfate. Marine
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labvonioa. Atti. Accad. Naz. Lincei. 32:703-706.
Zunarelli, R. 1962. Al' differenziamento citosessuale di tre specie di
Ophryotrocha. Boll. Zool. 29:417-423.
Zunarelli-Vandini, R. 1967. Azioni reciproche sulle gonadi in coppie
oseospecifiche ed eterospecifiche di Ophryotrocha puevLH-s siberti
ed Opkryotrochalabronica. Archo. Zool. Ital. 52:177-192.
69
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Figure 20. Ophryotrocha diadema, dorsal view of entire worm (after
Akesson, 1976}.
70
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Q
Figure 21. 0. diadema, egg mass (after Akesson, 1976)
71
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Figure 22. 0. diadema, larva from egg mass (after Akesson, 1976)
72
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Figure 23. 0. diadema, larva from egg mass (after Akesson, 1976)
73
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Figure 24. 0. diadema, released larva (after Akesson, 1976)
74
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APPENDIX E
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELID:
Janua (Dexiospira) brasiliensis
CULTURE TECHNIQUES
Equipment and Supplies
The equipment required for a continuous supply of Dexiospira consists of
a dissecting and compound microscope, a good microscope light, and a blender.
Approximately two m^ of shelf space is required to culture about 1,000
specimens. A constant temperature laboratory maintained at 19±1°C will in-
sure maximum results since this species is especially sensitive to tempera-
tures above 21° or 22°C.
Natural, filtered seawater at a salinity of 35.5 °/00 will yield the
maximum number of offspring since this species is sensitive to salinities be-
low about 32 °/00. Approximately 100 liters of seawater are required per
month to culture a population of 1,000 to 2,000 specimens. A central com-
pressed air system is the most convenient source of air for the culture. It
is advantageous to filter all seawater to be used in culturing Dexiospira
through a microporous filter (45 U) to minimize the growth of microorganisms.
Additional supplies required for culturing Dexiospira include 3.78 £ (1 gal)
glass jars, petri dishes, pasteur pipettes, beakers, forceps, microscope
slides, plastic string, air stones, plastic tubing, and scalpel. Since
Dexiosp-Lra requires living food (Dunaliella tertiolecta), the techniques,
equipment, and supplies required to culture this phytoplanktonic organism are
given as a separate section within Appendix E.
Collecting Techniques and Life History
Dexiospira occurs in abundance on floating boat docks in southern
California. This species grows on the surface of docks as well as fouling
organisms; it is easily collected from the surface of mussels (Mybilus
edulis) or sea lettuce (Ulva lactuca)- The calcareous tube of this species
is white, coiled dextrally (counter-clockwise), and measures one to two mm
in diameter (Figure 25). Sinistral tubes of spirorbids and other dextrally
coiled species may be found in association with Dex-iospiva so careful identi-
fication is required.
While Dexiospiva. is widespread only in the warmer waters of the world,
the techniques described herein could apply to other spirorbid polychaetes.
Identification of the Spirorbinae is extremely difficult. The key
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characteristics employed in identification are as follows: orientation and
morphology of the tube, method of brood protection, number of thoracic seg-
ments, and types of setae and uncini. As the setation is very similar in
many species, it may be necessary to send specimens' to an authority for
species confirmation. Consult the papers by Knight-Jones and others (1959,
1975, 1977) for keys to the species of spirorbids.
The tubes of Dexiospiva are easily removed from the fronds of Ulva with
a scalpel. If they are found on mussel valves, it is better to leave them
on, since many of the tubes would be crushed on removal. It is best to open
the mussel and remove all of the soft tissue. This will prevent possible
future fouling of the water by the mussel. The suspension of glass jars or
wooden blocks from a boat dock for about a month is a convenient method of
collection.
Animals removed from Ulva should be placed in glass petri dishes using
a pasteur pipette with an adequate opening. The cleaned mussel valves
should be placed in 3.78 £ (1 gal) jars. If the wooden blocks are small
enough, these also may be placed in small aquaria.
Dexiosp-ira is hermaphroditic. Although these animals are capable of
self-fertilization, the viability of the embryos seems to decrease with re-
peated self-fertilization. Therefore, it is advantageous to culture them
in groups rather than separately. After fertilization, the embryos are
transferred (method unknown) to the operculum, which is modified as a brood
chamber (Figure 26) . The brooding period lasts from six to eight days and
is followed by the discharge of the pelagic larvae. The larvae settle and
begin tube formation within one hour following their release. Sexual matur->
ity is attained in 30 days.
Techniques of Handling
Place a petri dish containing 50 to 100 tubes of Dexiospira on the bot-*
torn of a 3.78 & jar provided with aeration and containing 2,500 ml of
millipore-filtered seawater. Suspend microscope slides (2.54 x 7.62 cm) in
the water in order to provide a surface for larval settlement. In order to
facilitate larval settlement prepare the slides as follows: place fresh
Ulva in a blender and grind the alga into small pieces. Pour the material
into a beaker, and allow the larger pieces to settle. Dip a clean microscope
slide into the supernatant fluid for a moment and remove. It is not neces-
sary to dry the slide; it can now be suspended in the jar with a plastic
string. Since field specimens are reproductively active year round, only
two to three days are required for adequate larval settlement on the glass
slides. Transfer the slides to a clean aquarium provided with aerated seat!
water and Dunaliella as food. A new set of slides can be placed in the
original aquarium for additional settlement. This procedure can continue
until the field collected specimens are no longer in the reproductive state,
The green phytomastigophoran Dunaliella has been found to be the best
food source. Approximately 30 to 40 ml of DunaHella are added to each
aquarium twice a week to maintain the culture. The seawater should be
changed every two to three weeks.
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If the mussel valves with tubes are not properly washed of extraneous
material, the culture will be infested with a myriad of unnecessary
organisms. These are not necessarily harmful to Dexiospira, but they will
cause an increase in the amount of Dunal-iella required to maintain the
culture. This contamination can be reduced by changing the water every two
to three weeks and by using microporous-filtered seawater. Larvae of
Dex-iospira will often settle and metamorphose on the surface tension when
cultured in petri dishes. They can be dropped to the bottom by touching
them with forceps. Larvae will settle on the inside of the aquarium as well
as on the glass slides. Allow these specimens to mature, and then they can
be removed and utilized in establishing additional colonies. There are no
known enemies of Dexiospira.
TOXICITY TEST PROCEDURES
Only small plastic petri dishes (15x60 mm) are required in addition to
the equipment and supplies specified above.
96-Hour Experiments
Remove specimens from either the sides of the stock aquaria or from the
suspended glass slides. The specimen can be removed with the flanged tipped
pipette by scraping the sides with the tip while producing suction with the
rubber bulb. There is only a minimal amount of damage to the animal when
this technique is employed. Examine each specimen under the dissecting
microscope, and discard any injured specimens. Place five specimens in each
small petri dish containing 15 ml of seawater and toxicant. Use ten dishes
per concentration with a minimum of five concentrations plus control. These
worms will not require food during the 96-hour experimental period. Count
the number of living worms at 96 hours, and discard all specimens.
Long-Term Experiment Involving Reproduction
Establish the toxicity test as outlined under the 96-hour experiment.
Add 1.0 ml of Dunaliella to each container every other day, and change the
medium in the petri dish once a week during the experimental period. If this
procedure is not followed, then a population of microorganisms, in addition
to DunaHella, will build up rapidly. Allow the experiment to run 28 to 35
days to allow sufficient time for reproduction to occur. Count the number of
living specimens in each container. Mount each specimen on a slide, examine
under a compound microscope, and count the number of embryos within the oper-
culum. Discard all specimens at the completion of the experiment. It is
possible to combine both the 96-hour LCso experiment and the reproductive
experiment. Examine for deaths at 96-hours, record, feed with Dimaliella,
and allow the experiment to continue for the 28-35 days.
Death in Dexiospira—
An animal may be considered dead when there is no sign of movement or
when the body is decomposed. It may be necessary to open the tube to deter-
mine the animal's condition. The tube can be opened by tapping the tube with
a pair of fine forceps.
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LABORATORY CULTURE OF Dunaliella tevtiolecta
Medium
Seawater: Use about 30-32 °/oo salinity seawater (90% seawater and 10%
double distilled water). Filter through a microporous-filter system (45 y).
For 50 ml quantities, mix complete medium (below) and autoclave at 15 psi
for 15 minutes. For larger quantities, autoclave phosphate separately, and
then add to the medium. If a serious precipitation occurs, it may be neces-
sary to filter sterilized medium prior to use.
Major nutrients: Nitrate— 5 g KN03 made to 1,000 ml with H20. This
stock solution may be stored in a refrigerator for up to four months. Add
1.0 ml stock solution to one liter distilled water to make a final solution;
use ten ml of this solution per liter of culture medium. Do not store final
solution.
Phosphate—0.68 g KH2P04 made to 1,000 ml with H20. This solution may be
stored in a refrigerator for up to four months. Add 1.0 ml stock solution
to one liter distilled water to make a final solution; use ten ml of this
solution per liter of culture medium. Do not store final solution.
Minor trace metals—Stock metals—30 mg ZnS04«H20, plus 25 mg CuS04'5H20,
plus 20 mg CuS04'7H20 dissolve 5.0 g FeCl3-6H20 and 2.0 g MnS04'H20 in 1,000
ml distilled water (ignore slight precipitate).
Sodium Molybdate—25 mg Na2Mo04*H20 in 1,000 ml of distilled water.
Sodium Ethylenediaminetetraacetate—Dissolve 50 g of Na2E.D.T.A.-2H20 in
1,000 ml of distilled water.
Metal Mixture—Add 100 ml of the Na2E.D.T.A.-2H20 solution and ten ml each to
the three minor trace metal solutions listed above to about 800 ml of dis-
tilled water. Adjust the pH to 7.5 with dilute NaOH solution and make up to
1,000 ml with distilled water. Add 1.0 ml to each liter of culture medium.
This solution may be stored for up to four months in a refrigerator.
Vitamins: Vitamin B]_2—Dissolve 10.0 mg crystalline vitamin Bi2 in 100ml
of distilled water. Store in deep freeze until use. Immediately prior to
use, thaw solution, remove 1.0 ml and refreeze stock solution. Dilute the
one ml of vitamin B]_2 solution with 99 ml of distilled water. Add 1.0 ml of
diluted solution to each liter of seawater and throw away the remainder.
Biotin—Dissolve 10.0 mg biotin in 100 ml of distilled water and freeze.
Immediately prior to use, thaw the solution, remove 1.0 ml and refreeze the ,
stock solution. Dilute the 1.0 ml with 99 ml distilled water. Add 1.0 ml of
diluted solution to each liter of seawater and throw away the remainder.
Thiamine hydrochloride—Dissolve 100 mg of thiamine hydrochloride in 100 ml
of distilled water and freeze. Immediately prior to use thaw, remove 1.0 ml,
and refreeze the stock solution. Dilute the 1.0 ml with 99 ml of distilled
water. Add 1.0 ml of diluted solution to each liter of seawater and throw
away the remainder.
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Culture
Place 5 to 15 ml of Dunal'lel'l.a from a previous culture in a 500 ml
Erlenmeyer flask containing 500 ml of autoclaved medium. The amount of
Dunalie'Lla. depends upon its concentration in the previous culture. Place
stoppered Erlenmeyer flasks under grow-lux fluorescent lights. Depending
upon the amount of Dunaliella required, the alga can be exposed to continu-
ous light or to a light-dark cycle, which is maintained with an electrical
timer. The growth rate can be slowed, if desired, by placing the cultures
in a cold bath (17°C) with the lights placed overhead.
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BIBLIOGRAPHY
Bailey, J. H. 1969. Methods of brood protection as a basis for the reclass-
ification of the Spirorbinae (Serpulidae). Zool. Jour. Linnean Soc.
48:387-407.
deSilva, P.H.D.H. 1962. Experiments on choice of substrate by Spfoorbis
larvae. Jour. Exper. Biol. 39:483-490.
deSilva, P.H.D.H. 1967. Studies on the biology of Spirorbinae. Jour.
Zool. 152:269-279.
Gee, J. M. 1967. Growth and breeding of Spirorbis rupestris (Polychaeta:
Serpulidae). Jour. Zool. 152:235-244.
Gee, J. M., and G. B. Williams. 1965. Self and cross fertilization in
Spfaorbis borealis and S. pagensteohevi.. Jour. Mar. Biol. Assoc.
U.K. 45:275-285.
Hoglund, L. B. 1951. Notes on the Morphology and Biology of Some Spirorbid
Larvae. Zool. Bidr. Uppsala. 29:261-276.
Knight-Jones, P, and E. W. Knight-Jones. 1977. Taxonomy and ecology of the
British Spirorbinae (Polychaeta). Jour. Mar. Biol. Assoc. U.K.
57:453-499.
Knight-Jones, P., E. W. Knight-Jones, and T. Kawakara. 1975. A review of
the genus Jantia, including Dexiospira (Polychaeta:Spirorbinae). Zool.
Jour. Linnean Soc. 56:91-129.
Williams, G. B. 1964. "Effects of Extracts of Fuoas serratus in promoting
the settlement of the larvae of Spirorbis borealis". Jour. Mar. Biol.
Assoc. U.K. 44:397-414.
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Figure 25. Janua (Dex-Lospira) brasiliensis, tube.
81
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Figure 26. D. brasiliens-is, dorsal view of entire worm removed from tube.
82
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APPENDIX F
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELIDS:
Polydopa ligni and Boecardia proboscidea
CULTURE TECHNIQUES
Equipment and Supplies
The only equipment required to culture these two spionid polychaetes is
a dissecting and compound microscope. A blender is convenient to mix food.
About four m^ of shelf space is required to culture a large number of both
these species. A constant temperature laboratory set at 19±1°C provides the
ideal environment. For best results, filter all seawater through a micro-
rorous filter (45 y) to remove microorganisms; however, stock colonies do not
necessarily require it. Approximately 150-300 liters of seawater are re-
quired per month to culture these two species. A central compressed air
system is the most satisfactory method of supplying air to the aquaria. A
magnetic stirring device (Hinegardner, 1969) is convenient for culturing the
planktonic stages. Consumable supplies include 3.78 £ (1 gal) jars, petri
dishes (20 x 100 mm), pasteur pipettes, 400 ml plastic tri-pour beakers with
lids, air stones, plastic tubing, and glass tubing.
Collecting Procedures and Life History
Polydova lign-L—
This species of polychaete is found throughout the temperate regions of
the world and may inhabit substrata ranging from silty mud to sand. The worm
is generally found inhabiting tubes made of mud or sand lying within the up-
per few centimeters of sediment. Since this species has a rather wide
salinity tolerance, it may be found anywhere from the open coast to the estu-
arine environment.
Adult Polydova can be collected by use of a 0.5 mm screening box. In
the collection area, several scoops of the top two to five cm of the sub-
stratum are placed in the screening box and are gently shaken through the
screen. The adult worms and their tubes will remain on the screen and can be
washed into a container of seawater. Since many of the adults will remain
inside their tubes during the sifting process, all tubes, left on the screen
should be saved and inspected for worms in the laboratory under the dissect-
ing microscope. Adults measure from 3 to 25 mm in length.
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If no suitable collection area can be found for the adult worms, a
population can be started from the planktonic larvae. Polydora reproduces
seasonally in some locations and almost continuously in other areas (Rice,
1975). Generally, the planktonic larvae can be obtained by use of a shallow
(as near the surface as possible) plankton tow using a fine mesh plankton
net. Larvae obtained in this manner will settle out and metamorphose into
adults within one to two weeks under laboratory conditions. Place the
larvae in a 3.78 £ jar provided with 2,500 ml microporous-filtered seawater,
some fine sand, and an air supply.
Polydora (Figures 27 and 28) belongs to a subgroup of the Family
Spionidae known as the polydorid complex that contains the genera Polydora,
Boocardia, and Pseudopolydora. These worms share the common feature in
which the setae of their fifth setiger are modified into thick heavy spines
of various shapes (Figure 29). This feature is useful in identification of
species. Polydora is distinguished from the other members of the genus by
the presence of the following three structures: 1) a subdistal tooth near
the tip of the heavy spines of the fifth setiger (Figure 29), 2) small
brush-like companion setae lying directly on top of the heavy spines of the
fifth setiger (these can only be seen under a compound microscope) (Figure
29), and 3) a small nuchal antenna on the caruncle just between the palps
(Figure 27). For further data and keys to the species of Polydora see Blake
(1971) and Hartman (1969).
The planktonic larvae of Polydora are very difficult to distinguish
from the other members of the genus. Once the larvae have settled and meta-
morphosed into the adult form, the above characteristics may be used for
identification. For data on larval morphology and occurrence see Hannerz
(1956) and Blake (1969).
Spawning occurs throughout the year under laboratory conditions. The
male releases spermatophores out the anterior opening of his tube. The
spermatophores are picked up by the palps of the female and transported into
her tube where the spermatophore is broken and the sperm released. Fertili-
zation has not been observed, but apparently sperm are stored in the seminal
receptacle with fertilization occurring during formation of the egg capsule.
Early development, through the three-segmented larval stage, takes place in
these egg capsules (Figure 30). Development from the 3 to the 14-segmented
stages takes place in the water mass, then the larvae metamorphose and
settle (Figures 31 and 32). Sexual maturity is reached in 50 to 60 days.
Boocardia proboscidea—
This species has been found from the intertidal and shallow subtidal
waters of British Columbia south to southern California. It constructs sand
tubes in the sediments of bays and estuaries, among clumps and holdfasts of
rocky intertidal algae, within shells of mollusks, and within soft sandstone
rocks. Adult Boccardia are collected conveniently by either screening inter-
tidal sediments through a 0.5 mm screening box or finding populations within
sandstone rocks. Boooardia is recognized by the snout-like prostomium and
by the presence of sooty pigmentation along either side of the prostomium
(Figure 33). Confirmation of this species depends upon the microscopic ex-
amination of the setae of the fifth setigerous segment (Figure 34). Adults
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measure up to 5 mm in length.
Booaardia produces two different types of eggs: one measures 100 u in
diameter and develops into planktotrophic larvae, and the other egg type
measures 150 U in diameter and develops into lecithotrophic larvae. Addi-
tional differences that result from these two egg types are noted below.
In the population of Boocardia that lays the smaller egg, the eggs are
presumably fertilized with sperm stored in the seminal receptacle, similar
to the Polydora. The zygotes are deposited in egg capsules (Figure 35),
which number one per segment, and may total up to 75 per specimen. The num-
ber of zygotes per capsule varies from 4 to 250 with an average of 150.
Each zygote produces a larvae that hatches from the capsule. After five
days, setae appear on three segments (Figure 36). The setae elongate and
the larvae emerge from the capsule after eight days. New capsules are
formed as soon as the larvae from the first laying are released. Larvae re-
main planktonic from day 8 to 29 at which time they possess about 12 to 14
segments. Metamorphosis to the benthic stage was first noted after 29 days
with larvae possessing 15 segments (Figure 37). Sexual maturity is reached
as early as 110 days.
Development in the population of Bocoardia that produces the larger egg
proceeds somewhat differently with regard to timing of the various stages
and the number of young produced. Three setigerous larvae are noted at 96
hours with additional segments appearing more or less daily; a 13-segmented
larvae is ten days old. After 17 days, 13 to 15 segmented larvae hatch from
the capsule. Only one or a few larvae emerge from a single capsule; the re-
maining zygotes served as nurse cells during development of these few larvae.
This form has not yet been reared to sexual maturity in the laboratory.
Techniques of Handling Polydora and Boeeardia
Polydora ligni,
Since the life cycle of this polychaete involves both a benthic and
planktonic stage, it is necessary to employ special techniques in handling
the adults and larvae. Some of the techniques for handling both stages have
been described under collecting techniques above. About 50 to 100 adults
can be maintained in 3.78 £ (1 gal) jars filled with 2,500 ml of filtered
seawater. Aeration is provided with an air stone attached via plastic
tubing to a compressed air system. Adults are fed an Enteromorpha sp. and
Biorell (or Tetramin) food mixture (4:1 ratio). Enteromorpha sp. is col-
lected from estuarine areas and washed repeatedly to remove sediment and
debris. The Enteromorpha sp. is air-dried and stored in plastic bags. The
food mixture for Polydora is prepared by drying the alga in an oven at 35°C
for 15 minutes. Then the crisp Enteromorpha sp. is placed in a blender and
chopped into a fine powder. This powder is shaken through a screen with a
0.05 mm mesh. The Enteromorpha sp. powder is collected on a pan underneath
the screen. This food can be stored for an indefinite length of time.
Biorell or any commercially available fish food can be used as a high
protein supplement along with Enteromorpha sp. The Biorell is prepared by
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chopping the food in a blender and then shaking it through a 0.06 mm screen.
The fine powder is collected from the pan below the finest screen. The food
can be stored indefinitely in this form in a moisture proof container.
For best results, 0.25 g of food mixture should be added to each jar
twice a week, and the water should be changed once a month. Larger numbers
of individuals can be maintained in 18.9 £ aquaria using similar techniques.
A good indication of a healthy culture is the presence of the worm's palps
extending out of the end of its tube in search of food. Also, it is often
possible to directly observe the worms in their tubes by use of a hand lens
when the tubes are constructed on the side of the jar.
Planktonic larvae may periodically appear, usually in large numbers in
the adult culture aquaria. The larvae are small and hard to see, but by
turning off the air supply and directing a strong beam of light through the
container, the swimming movements of the larvae can be observed. The larvae
are strongly attracted to light in their early stages, and they can be con-
centrated and removed from the adult colony by directing a light through the
aquarium near the surface of the water. Within 10 minutes, nearly all of
the larvae will have accumulated at the point nearest the light source and
they can be removed with a pipette. Most field-collected or laboratory-
cultured larvae will not survive to maturity. Generally, less than 10 per-
cent will reach adulthood. For maximum survival, the larvae should be
placed in about 300 ml of microporous-filtered seawater in an appropriate
container, such as a plastic Tri-pour beaker with a perforated cardboard
cover, and provided with about 20 ml of DunaHella as a food source twice a
week. Put about 300 larvae in each container. Then place the containers
onto a magnetic stirring device (Hinegardner, 1969). Within two or three
weeks the larvae will settle on the bottom and metamorphose into the adult
form. The metamorphosed adults can be kept for two or three months in the
same beakers. When it is observed that most of the larvae have meta-
morphosed into adults, their diet should be changed to the Enteromovpha-
biorell mixture and the water in each container changed every two weeks.
Boceardia probosc-idea—
The same techniques used in culturing the adults and larvae of Polydora
ligni are used in culturing Boccard'ia.
Problems in Culturing Polydora and Boooardia—
The primary problem in culturing these adult spionids is overcrowding
and overfeeding. Since these species may be present in large numbers in the
field (10,000 to 50,000 per m2 or more), the tendency is to attempt to main-
tain all collected specimens. This results in overcrowding that in turn will
result in death of some specimens. These deaths will foul the water and
lead to additional deaths. If too much food is added to the culture, the '
worms will begin to show stress as a result of lowered dissolved oxygen and
leave their tubes. When this is observed, the water in the container should
be changed immediately and the worms allowed to recover for one to two days
before feeding again. Another common problem occurs as a result of too much
Biorell in the food mixture. Large amounts of Biorell, if left uneaten in
the culture, will develop a white mold after a few days and may cause the
worms to leave their tubes. Again, this problem can be alleviated by
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changing the water and reducing the amount of Biorell in the food.
Maximum survival of larvae depends largely upon keeping the cultures as
free from contamination as possible. The most common source of contamina-
tion is the presence of an adult culture of protozoans and bacteria. These
organisms do not harm the adults but may be deleterious to the larvae. It
is often helpful to put the larvae through several changes of microporous-
filtered seawater in small petri dishes (use the light attraction technique
described above to concentrate the larvae) before adding the Dunal-iella.
TOXICITY TEST PROCEDURES
Equipment and Supplies
No additional equipment or supplies are required to conduct toxicity
tests with Polydora ligni and Bocoardia proboscidea; however, most of the
studies have been conducted with Polydora.
Remove the required number of individuals from their tubes in the stock
culture and transfer with a pipette to a glass petri dish. Examine all spe-
cimens under the dissecting microscope for injured or reproductive
individuals. Discard all injured worms, and return reproductive specimens
to the stock colony. Maturing males usually appear milky white, posterior
to setiger 16, and females appear grey to pink when eggs are present. Place
15 to 20 ml of the desired toxicant solution in each 60 x 15 mm petri dish.
One or two worms can be kept in each dish. Use a minimum of 20 worms per
concentration. Use a total of five concentrations plus control. Add a
small amount (0.14) of fine sand (diameter 0.125-0.25 mm) to each dish as
material for tube construction. Examine for deaths daily (see below) and
discard all worms at completion of the experiment.
Long-Term Experiments (28 days)
Follow the same procedures as outlined for the 96-hour bioassay. Since
both species begin to show stress in about seven days if they have not been
fed or do not have materials for tube construction, it will be necessary to
feed the worms weekly. Provide each dish with approximately 0.1 g of fine
sand or silt for the tube construction material. Feed five pasteur pipette
drops of the Enteromorpha-'Bi.orel.l mixture (0.05 g) suspended in microporous-
filtered seawater per dish. The water should be renewed in each dish prior
to the weekly feeding. The condition of each worm should be determined
prior to feeding each week. If no movement can be seen (usually the palps
move about and can be easily seen with the unaided eye after some experi-
ence) , decant the fluid, invert the dish, examine the tube under the
dissecting microscope, and note the health of the worm within its tube.
Death in Adult Polydova and Boooardia—
Stressed spionids generally abandon their tubes, which will ultimately
lead to their death. While normal healthy spionids can construct a new tube,
a stressed specimen cannot. Once the specimen abandons its tube, it may
fragment, lose its palps, and be attacked by microorganisms. It is impor-
tant in both the 96-hour and 28-day bioassay experiment to note whether or
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not the specimen is within its tube since this can be an indication of sub-
lethal effects.
96-Hour Toxicity Tests with Larvae
Only limited success has been obtained thus far in conducting bioassays
with Polydora larvae; no tests have been carried out with Boeeardia. Be-
cause of the large number of larvae produced and because of the ease in
concentrating and collecting them with a light, the use of the larvae of
these species have a potential for being a useful larval bioassay organism
if techniques can be developed to greatly increase the survival rate.
Concentrate the larvae with a light, and pipette them into a petri dish.
Place 10 larvae in a petri dish containing toxicant. Use 10 dishes per con-
centration and a minimum of five concentrations plus control. Examine for
living larvae at 96 hours; dead ones may have decomposed by 96 hours. Since
many specimens may die during the experimental period, at least 85 percent of
the larvae should be living within the control to be considered a valid
experiment. Discard all specimens at the termination of the experiment.
Long-Term Experiments through Reproductive Period
No toxicity tests through a reproductive period have been conducted thus
far. The procedures outlined herein are intended to serve as a guide to con-
duct such an experiment. Place ten worms in a 100 x 20 mm petri dish together
with microporous-filtered seawater and 0.1 g of powdered sand or clay. Allow
24 hours for the specimens to construct a tube. Remove all worms that failed
to construct a tube. Decant off all seawater, and fill with the desired toxi-
cant solution. Since it is quite possible that a different number of
specimens will be present in each petri dish, a count of specimens should be
made so as to insure the same total number of specimens in each concentration.
There should be a minimum of five specimens per dish to insure fertilization
of eggs. The water should be changed and the worms fed at weekly intervals
as described for long-term adult bioassay experiments above. Each dish
should be checked every four days for the presence of egg capsules (Figures
20 and 25) inside the tubes of the female worms. This can be done by decant-
ing the water and inverting the bottom of the petri dish on the stage of the
dissecting microscope and examining each tube for egg capsules. The worms
can be exposed to air for up to five minutes without damage. The number of
capsules is noted and the average number of eggs per capsule estimated. After
a period of several weeks, the reproductive activity for each concentration
is expressed as total number of capsules and/or eggs produced per female per
concentration.
LABORATORY CULTURE OF DunaHella tertioleota
Medium
Seawater: Use about 30-32 °/oo salinity seawater (90 percent seawater
and 10 percent double distilled water). Filter through a microporous filter
system (0.45 y). For 50 ml quantities, mix complete medium (below) and
autoclave phosphate separately and then add to the medium. If a serious
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precipitation occurs, it may be necessary to filter sterilized medium prior
to use.
Major nutrients: Nitrate— 5 g KN03 made to 1,000 ml with H20. This
stock solution may be stored in a refrigerator for up to four months. Add
1.0 ml stock solution to one liter of distilled water to make a final
solution; use ten ml of this solution per liter of culture medium. Do not
store final solution.
Phosphate— 0.68 g KH2P04 made to 1,000 ml with H20. This solution may be
stored in a refrigerator for up to four months. Add 1.0 ml stock stolution
to one liter distilled water to make a final solution; use ten ml of this
solution per liter of culture medium. Do not store final solution.
Minor trace metals— Stock metals—30 mg ZnS04'7H20, plus 25 mg CuS04'5H20,
plus 20 mg CuS04'7H20 dissolved in 1,000 ml of distilled water.
Iron + Manganese— Dissolve 5.0 g FeCl3'6H20 and 2.0 g MnS04'H20 in 1,000 ml
of distilled water (ighore slight precipitate).
Sodium Molybdate— 25 mg Na2Mo04'2H20 in 1,000 ml of distilled water.
Sodium Ethylenediaminetetraacetate— Dissolve 50 g of N32E.D.T.A.-2H20 in
1,000 ml of distilled water.
Metal Mixture— Add 100 ml of the Na2E.D.T.A.-2H20 solution and ten ml each
to the three minor trace metal solutions listed above to about 800 ml of
distilled water. Adjust the pH to 7.5 with dilute NaOH solution and make up
to 1,000 ml with distilled water. Add 1.0 ml to each liter of culture
medium. This solution may be stored for up to four months in a refrigerator.
Vitamins: Vitamin Bi2—Dissolve 10.0 mg crystalline vitamin B]_2 in
100 ml of distilled water. Store in deep freeze until use. Immediately
prior to use, thaw the solution, remove 1.0 ml, and refreeze the stock solu-
tion. Dilute the one ml of vitamin Bi2 solution with 99 ml of distilled
water. Add 1.0 ml of diluted solution to each liter of seawater and then
throw away remainder.
i
Biotin— Dissolve 10.0 mg biotin in 100 ml of distilled water and freeze.
Immediately prior to use, thaw the solution, remove 1.0 ml, and refreeze the
stock solution. Dilute the 1.0 ml with 99 ml distilled water. Add 1.0 ml
of diluted solution to each liter of seawater and throw away the remainder.
Thiamine hydrochloride— Dissolve 100 mg of thiamine hydrochloride in 100 ml
of distilled water and freeze. Immediately prior to use thaw, remove 1.0ml.
and refreeze the stock solution. Dilute the 1.0 ml with 99 ml of distilled
water. Add 1.0 ml of diluted solution to each liter of seawater and throw
away the remainder.
Culture
Place 5-15 ml of DunoH-ella from a previous culture in a 500 ml Erlen-
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meyer flask containing 500 ml of autoclaved medium. The amount of
Dimal'iel'la depends upon its concentration in the previous culture. Place
stoppered Erlenmeyer flasks under grow-lux fluorescent lights. Depending
upon the amount of Dunal-iella required, the alga can be exposed to continu-
ous light or to a light-dark cycle, which is maintained with an electrical
timer. The growth rate can be slowed, if desired, by placing the cultures
in a cold bath (17°C) with the lights placed overhead.
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BIBLIOGRAPHY
Blake, J. A. 1969. Reproduction and larval development of Polydora from
northern New England (Polychaeta:Spionidae). Ophelia, 7:1-63.
Blake, J. A. 1971. Revision of the genus Polydora from the east coast of
North America (Polychaeta:Spionidae). Smithsonian Contributions to
Zoology, No. 75, 32 pp.
Hannerz, L. 1956. Larval development of the families Spionidae Sars,
Disomidae Mesnil, and Poecilochaetidae N. fam. in Gullmar Fjord
(Sweden). Zoologiska Bidrog Fran Uppsala, 31:1-204.
Hartman, C. 1941. Polychaetous annelids. Ill Spionidae. Some contribu-
tions to the biology and life history of Spionidae from California.
Allan Hancock Pacific Exped. 7:289-324.
Rice, S. A. 1978. Spermatophores and sperm transfer in spionid polychaetes.
Trans. Amer. Micros. Soc. 97:160-170.
Rice, S. A. 1975. The life history of Polydora li-gni, (Polychaeta:Spionidae)
including a summary of reproduction in the family Spionidae. Master's
Thesis, California State University, Long Beach, Long Beach, California,
129 pp.
Rice, S. A., and D. J. Reish. 1976. Egg capsule formation in the polychaete
Potydova ligni: confirmation of an hypothesis. Bull. So. Calif. Acad.
Sci. 75:285-286.
Woodwick, k. H. 1977. Lecithotrophic larval development in Boccardia
proboscidea Hartman. In: Essays on polychaetous annelids in memory of
Dr. Olga Hartman, D. J. Reish and K. Fauchald, eds. Allan Hancock
Foundation, University of Southern California, pp. 347-371.
91
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Figure 27. Polydora ligni, anterior end, dorsal view showing location of
palps, nuchal antenna, and fifth setiger.
92
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Figure 28. P. lign-L, posterior end, dorsal view showing disc-like
pygidium.
93
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Figure 29. P. 1i,gni, setae of modified fifth setiger showing one spine
with a subapical tooth and companion seta with forked tip.
94
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Figure 30. P. ligni, portion of a string of egg capsules as they appear
inside the tube of the female.
95
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Figure 31. P. ligni, three setiger larva just after release from the
female's tube.
96
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Figure 32. P. ligni,, 14 setiger larva ready to metamorphose into the
adult form.
97
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Figure 33. Boccardia proboscidea, anterior end, dorsal view.
98
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Figure 34. B. proboscidea, setae of the modified fifth setiger.
99
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Figure 35. B. probosaidea, portion of a string of egg capsules.
100
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Figure 36. B.' proboseidea, three setiger larva.
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Figure 37. B. proboscidea, 15 segmented larva.
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APPENDIX G
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELID:
jyinophilus sp.
CULTURE TECHNIQUES
Equipment and Supplies
A dissecting and compound microscope are required for examination and
identification of the archiannelid D-inophilus sp. Approximately one m^ of
shelf space is required in order to culture hundreds to thousands .of this
species per month. A piped compressed air system is the most suitable method
of supplying air to aquaria. While Dinophilus sp. is quite capable of with-
standing broad ranges of temperature, best results have been obtained when
the laboratory temperature is maintained at 19±1°C. Consumable supplies in-
clude 3.78 £ (1 gal) jars with covers, pasteur pipettes, petri dishes,
sieves, microporous filter system, air stones, and plastic tubing. A blender
is convenient for grinding food into finer particles for Dinophi-lus. Since
this polychaete is so small, it is best to pass all seawater through a micro-
porous filter (45 y) to remove microorganisms. Approximately 50 liters of
seawater will be more than adequate to culture this species per month.
Collecting Techniques and Life History
The genus Dinopkllus sp. is a minute polychaete that has been reported
from many regions of the world. Because of its small size, it is probably
more common than the previous reports indicate. It is impossible to observe
in the field because of its size. The most convenient way to collect
Dinophilus is to bring clumps of fouling organisms from boat floats or pil-
ings into the laboratory and place them in a pan of seawater. After a period
of time, examine the animals that crawl up along the sides of the pan under
the microscope and look for Dinophilus sp. (Figure 38). Remove with a pas-
teur pipette, and place in a petri dish containing seawater.
Species of the genus Dinophilus are very difficult to identify and re-
quire the help of an expert. Thus far, this laboratory population has not
yet been identified to species.
Members of the genus Dinophilus are dioecious. Eggs are laid in a small
capsule and generally number three to four per capsule (Figure 39). Sexual
maturity is generally reached within one week. Since this species has such
a short life history, it is unnecessary to use special techniques with
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different stages. Stock colonies can be maintained at 19±1°C with a minimum
of time by utilization of 3.78 & (1 gal) jars. Add 2,500 ml microporous-
filtered seawater to each container. Provide aeration with an aquarium stone
connected to a plastic tube and air supply. The top of the jar can be cov-
ered with a plastic top to minimize evaporation. Many specimens (20-50) can
be placed in a single gallon jar. Add about 1.0 ml of Tetramin solution
(see below) about every seven days. Since this species reproduces rapidly,
the population within the aquarium reaches its peak in about two to three
weeks then declines. Therefore, each jar should be reestablished about once
a month. The reproductive capacity of Dinophilus can be slowed by placing
the aquarium in a cold bath at 13-17°C.
Food for Dinophllus consists of finely-powdered Tetramin that is pre-
pared by grinding the flakes in a blender. The powdered Tetramin is then
placed on a 0.06 mm mesh sieve and shaken for a few minutes. Use only the
powder that passes through the sieve and collects within the pan. A 1 per-
cent mixture of fine powder to seawater is prepared, and 1.0 ml is pipetted
into each aquarium per week.
Problems in Culturing—
Most of the difficulty in culturing Dinophilus can be attributed to the
lack of regular, systematic care. However, since this species reproduces so
rapidly, ten aquaria should be sufficient to meet most laboratory needs.
Since the population within an aquarium reaches a peak within two to three
weeks then declines, it is important to keep a record of its establishment.
A fungal growth occurs whenever the animals are overfed. This fungal growth
can lead to a poor yield. If a fungal growth occurs, stir up the material
on the bottom and observe the next day. If the fungal growth reappears, re-
establish the aquarium.
BIOASSAY PROCEDURES
Equipment and Supplies
The only additional supplies required for conducting bioassays with
Dinoph'ilus are stender dishes or small plastic petri dishes.
96-Hour Experiments with DinopTritus
Because of its rapid life history, 96-hour bioassays may be difficult to
conduct since it is possible for this species to have reproduced during this
short experimental period. Only the smallest specimens should be used in
this experiment. Trial experiments should be carried out to determine
whether or not 96-hour tests can be conducted under conditions in the partic-
ular laboratory. Assuming it is possible to conduct 96-hour bioassays
successfully, pipette material from the bottom of an aquarium into a petri
dish and examine for young specimens under the dissecting microscope.
Transfer appropriate sized specimens into a second petri dish with a pasteur
pipette. Examine all specimens for injuries, and discard those that are not
suitable for experimentation. Place five specimens within a stender dish
containing 20 ml of seawater, use ten dishes per concentration. A minimum of
five concentrations of toxicant plus control should be employed for each
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test. Since these worms are so small, it is necessary to feed them during
the 96-hour experimental period. Add 0.1 ml of food to each container.
Count the number of living Dinophilus at the end of the experimental period.
Discard all specimens at the conclusion of the experiment.
Long-Term Experiment through Reproductive Period
Follow the same procedures outlined for the 96-hour experiment. Gener-
ally, the same experiment can serve both as a 96-hour and long-term bioassay.
Allow the experiment to extend to 21 days at which time all specimens should
be counted. The number of egg capsules containing embryos can be counted
and recorded separately.
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BIBLIOGRAPHY
Since this species of Dinophilus has not yet been identified (it pro-
bably represents an undescribed one), references are included herein to all
published work dealing with the living studies within the genus and some
more recent or important systematic studies of DinophLlus.
Akesson, B. 1973. Dinophilidermas (Archiannelida) systematiska stallning.
Zoologisk Revy. 35:76-78.
Bacci, G. 1956. Indicazioni di un effecto materna nella determinazione
sessuale di Dinophilus gyrociliatus ( = D. apatris). Revista di
Biologia, 26:71-76. *
Basedow, T. 1969. liber die Ausevirkung von Temperaturschocks auf die
Temperaturresistenz pokilothermer Wassertiere. Eine Untersuchung zum
Problem der thermischen Schockanpassung bei Tieren. Int. Revue ges.
Hydrobiol. 54:765-789.
Beauchamp, P. de. 1912. Contribution a I1etude experimentales de la sexu-
alite chez Dinophilus. Acad. Sci. Paris, C. R. 154:1836-1838.
Conklin, E. G. 1906. Sex differentiation in Dinophilus. Science.
24:294-296.
Gray, J. S., and Ventilla, R. J. 1971. Pollution effects on micro- and
meio-fauna of sand. Mar. Pollut. Bull. 2:39-43.
Griin, G. 1972. Uber den Eidimorphismus und die Oogeneses von Dinophilus
gyrociliatus (Archiannelida). Zeitschrift Zellforsch, mikrosk. Anat.
130:70-92.
Jagersten, G. 1944. Zur kenntnis der Morphologic, Enzystierung und
Taxonomic von Dinophilus. Kungliga Svenska vetenskapsakademiens
Handlingar. 21:1-90.
Jennings, J. B., and Gelder, S. R. 1969. Feeding and digestion in
THnopTrLlus gyioeil'Latus (Annelida:Archiannelida). J. Zool. Lond.
158:441-451.
Jones, E. R., Jr., and F. F. Ferguson. 1957. The genus Dinophilus
(Archiannelida) in the United States. Amer. Midi. Nat. 57:440-449.
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Jouin, C. 1971. Status and knowledge of the systematics and ecology of
Archiannelida, In: Proceedings of the First International Conference
on meiofauna. Smithsonian Contri. Zool. 76:47-56.
Korschelt, E. 1882. Uber Bau and Entwicklung des Dinophilus apatris.
Inaugural Dissertation. Zeits. wiss. Zool. Leipzig 37:315-333
Korschelt, E. 1887. Die Gattung Dinophilus under bei ihr auftretende gesch-
lechtsdimorphismus. Eine dritische Ausammenfassung neuerer und alterer
Forschungsergebnesse. Zool. Jahrb. Jena. Abt. Syst. 2:955-967.
Malikova, I. G. 1971. Epitalisation of the wound and encystment in
Dinophilus taeniatus Harmer after the cross sections. Vest. Leningr.
gos. Univ. (Biol.). 21:43-47 [In Russian].
Malikova, I. G. 1973. Observation of the regeneration in Dinophilus
taeniatus. H. Vestnik Leningr. gos. Univ. (Biol.). 24:14-19 [In
Russian].
I
Malsen, H. 1906. Geschlects-bestimmende Einflusse and Eiblidung des
Dinophilus apatris. Arch micr. Anat. Bonn. 69:63-99.
Nelsen, J. A. 1904. The Early Development of Dinophilus: A Study in Cell-
lineage. Proc. Acad. Sci. Nat. Phila. 56:687-737.
Repiachoff, W. 1886. On the anatomy and history of the development of
Dinophilus gyrociliatus. Odessa. 77 pp. [In Russian].
Schimkewitsch, W. 1895. Zur Kenntnis des Baues und der Entwicklung des
Dinophilus von Weissen Meere. Zeits. wiss. Zool., Leipzig. 59:46-79.
Shen, T. H. 1936. Beitrage zur Studium der Geschlechtsbestimmung bei
Dinophilus apatris. Zool. Jahrb. Jena, Physiol. 56:219-238.
Stiasny, G. 1910. Dinophilus apatris, var. nov. tergestina. Zool. Anz.
Leipzig. 35-587.
Sudzuki, M., and Sekiguichi> K. 1972. Some remarks on five abberant anne-
lids from the culture water of Japanese horse-shoe crabs. Science Rep.
Tokyo Kyoiku Daig. (b) 15:39-56.
Traut, W. 1966. Eine Mutante mit vergrosserten Mannchen-Eiern bei
Dinophilus gyrociliatus (Archiannelida). Experientia 22:237-238.
Traut, W. 1966. Uber due Kopulation bei Dinophilus gyrociliatus
(Archiannelida). Zool. Anz. 177:402-411.
Traut, W. 1968. Genetische Analyse zweier Mutauten von Dinophilus
gyrociliatus (Archiannelida) mit veranderter Eigrosse. Helgolander
Wiss. Meeresunters. 18:296-316.
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Traut, W. 1969. Zur Sexualitat von Dinophilus gyroeiliatus (Archiannelida)
1. Der Einfluss von asussenbedingungen und genetiachen Faktorene auf
das Geschlechtsverhaltnis. Biol. Zbl. 88:469-495.
Traut, W. 1969. Zur Sexualitat von Dinophilus gyriociliatus (Archianne-
lida) . 2. Der Auflbau des Ovars und die Oogenese. Biol. Zbl.
88:695-714.
• •
Traut, W. 1969. Uber den Geschlechtsbestimmungomodus bei Dinophilus.
Zool. Anz. Suppl. 32:260-265.
Traut, W. 1970. Zur sexualitat von Dinophilus gyrioeiliatus (Archianne-
lida) . 2. Die geschlechtsbestimmung. Biol. Zbl. 89:137-161.
Tzonis, K. 1938. Beinflussing der Geschlechtsverhaltnisse bei Dinophilus
apatris Korschelt durch Assenbedingungen. Zool. Jahrb. Jena.
Physiol. 58:539-550.
Wieser, W. Archiannelids from the intertidal of Puget Sound. Trans. Amer.
Micros. Soc. 76:275-285.
108
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Figure 38. Dinopkilus sp., dorsal view of entire worm.
109
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Figure 39. Di.nophilus sp. , egg caps-ule.
110
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APPENDIX H
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELIDS:
Cirriformia luxwiosa and Cirriformia
spirabrancha
CULTURE TECHNIQUES
Equipment and Supplies
Dissecting and compound microscopes and a blender are the only pieces of
equipment required to culture these two species of Cirriformia. A minimum of
10-12 m of shelf space is necessary in order to culture populations of 1,000-
3,000 specimens each of these two species. A constant temperature laboratory
set at 19±°C is an ideal environment for both of these species; however, if
time becomes a critical factor, then placing the aquaria in a cold room (or
cold bath) at 15-17°C slows the metabolic rate sufficiently to reduce the
amount of time necessary for routine maintenance of the populations. Natu-
ral, filtered seawater (35 °/oo salinity) is the most satisfactory source of
seawater; approximately 5,000 liters of seawater per month is needed to main-
tain populations of 2,500 specimens of these two species. Because of the
frequent feedings and periodic water changes, it will be necessary to spend
15 hours per week to maintain these populations. A central compressed air
system is the most satisfactory source of air for Cirrifornria. Consumable
supplies include 3.78 £ (1 gal) jars and 56.1 £ aquaria, medium grade aquar-
ium sand, forceps, brushes, petri dishes, pasteur pipettes, aquarium stones,
plastic tubing, and food supply.
Collecting Techniques and Life History
Both species of Cirriformi-a are known primarily from California, al-
though both species have been reported from the gulf of California. A
related species, C. tentaculta, found in European waters, is similar, and the
techniques described for C. luxuriosa and C. spirabrancha would probably
apply equally well to C. tentaaulata. Both species were collected from
Aland.tos Bay in Long Beach but from different localities. Both were taken
from fine sediments that generally possessed sulfide odors. The worms live
in the upper three to six cm of sediment and can be collected either by hand
or by sieving the sediment through a course screen (2.0 mm mesh).
Ciwiformia Zuxuriosa also occurs within Mytilus edulis communities attached
to floating boat docks, especially where the water circulation is limited.
Because of the possession of numerous gills throughout much of the length of
the body, specimens become entangled and are nearly impossible to separate.
Ill
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The field-collected specimens should be placed in pans with about 1 cm thick-
ness of medium grade aquarium gravel (2.0 mm diameter) and 3-5 cm of
seawater. The specimens will burrow into the gravel and become separate from
one another. The untangled worms can then be placed in either 3.78 £ jars
(5 specimens/aquarium) or 56.1 H (15 gal) aquaria (50-100) and provided with
one to two cm thickness of gravel, seawater, an air supply, and food.
Living specimens of these two species are easily distinguished from one
another by color of the tentacles; C. luxuriosa have red tentacles (Figure
40) , and C. spirdbrancha (Figure 41) have green ones. Unfortunately, this
color difference disappears upon preservation. Preserved specimens are dis-
tinguished by one or rarely two heavy black spines in both the posterior
notopodial and neuropodial regions in C. luxwpiosa (Figure 42) and several
yellow spines in both the posterior notopodial and neuropodial regions in
C. spirabranaha (Figure 43) . The differentiation of preserved juvenile spe-
cimens is more difficult because the spines in C. luxuriosa may not yet be
either big or black.
The life cycles of both CiTviformia luxuri-osa and C. spi-vabrano'ha are
long, apparently two years and one year, respectively, and as a result, only
a limited amount of success in culturing this species from egg to egg has
been accomplished. Since both of these species are large (lengths up to
16 cm and wet weight up to 4.0 g) and their life cycles long, the potential
value of these species lies in long-term studies involving the uptake of
toxicants. The sexes are separate and the eggs and sperm are released
through pores just anterior to the neurosetae. Both sexes emerge from the
sediment to release their gametes onto the surface where fertilization
occurs. Females spawn an estimated 100,000 to 500,000 oocytes at one time.
Trochophores appear at 16 hours and become elongated (Figure 4) at 2 days.
Larval settlement occurs on the fifth day, and feeding commences on the
eighth day. Two setigerous segments appear on the eleventh day when the
first and second branchial buds are noted (Figure 45) . Growth continues at
the rate of one new segment per day (Figure 46) . Gametes are noted in both
species at nine months. Spawning occurred after one year in C. spirabranc'ha
but had not yet occurred in C. luxuriosa after two years, at which time the
colony was destroyed.
A mixture of Enteromorpha sp. and Biorell is the most satisfactory food
source. Large quantities of the green alga Entevomoypha sp. can be collected
from estuaries during high tides. The alga is collected by hand and washed
at the site to remove as much adhering debris and sediments as possible. The
algae is spread out on chicken wire and allowed to air-dry. When it is dry,
usually within 24 hours, it is gathered and can be stored for an indefinite
period of time. Biorell is a commercially available fish food flake. The
mixture of EntePomoYpha and Biorell is made at the ratio of 4:1 by dry ,
weight. This mixture is pulverized in a blender and shaken through a sieve
possessing a 0.06 mm opening. Feed smaller sized worms 0.05 g per worm five
times a week. The amount of food should be increased with the growth of the
worms. Feed worms over ten cm in length 0.3 g per worm, five times a week.
To judge whether the colony of Civvifoicmia is being overfed, examine the
condition of the surface of the gravel. If the gravel is kept clean, the
worms are not being overfed. Increase the amount of food per feeding until
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debris begins to accumulate on the surface of the gravel The seawater
should be changed about every two weeks at which time the gravel should be
rinsed to remove any buildup of organic material.
Problems in Culturing—
Maintaining a colony of Cirri-formi-a luxuriosa and C. spirabranc'ha re-
quires daily care. If such care is not given these species, then growth may
be slowed and black sulfide deposits may build up, which could lead to death
of the specimens. The surface of the gravel within each aquarium should be
examined daily prior to feeding to determine whether or not the colony is
being underfed or overfed. It is important that the gravel be washed every
two weeks to remove the fecal material. The seawater should be changed at
this time. There are no known enemies of either species, nor are the
species cannibalistic.
TOXICITY TEST PROCEDURES
Equipment and Supplies
Toxcity tests can be conducted in 500 ml Erlenmyer flasks; other sup-
plies required were specified under culture techniques. Stender dishes or
small petri dishes are satisfactory containers for larval bioassays.
96-Hour Experiments with Adults
Remove specimens from stock cultures individually and place in separate
petri dishes. This procedure eliminates any possibility of the gills of
specimens becoming entangled with one another, which would greatly increase
the time in setting up a bioassay test. Examine each specimen individually
under a dissecting microscope for injuries, and discard such specimens.
Place a single worm in a 500 ml Erlenmeyer flask containing 100 ml of toxi-
cant. Close the flask with a silicone stopper. Use 20 worms per concentra-
tion and a minimum of five concentrations plus controls. Examine each worm
daily for death, and discard all specimens.
96-Hour Experiments with Trochophore Larva
Since both species of Civpi-formla lay tens of thousands of eggs, many
larval toxicity tests can be conducted from the fertilized eggs from one
spawning. Draw up ten trochophore larvae with a pasteur pipette, and place
in a dish containing seawater and the toxicant; use a minimum of five dishes
per conentration and a minimum of five concentrations plus control. No food
is required for a 96-hour experiment. Examine at the end of the experiment-
al period for living trochophores, and destroy all specimens at this time.
Dead torchophores will decompose within a short period of time and cannot
generally be found by the end of the experimental period.
Death in Cirriformia—
Death in both species of Cirriformia is defined as the absence of move-
ment when the flask is gently rolled. Rupture of the body wall often occurs
just prior to death. The body gills of stressed Cirriforrrria are usually
fragmented.
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Long-Term Experiments with Adults
Since both species of Cirriformia are capable of surviving one to two
months without feeding, the same experiment can be utilized for both the 96-
hour and 28-day bioassay. If the objectives are to conduct a long-term
experiment over several months, it will be necessary to provide specimens
with food (in the form and amounts specified under the procedures outlined
for culturing these worms). Because of the large size of the worm, both of
these species may be convenient test organisms for long-term experimentation
to study uptake, retention, and metabolism of a toxicant. However, it is
important to note that such an experiment has not yet been conducted with
either species. If such an experiment is contemplated, the procedures out-
lined under the section on culturing should be followed. A large number of
specimens (25 to 50} could be placed in a 56.1 liter aquarium (Provide ap-
proximately one liter of seawater per specimen). Gravel must be provided
and the worms fed the required food five times a week. Either species could
be used to conduct long-term studies in a flow-through system.
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BIBLIOGRAPHY
Blake, J. A. 1975. The larval development of polychaeta from the northern
California coast I. Cirriformia spirabranchia (Family Cirratulidae).
Trans. Amer. Micros. Soc. 94:179-188.
Carr, R. S., and D. J. Reish. 1977. The effect of petroleum hydrocarbons on
the survival and life history of polychaetous annelids. In: Rate and
Effects of Petroleum Hydrocarbons in Marine Organisms and Ecosystems.
D. A. Wolfe, ed. Pergamon Press, New York, pp. 168-173.
Milanovich, F. P., R. Spies, M. S. Goram, and E. E. Sykes. 1976. Uptake of
copper by the polychaete Cirri-fo^mia sp-irabranch-ia in the presence of
dissolved yellow organisms of natural origin. Estuarine Coastal Mar.
Sci. 4:585-588.
Moore, J. P. 1904. New polychaeta from California. Proc. Acad. Nat. Sci.
Phila. 56:484-503.
Reish, D. J., and T. L. Richards. 1966. A culture method for maintaining
large populations'of polychaetous annelids in the laboratory. Turtox
News. 44:16-17.
115
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Figure 40.
ia luxuriosa, anterior end, dorsal view.
116
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Figure 41. C. spirabranefia,, anterior end, dorsal view.
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Figure 42. C. luxuviosa, outline of segment 150 (after Moore, 1904)
118
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Figure 43. C. spirdbranaha, outline of segment 150 (after Moore, 1904)
119
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Figure 44. C. luxuri-osa, late trochophore stage.
120
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Figure 45. C1.
metamorphosed juvenile.
121
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Figure 46. C. luxwrlosa, late juvenile stage.
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APPENDIX I
CULTURE AND BIOASSAY PROCEDURES
FOR POLYCHAETOUS ANNELID:
Halosydna johnsoni,
CULTURE TECHNIQUES
Equipment and Supplies
A dissecting and 'compound microscope are needed for observation of the
various stages in the life history of Halosydna johnsoni. Some method of
maintaining a temperature of 18°C (constant temperature room or water bath)
is needed. A cold bath measuring 1.2 x 2.4 m will provide sufficient space
to culture and maintain approximately 500 adults in petri dishes, which are
placed in enamel pans and 25 3.78 H jars containing juveniles. Since each
adult must be fed individually twice a week, approximately ten hours a week
is necessary to maintain a population of Halosydna of this size. Supplies
required include petri dishes, glass tubing, aquarium stones, plastic tubing,
plankton netting (Nitex 7 y), pasteur pipettes, microporous filter system,
brushes, 25 x 35 cm trays, forceps, and screens with 0.147 and 0.06 mm open-
ings. Food consists of frozen brine shrimp, a mixture of Biorell, Tetramin,
and alfalfa powder, and a Dunaliella culture.
Collecting Techniques and Life History
Halosydna johnsoni is the most commonly occurring scale worm in southern
California (Figure 47). A closely related species, H. brevisetosa, is found
in colder waters of the eastern Pacific Ocean; however, the distribution of
the two species can overlap. These two species differ primarily in the type
of neurosetae. Some question of the validity of the two species has been
raised, since it has been demonstrated experimentally that each "species"
can develop the other "species" setal type in warm or cold temperatures
(Hillger and Reish, 1970). Since all specimens reared from fertilized eggs
to adults developed only bifid neurosetaes, the specific name H. jolmsoni. is
retained herein as the valid name for this population with the realization
that subsequent studies may not substantiate this conclusion.
Halosydna is most easily collected from the fouling community attached
to floating boat docks since collections can be made independent of tidal
conditions. Clumps of mussels (Mytilus eduHs) and other organisms are re-
moved from the dock float and placed in shallow pans. Separate this material
to free the worms. Remove the worms from the pan with a brush, and place in
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a separate container for transport to the laboratory. In the laboratory,
transfer the specimens to individual petri dishes. Place the dishes in trays
in an 18°C cold water bath. Two to three days later place a piece of glass
tubing, which is of the appropriate length and diameter for the given worm,
in each petri dish. The worm is maneuvered into the tubing with a small
brush. If the worm attempts to vacate the tube, the protruding end is tapped
lightly. This process is repeated until the worm accepts the tube as its
new home. One week after collection the worms are fed a few frozen brine
shrimp. For the first few feedings it is necessary to grasp several brine
shrimp with a pair of forceps and wiggle them near the anterior end of the
worm to initiate a feeding response. After three to five feedings most
specimens are trained sufficiently to accept the food so that it is only
necessary to place the brine shrimp in the vicinity of the anterior end. The
worms are fed until satiated twice a week. Change the seawater after each
feeding.
The sexes are separate and can only be distinguished after development
of the gametes. To distinguish one sex from the other, examine the ventral
surface; females appear olive-green because of the presence of maturing ova,
and the males are white as the result of maturing sperm. Spawning is in-
duced by transferring mature specimens from 18°C to a petri dish containing
water with a temperature of 20 to 25°C. Actual release of sperm has not been
observed, but all gametes are not released at the same time. Females release
their ova through paired ventrally situated nephridial papillae. Initially,
the ova are released individually at a more or less staccato rate that then
becomes a steady emission followed again by the staccato rate of release.
Nearly all ova are released at one spawning. Up to an estimated 240,000 ova
are produced by one female; the ova are small, flattened somewhat, and
measure 88 to 94 y long and 40 y thick. Gametes of both sexes are trans-
ferred from petri dishes to a 3.78 & jar containing 18°C seawater; the water
is stirred intermittently over a 15-minute period. The jar is filled with
seawater and divided into three additional jars that in turn are filled with
18°C seawater. Place the jar in an 18°C cold bath and provide it with
aeration. In order to measure the success of fertilization, remove the jar
from the cold bath and place a strong light at the top to attract the trocho-
phore larvae (Figure 48). The density of the trochophores at the surface is
noted as well as the number of unfertilized eggs or undeveloped zygotes ad-
hering to the bottom of the aquarium. That portion of the water containing
trochophores is decanted, diluted two to three times, and poured into the
appropriate number of jars; the amount of dilution is dependent upon the
density of the trochophores in the initial jars. After dilution, place the
jars in the cold bath at 18°C and provide aeration. At six days, 60 ml of
Uwnal'lella -tevb-loteo-ta culture are added to each aquarium followed by an
additional 20 ml of culture every five days for three times. The majority
of the larvae metamorphose (Figure 49) at about 20 to 25 days and settle to/
the bottom. Do not feed between days 25 to 40 at which time the contents of
the jar are filtered through a 7 y nitex plankton netting. The netting is
placed in a petri dish containing seawater and examined under a dissecting
microscope for presence of larvae. Transfer the young Hdtosydna with an
appropriate sized pasteur pipette and place in a clear 3.78 & aquaria con-
taining 3:1 of millipore-filtered seawater and 0.6 g of a mixture of two
parts (dry weight) of alfalfa powder, one part of Biorell powder, and one
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part Tetramin powder. A maximum of 50 larvae (Figure 50) can be placed in
each jar, which is then placed in the 18°C cold bath. After 60 to 70 days
of development, specimens with 25 or more segments can be removed and placed
in individual petri dishes and fed brine shrimp as described for maintaining
adults. Sexual maturity is reached after nine months of growth.
Problems in Culturing—
Since the adults are aggressive and cannibalistic, they must be kept in
separate containers. In addition, each one must be fed individually with
frozen brine shrimp. The individual personal attention required by each
adult necessitates a considerable amount of time. If the proper attention
and care are not given the adults, death can follow quite rapidly because of
fouling of the water. As typical for most species having pelagic larvae,
many larvae fail to metamorphose. Undoubtedly, if more time and attention
were spent caring for the culture a higher percentage of the larvae could
survive.
TOXICITY TEST
Equipment and Supplies
No additional equipment and supplies are required to carry out bio-
assays other than those specified above.
96-Hour Experiments with Adults
Place 50 ml of seawater and toxicant in a ten cm diameter petri dish.
Transfer each H. johnsoni in its glass tubing to a petri dish containing
clean seawater, and examine for injuries under the dissecting microscope.
Then transfer all healthy specimens to the experimental container. Use 20
worms per concentration with a minimum of five concentrations plus control
for each bioassay. Do not feed the animals during the course of the experi-
ment. Examine for deaths daily, and discard all specimens at the conclusion
of the experiment.
Long-term Experiment with Adults (28 Days)
Set up the experiment the same as the 96-hour test; in fact, the same
experiment can serve as both by following the procedures given below. After
examination for deaths at 96 hours feed each animal with frozen brine shrimp
as indicated under the techniques of culturing the adults. After feeding,
renew the seawater and toxicant; feed twice a week with water changes made
after each feeding.
96-Hour Experiments with Trochophore Larvae
The trochophore larvae of H. johnsoni, are sensitive and especially use-
ful for 96-hour tests. Separate out the trochophore larva from the top of
the aquarium as specified above. Pipette larvae into a petri dish containing
seawater. Use ten larvae per dish with ten petri dishes per concentration
and five test concentrations plus control. Count the number of living tro-
chophores at the end of 96 hours and discard all specimens at the conclusion
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of the experiment.
Death in Hdlosydna—
Stressed adult Ealosydna will usually leave the glass tubing and may or
may not exhibit erratic movements such as turning over and over. Specimens
under severe stress, such as those specimens placed in a highly toxic solu-
tion, may go into convulsions and fragment at the mid-body region. Death
follows rapidly in this case. Specimens in a solution at or near the calcu-
lated LCso will generally leave the tube, leytrae (scales) will be cast off,
and dorsal cirri may fragment. These specimens may live beyond a 28-day
period, but they will not feed.
Dead trochophore larvae are often difficult to find because decomposi-
tion is so rapid in these young worms. It is much more convenient to count
living specimens. Trochophore larvae are considered dead if they fail to
move through the water mass and if there is an absence of ciliary movement.
A stressed trochophore is generally more or less motionless, but its cilia
will continue to move at a reduced rate.
Use of Ealosydna johnsoni in Bioassays
Ealosydna johnsoni, has only been utilized to a limited extent in bio-
assays. The advantages of this species as a test organism are the ease in
obtaining a large number of trochophore larvae, the sensitivity of both lar-
vae and adults to many toxicants, and the large size of adults that permits
body burden analyses of individual specimens. The primary disadvantage of
this species is the considerable amount of time that must be spent in main-
taining a relatively small population.
LABORATORY CULTURE OF Dunaliella tevtioleota
Medium
Seawater: Use about 30-32 °/oo salinity seawater (90% seawater and 10%
double distilled water). Filter through a microporous-filter system (0.45y).
For 50 ml quantities, mix complete medium (below) and autoclave at 15 "psi for
15 minutes. For larger quantities autoclave phosphate separately and then
add to the medium. If a serious precipitation occurs, it may be necessary
to filter sterilized medium prior to use.
Major nutrients: Nitrate—5 g KN03 made to 1,000 ml with H20. This
stock solution may be stored in a refrigerator for up to four months. Add
1.0 ml stock solution to one liter distilled water to make a final solution;
use ten ml of this solution per liter of culture medium. Do not store final
solution.
Phosphate—0.68 g KH2P04 made to 1,000 ml with H20. This solution may
be stored in a refrigerator for up to four months. Add 1.0 ml stock solution
to one liter distilled water to make a final solution; then ten ml of this
solution per liter of culture medium. Do not store final solution.
Minor trace metals-- Stock metals—30 mg ZnS04'7H20, plus 25 mg
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CuS04'5H20, plus 20 mg CuS04-7H20 dissolved in 1,000 ml of distilled water.
Iron + Manganese—Dissolve 5.0 g FeCl3'6H20 and 2.0 g MnS04'H20kn 1,000
ml distilled water (ignore slight precipitate).
Sodium Molybdate—25 mg Na2Mo04'2H20 in 1,000 ml of distilled water.
Sodium Ethylenediaminetetraacetate—Dissolve 50 g of Na2E.D.T.A.-2H20
in 1,000 ml of distilled water.
Metal Mixture—Add 100 ml of the Na2E.D.T.A.-2H20 solution and ten ml
each of the three minor trace metal solutions listed above to about 800 ml
of distilled water. Adjust the pH to 7.5 with dilute NaOH solution and make
up to 1,000 ml with distilled water. Add 1.0 ml to each liter of culture
medium. This solution may be stored for up to four months in a refrigerator.
Vitamins:Vitamin Bi2—Dissolve 10.0 mg crystalline vitamin B]_2 in
100 ml of distilled water. Store in deep freeze until use. Immediately
prior to use, thaw the solution, remove 1.0 ml, and refreeze the stock solu-
tion. Dilute the ten1 ml of vitamin Bi2 solution with 99 ml of distilled
water. Add 1.0 ml of diluted solution to each liter of seawater and then
throw away remainder.
Biotin—Dissolve 10.0 mg biotin in 100 ml of distilled water and freeze.
Immediately prior to use thaw the solution, remove 1.0 ml, and refreeze the
stock solution. Dilute the 1.0 ml with 99 ml distillled water. Add 1.0 ml
of diluted solution to each liter of seawater and throw away the remainder.
Thiamine hydrochloride—Dissolve 100 mg of thiamine hydrochloride in
100 ml of distilled water and freeze. Immediately prior to use thaw, remove
1.0 ml, and refreeze the stock solution. Dilute the 1.0 ml with 99 ml of
distilled water. Add 1.0 ml of diluted solution to each liter of seawater
and throw away the remainder.
Culture
Place 5 to 15 ml of Dunaliella from a previous culture in a 500 ml
Erlenmeyer flask containing 500 ml of autoclaved medium. The amount of
Dunaliella depends upon its, concentration in the previous culture. Place
stoppered Erlenmeyer flasks under grow-lux florescent lights. Depending
upon the amount of Dunal'Le'l'la required, the alga can be exposed to continuous
light or to a light-dark cycle, which is maintained with an electrical timer.
The growth rate can be slowed, if desired, by placing the cultures in a cold
bath (17°C) with the lights placed overhead.
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BIBLIOGRAPHY
Gaffney, P. M. 1973. Setal variation in Halosydna brevisetosa polychaeta.
Sys. Zool. 22:171-175.
Hillger, K., and D. J. Reish. 1970. The effect of temperature on the setal
characteristics in Polynoidae (Annelida:Polychaeta). Bull. So. Calif.
Acad. Sci. 57:99-106.
Pettibone, M. H. 1953. Some scale-bearing polychaetes of Puget Sound and
adjacent waters. Univ. Washington Press, Seattle. 89 pp.
Skosberg, T. 1942. Redescription of three species of the polychaetous
family Polynoidae from California. Proc. Calif. Acad. Sci., Ser. 4,
23:481-502.
MacGinitie, G. E., and N. MacGinitie. 1949. Natural history of marine
animals. McGraw Hill Book Co., New York, 473 pp.
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Figure 47. H. johnsoni, anterior end, dorsal view.
129
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Figure 48. H. johnsvni, trochophore larva.
130
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Figure 49. H. Qohnsoni, metatrochophore larva.
131
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Figure 50. H. johnsoni, metamorphosed juvenile.
132
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APPENDIX J
LIST OF PUBLICATIONS AND THESES
SUPPORTED BY THIS RESEARCH GRANT
PUBLICATIONS
Reish, D. J., J. M. Martin, F. M. Piltz, and J. Q. Word. 1974. The
Induction of Abnormal Polychaete Larvae by Heavy Metals. Marine
Pollution Bulletin, 5:125-126.
Oshida, P. S. , and D. J. Reish. 1974. The Effect of Various Water
Temperatures on the Survival and Reproduction in Polychaetous Annelids:
Preliminary Report, Marine Studies of the San Pedro Bay, Calif. Part 3.
Allan Hancock Foundation, pp. 63-77.
Oshida, P. S. , and D. J. Reish. 1975. Effect of Chromium on Reproduction
in Polychaetes. Annual Rept., So. Calif. Coastal Water Res. Project.
pp. 55-60.
Oshida, P. S., A. J. Mearns, D. J. Reish, and C. S. Word. 1976. The Effect
of Hexavalent and Trivalent Chromium on Neanthes arenaceodentata.
(Polychaeta: Annelida). So. Calif. Coastal Water Research Project,
Tech. Mem. No. 225, 58 pp.
Reish, D. J., J. M. Martin, F. M. Piltz, and J. Q. Word. 1976. The Effect
of Heavy Metals on Laboratory Populations of Two Polychaetes with
Comparisons to their Water Quality Conditions and Standards in
Southern California. Water Res., 10:299-302.
Anon., 1976. Bioassay Procedures for Marine Polychaete Annelids (Tentative)
In: Standard Methods for the Examination of Water and Wastewater. 14th
Edition, APHA, AWWA, and WPCF, Washington, D.C., pp. 785-793. (Note:
The editorial policy of "Standard Methods" does not allow for the in-
clusion of research grant support acknowledgement nor author. This
section was written by D. J. Reish during the tenure of this research
grant.)
Reish, D. J. 1976. The Establishment of Laboratory Colonies of Polychaetous
Annelids. Thalassia Jugoslavia., 10:181-195.
Rice, S. A., and D. J. Reish. 1976. Egg Capsule Formation in the Polychaete
Polydora l-Lgni: Confirmation of a Hypothesis. Bull. So. Calif. Acad.
Sci., 75:285-286.
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Mearns, A. J., P. S. Oshida, M. J. Sherwood, R. R. Young, and D. J. Reish.
1976. Chromium Effects on Coastal Organisms. Jour. Water Pollution
Control Fed., 49:1929-1939.
Reish, D. J. 1977. Effect of Chromium on the Life History of Capitella
capitata. In: Physiological Responses of Marine Biota to Pollutants,
F. J. Vernberg, A. Calabrese, F. P. Thurberg, and W. B. Vernberg, eds.,
Academic Press, New York, pp 199-207.
Reish, D. J. 1977. The Role of Life History Studies in Polychaete
Systematics. In: Essays in Memory of Dr. Olga Hartman, D. J. Reish
and K. Fauchald, eds. Allan Hancock Foundation, Univ. of So.
California, pp. 461-476.
Carr, R. S., and D. J. Reish. 1977. The Effect of Petroleum Hydrocarbons
on the Survival and Life History of Polychaetous Annelids. In: Fate
and Effect of Petroleum Hydrocarbons in Marine Organisms and Ecosystems,
D. A. Wolfe, ed. Pergamon Press, New York. pp. 168-173.
Reish, D. J., and R. S. Carr. 1978. The Effect of Heavy Metals on the
Survival, Reproduction, and Life Cycles for Two Species of Polychaetous
Annelids. Marine Pollution Bull., 9:24-27.
Reish, D. J., T. V. Gerlinger, C. A. Phillips, and P. Schmidtbauer. 1977.
Toxicity Formulated Mine Tailings on Marine Polychaeta. Rept. to
Environment Canada, Winnipeg, Manitoba. Marine Biological Consultant,
Inc., Costa Mesa, California, 104 pp.
Reish, D. J. 1978. The Effect of Heavy Metals on Polychaetous Annelids.
Rev. Int. Oceanogr. Med., 49:99-104.
Reish, D. J., C. E. Pesch, J. H. Gentile, G. Bellan, and D. Bellan-Santini.
1978. Interlaboratory Califration Experiments Using the Polychaetous
Annelid Capitella. oop-itata. Marine Environmental Research, 1:109-118.
Rice, S. A. 1978. Spermatophores and Sperm Transfer in Spionid Polychaetes.
Trans. Amer. Micros. Soc., 97:160-170.
Anon. Bioassay Procedures for Marine Polychaete Annelids (Tentative). In:
Standard Methods for the Examination of Water and Wastewater, 15th
Edition, APHA, AWWA, and WPCF, Washington, D.C. In press, 1979.
(Note: written by D. J. Reish.)
THESES
Piltz, F. M. 1974. The Effect of Copper on Reproduction of Two Polychaetous
Annelids, Capitella capitata (Fabricius) and Ophpyotrocha ap. Master's
Thesis, Calif. State Univ., Long Beach, 107 pp.
Word, J. Q. 1974. The Effect of Zinc on Reproduction of Two Polychaetous
Annelids, Capitella oapi>tata (Fabricius) and Opkryotrocha sp. Master's
Thesis, Calif. State Univ., Long Beach, 43 pp.
134
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Rice, S. A. 1975. The Life History of Polydora ligni Polychaeta Spionidae)
Including a Summary of Reproduction in Family Spionidae. Master's
Thesis, Calif. State Univ., Long Beach. 129 pp.
Shisko, J. S. 1975. The Life History of the Annelid (Polychaeta:
Serpulidae) Jania (Dexiospira) brasiliensis (Grube). Master's Thesis,
Calif. State Univ., Long Beach. 55 pp.
Morgan, D. E. 1975. Life Histories of the Cirratulid worms Cirriformi-a
luxur-iosa and C'LvvLform-la spirabrancha (Annelida:Polychaeta) . Master's
Thesis, Calif. State Univ., Long Beach. 172 pp.
Carr, R. S. 1976. The Effects of Petrochemicals on Five Species of
Polychaetous Annelids, Ctenodrilus serratus3 Ophryotrocha sp.,
Ophryotrocha puerilis, Capitella capitata, and Cirriformia spirabrancha.
Master's Thesis, Calif. State Univ., Long Beach. 92 pp.
King, K. M. 1976. The Life History of Bocccaedia proboscidea Hartman
(Polychaeta:Spionidae). Master's Thesis, Calif. State Univ., Long
Beach. 118 pp.
Rossi, M. 1976. The Life History of Halosydna brevisetosa (Kinberg)
(Polychaeta:Polynoidae). Master's Thesis, Calif. State Univ., Long
Beach. 132 pp.
135
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APPENDIX K
List of Laboratories Which Have Utilized Specimens
Cultured in California State University, Long Beach
Many laboratories have obtained specimens or subcultures of polychaetes
from this laboratory for use in their own studies. One valuable spin-off
from developing the techniques for culturing polychaetes through complete
life cycles and establishing laboratory inbred colonies is that it makes it
possible for others to conduct studies which would otherwise not have been
possible. Specimens have been sent to the following laboratories through
1979.
1. Environmental Research Laboratory, Narragansett, Rhode Island
(Bioassays with metals).
Capitella capitata
Cirrifornria luxuriosa
Ctenodpilus serratus
Neanthes arenaoeodentata
2. U.S. Navy, Anapolis, Maryland (Bioassays with dredge spoils).
Neanthes arenaeeodentata
3. U.S. Navy, San Diego, California (Bioassay with dredge spoils).
Neanthes arenaceodentata
4. U.S. Army Corps of Engineers, Vicksberg, Mississippi (Bioassays).
Neanthes arena.ceodenta.ta.
5. Texas A&M University (Bioassays with petroleum hydrocarbons).
Capitelrla. capitata
Neanthes arenooeodentata
6. Scripps Institute of Oceanography (Bioassays with petroleum
hydrocarbons).
Capitella capitata
Neanthes arenaoeodentata
7. Allan Hancock Foundation, University of Southern California
(Bioassays with many different types of toxicants).
Neanthes arenaoeodentata
Ophryotrooha d-iadema
136
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8. School of Medicine, University of California Los Angeles (Medical
research with eggs).
Neanthes arenaceodentata
9. University of California, Irvine, California (polychaete metabolism).
Neanthes arenaceodentata
10. University of Alberta, Calgary, Alberta, Canada (trematod life cycles).
Neanthes arenaceodentata
11. McGill University, Montreal, Quebec, Canada (experimental studies with
eggs).
Capitella capitata
12. Station Marine d'Endoume, Marseille, France (Bioassays with metals).
Capitella capitata
13. Southern California Coastal Water Research Project, El Segundo,
California (Bioa'ssays) .
Ctenodrilus serratus
Neanthes arenaceodentata
14. Orange County Sanitation District, Fountain Valley, California
(Bioassays).
Neanthes arenaceodentata
15. NUS Corporation, Houston, Texas (Bioassays).
Neanthes arenaceodentata
16. EG&G, Bionomics, Pensicola, Florida (Bioassays).
Neanthes arenaoeodentata
17. Lockheed Corporation, Carlsbad, California (Bioassays).
Neanthes arenaceodentata
18. Marine Biological Consultants, Inc., Costa Mesa, California
(Bioassays).
Capitella capitata (
Ctenodrilus serratus
Neanthes arenaceodentata
Ophryotrocha diadema
19. Batelle Northwest Laboratories, Washington (Bioassays with petroleum
hydrocarbons).
Neanthes arenaceodentata
137
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TECHNICAL REPORT DATA
(Please read Instructions on the reverse before completing)
1. REPORT NO.
EPA-6QO73-80-0S3
3. RECIPIENT'S ACCESSION-NO.
4. TITLE AND SUBTITLE
The Effect of Different Pollutants on Ecologically
Important Polychaete Worms
5. REPORT DATE
JUNE 1980 ISSUING DATE.
6. PERFORMING ORGANIZATION CODE
7. AUTHOR(S)
Donald J. Reish
8. PERFORMING ORGANIZATION REPORT NO.
9. PERFORMING ORGANIZATION NAME AND ADDRESS
Department of Biology
California State University
Long Beach, CA 90840
10. PROGRAM ELEMENT NO.
1BA022
11. CONTRACT/GRANT NO.
R800962
12. SPONSORING AGENCY NAME AND ADDRESS
Environmental Research Laboratory -
Office of Research and Development
U.S. Environmental Protection Agency
Narragansett. Rhode Island 02882
Narra., RI
13. TYPE OF REPORT AND PERIOD COVERED
Final
14. SPONSORING AGENCY CODE
EPA/600/05
15. SUPPLEMENTARY NOTES
is. ABSTRACT
procedures for culturing marine polychaetous annelids from egg to egg
under laboratory conditions were described. A manual was prepared detailing the
procedures used in culturing 12 species of polychaetes. The polychaetes which have
been successfully cultured and the number of cycles completed in the laboratory are:
Neanthes arenaceodentata (50+), Capitella capitata (50+), Ctenodrilus serratus (50+),
Ophryotrocha diadema (50+), 0. puerilis (20+), Dinophilus sp. (50+), Dexiospira
brasiliensis (3), Polydora ligni (3), Boccardia proboscidea (3), Cirriformia
luxuriosa (1) , £. Spirabrancha (1) , and Halosydna j ohnsoni (1) .
The effects of heavy metals and the water soluble fractions of petroleum hydro-
carbons were measured over 96 hours, 28 days, and with some of the toxicants, over
a complete reproductive cycle for some pf these species of polychaetes. Mercury and
copper were the most toxic of the six jaetals tested and cadmium was the least toxic.
The 28-day LC^Q was less than the 96-hour value in most experiments. Larval stages
were more sensitive than the adults to heavy metals. Dexiospira was the most sensi-
tive species and Cirriformia luxuriosa was the most tolerant. Suppression of repro-
duction occurred with each species studied when exposed to heavy metals; the concen-
trations at which this occurred was less than the 28-day LC-,..
17.
KEY WORDS AND DOCUMENT ANALYSIS
DESCRIPTORS
b.IDENTIFIERS/OPEN ENDED TERMS
COS AT I Field/Group
Annelida
Toxicology
Toxicity
Pollution
Mercury
Cadmium
Copper
Lead
Chromium
Zinc
Hydrocarbons
Polychaeta
Polychaete culture
methods
Heavy metal toxicity
Petroleum hydrocarbon
toxicity
Interlaboratory calibrat
06, F
on
13. DISTRIBUTION STATEMENT
UNLIMITED
19. SECURITY CLASS (ThisReport}
UNCLASSIFIED
21. NO. OF PAGES
148
20. SECURITY CLASS (Thispage)
UNCLASSIFIED
22. PRICE
EPA Form 2220-1 (9-73)
138
•fr U.S. GOVERNMENT PRINTING OFFICE: 1980—657-165/0156
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